Sample Preparation for Trace Element Analysis

COMPREHENSIVE ANALYTICAL CHEMISTRY ELSEVIER B.V. Sara Burgerhartstraat 25 P.O. Box 211, 1000 AE Amsterdam, The Nether...

Author: Zoltan Mester | Ralph E. Sturgeon

234 downloads 1792 Views 10MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!

Report copyright / DMCA form

DOWNLOAD PDF

COMPREHENSIVE ANALYTICAL CHEMISTRY

ELSEVIER B.V. Sara Burgerhartstraat 25 P.O. Box 211, 1000 AE Amsterdam, The Netherlands q 2003 Elsevier B.V. All rights reserved. This work is protected under copyright by Elsevier Science, and the following terms and conditions apply to its use: Photocopying Single photocopies of single chapters may be made for personal use as allowed by national copyright laws. Permission of the Publisher and payment of a fee is required for all other photocopying, including multiple or systematic copying, copying for advertising or promotional purposes, resale, and all forms of document delivery. Special rates are available for educational institutions that wish to make photocopies for non-proﬁt educational classroom use. Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK; phone: (+44) 1865 843830, fax: (+44) 1865 853333, e-mail: [email protected]. You may also complete your request on-line via the Elsevier Science homepage (http://www.elsevier.com), by selecting ‘Customer Support’ and then ‘Obtaining Permissions’. In the USA, users may clear permissions and make payments through the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, USA; phone: (+1) (978) 7508400, fax: (+1) (978) 7504744, and in the UK through the Copyright Licensing Agency Rapid Clearance Service (CLARCS), 90 Tottenham Court Road, London W1P 0LP, UK; phone: (+44) 207 631 5555; fax: (+44) 207 631 5500. Other countries may have a local reprographic rights agency for payments. Derivative Works Tables of contents may be reproduced for internal circulation, but permission of Elsevier Science is required for external resale or distribution of such material. Permission of the Publisher is required for the other derivative works, including compilations and translations. Electronic Storage or Usage Permission of the Publisher is required to store or use electronically any material contained in this work, including any chapter or part of a chapter. Except as outlined above, no part of this work may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without prior written permission of the Publisher. Address permissions requests to: Elsevier Science Global Rights Department, at the mail, fax and e-mail addresses noted above. Notice No responsibility is assumed by the Publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent veriﬁcation of diagnoses and drug dosages should be made. First edition 2003 Library of Congress Cataloging in Publication Data / edited by Zolta´n Mester and Ralph Sturgeon p. cm. -- (Comprehensive analytical chemistry ; v. 41) Includes bibliographical references and index. ISBN 0-444-51101-6 (pbk. : alk. paper) -- ISBN 0-444-51101-6 (hardbound : alk. paper) 1. xxxx 2. xxxx 3. xxxx I. Mester, Zolta´n and Sturgeon, Ralph II. Series. QD75. W75 v. 41 [QD75.4.S24] 5430 .02--dc21

2002072248

British Library Cataloguing in Publication Data A catalogue record from the British Library has been applied for. ISBN: 0-444-51101-6 ISSN: 0166-526X 1 The paper used in this publication meets the requirements of ANSI/NISO Z39.48-1992 (Permanence of Paper). * Printed in The Netherlands.

COMPREHENSIVE ANALYTICAL CHEMISTRY ADVISORY BOARD

Professor A.M. Bond Monash University, Clayton, Victoria, Australia Dr T.W. Collette US Environmental Protection Agency, Athens, GA, U.S.A. Professor M. Grasserbauer Director of the Environment Institute, European Commission’ Joint Research Centre, Ispra, Italy Professor M.-C. Hennion Ecole Supe´rieure de Physique et de Chimie Industrielles, Paris, France Professor G. M. Hieftje Indiana University, Bloomington, IN, U.S.A. Professor G. Marko-Varga AstraZeneca, Lund, Sweden Professor D.L. Massart Vrije Universiteit, Brussels, Belgium Professor M.E. Meyerhoff University of Michigan, Ann Arbor, MI, U.S.A.

Wilson & Wilson’s COMPREHENSIVE ANALYTICAL CHEMISTRY

Edited by ´ D. BARCELO Research Professor Department of Environmental Chemistry IIQAB-CSIC Jordi Girona 18-26 08034 Barcelona Spain

Wilson & Wilson’s COMPREHENSIVE ANALYTICAL CHEMISTRY

VOLUME XLI

SAMPLE PREPARATION FOR TRACE ELEMENT ANALYSIS Edited by Z. MESTER R. STURGEON Institute for National Measurement Standards National Research Council 1500, Montreal Rd Ottawa, ON, KIA 0R6, Canada

2003 ELSEVIER AMSTERDAM – BOSTON – HEIDELBERG – LONDON – NEW YORK – OXFORD – PARIS – SAN DIEGO SAN FRANCISCO – SINGAPORE – SYDNEY – TOKYO

CONTRIBUTORS TO VOLUME XLI Freddy C. Adams Department of Chemistry, University of Antwerpen, Universiteitsplein 1, B-2610 Antwerpen, Belgium. [email protected] Roberto Alzaga Environmental Chemistry Department, IIQAB-CID-CSIC, Jordi Girona, 18, E-08034 Barcelona, Spain. Scott Anderson Air Liquide – Balazs Analytical Services, 46409 Landing Pky, Frement, CA 94538, USA. Kevin Ashley U.S. Department of Health and Human Services, Centers for Disease Control and Prevention, National Institute for Occupational Safety and Health, 4676 Columbia Parkway, Mailstop R-7, Cincinnati, OH 45226-1998, USA. [email protected] David P. Baldwin Ames Laboratory, Iowa State University, Ames, IA 50011,USA David Barclay CEM Corporation, 3100 Smith Farm Road, P.O. Box 200, Matthews, North Carolina 28106-0200, USA. [email protected] Ronald R. Barefoot Department of Geology, Earth Sciences Centre, 22 Russell Street, Toronto, Ontario, M5S 3B1, Canada. Douglas C. Baxter Division of Chemistry, Lulea˚ University of Technology, SE-971 87 Lulea˚, Sweden. [email protected] Josep M. Bayona Environmental Chemistry Department, IIQAB-CID-CSIC, Jordi Girona, 18, E-08034 Barcelona, Spain. [email protected] Maria Betti European Commission, JRC-ITU, P.O. Box 2340, 76125 Karlsruhe, Germany vi

Contributors to volume XLI

Robert I. Botto Analytical Services Laboratory, Baytown Chemical Plant Laboratory, 4500 Bayway Dr. Baytown, TX 77520, USA. [email protected] Brice Bouyssiere CNRS UMR 5034 Helioparc 2, av. Pr. Angot F-64053 PAU, France. Yong Cai Department of Chemistry and Southeast Environmental Research Center, Florida International University, Miami, Florida 33199, USA. cai@ﬁu.edu Carmen Camara Departamento de Quimica Analitica Facultad de Ciencias Quimicas, Universidad Complutense de Madrid 28040 Madrid, Spain. [email protected] Vale´rie Camel Institut National Agronomique Paris-Grignon, Laboratoire de Chimie Analytique, 16 rue Claude Bernard, 75231 Paris Cedex 05, France. [email protected] Joseph A. Caruso Department of Chemistry, University of Cincinnati, P.O. Box 210172, Cincinnati, OH 45221-0172, USA. [email protected] Maria Dolores Luque de Castro Department of Analytical Chemistry Annex C-3, Campus of Rabanales, University of Cordoba, E-14071 Cordoba (Spain). [email protected]. [email protected] Fernand Claisse 2780 Bd de Monaco, Quebec QC, Canada G1P3H2. [email protected] Ray Clement Ministry of the Environment, 125 Resources Road, Etobicoke, Ontario, Canada M9P 3V6. [email protected] Alberto de Diego Kimika Analitikoa Saila; Euskal Herriko Unibertsitatea; 644 P. K.; 48080, Bilbao, Spain. [email protected]

vii

Contributors to volume XLI

Sergi Dı´ez Environmental Chemistry Department, IIQAB-CID-CSIC, Jordi Girona, 18, E-08034 Barcelona, Spain. Olivier F.X. Donard CNRS, Laboratoire de Chimie Analytique Bio-inorganique et Environnement, He`lioparc, 2 avenue du President Angot, F-64000 Pau, France. [email protected] Peter Drouin Spectroscopy Section, Laboratory Services Branch, Ontario Ministry of the Environment, Ontario, Canada M9P 3V6. [email protected] Les Ebdon School of Environmental Sciences, University of Plymouth, Drake Circus, Plymouth PL4 8AA, UK. [email protected] John Ezzell Dionex Corporation, 1515 West 2200 South, Suite A, Salt Lake City, UT 84119-7209, USA [email protected] Zhao-Lun Fang Research Center for Analytical Sciences, Northeastern University, Chemistry Building, Box 332, Shenyang 110006, P.R. China. [email protected] Jo¨rg Feldmann Department of Chemistry, University of Aberdeen, Meston Walk, Old Aberdeen AB24 3UE, Scotland, UK. [email protected] Andrew S. Fisher School of Environmental Sciences, University of Phymouth, Drake Circus, Phymouth PL4 8AA, UK. a.ﬁ[email protected] Jose Luis Luque Garcı´a Department of Analytical Chemistry Annex C-3, Campus of Rabanales, University of Cordoba, E- 4071 Cordoba (Spain). [email protected] Walter Goessler Institute of Chemistry, Analytical Chemistry, Universita¨tsplatz 1, 8010 Graz, Austria. [email protected] viii

Contributors to volume XLI

Miguel de La Guardia Department of Analytical Chemistry, Faculty of Chemistry, University of Valencia, Dr Moliner St. 50. Burjassot, 46100-Valencia, Spain. [email protected] Monika Heisterkamp Mettler – Toledo GmbH, D-35396 Giessen, Germany. [email protected] Holger Hintelmann Department of Chemistry, Trent University, Peterborough, Ontario, K9J 7B8, Canada. [email protected] Jo´zsef Hlavay University of Veszpre´m, Department of Earth and Environmental Sciences, H-8200 Veszpre´m, Egyetem str. 10, P.O. Box 158 Hungary. [email protected] Michel Hoenig Centre for Veterinary and Agrochemical Research (CERVA), Leuvensesteenweg 17, B-3080 Tervuren, Belgium. [email protected] Milan Ihnat Paciﬁc Agri-Food Research Centre, Agriculture and Agri-Food Canada Summerland, BC, Canada, V0H 1Z0. [email protected] Hideyuki Itabashi Department of Applied Chemistry, Faculty of Engineering, Gunma University, Kiryu, Gunma 376-8515, Japan. [email protected] ˚ ke Jo¨nsson Jan A Analytical Chemistry, Lund University, P.O. Box 124, S-221 00 Lund, Sweden. [email protected] Katsu Kawabata PerkinElmer Instruments, 71 Four Valley Drive, Concord, Ontario, Canada, L4K 4V8. [email protected] Edward E. King CEM Corporation, 3100 Smith Farm Road, P.O. Box 200, Matthews, North Carolina 28106-0200, USA. [email protected] Yoko Kishi PerkinElmer Instruments, 71 Four Valley Drive, Concord, Ontario, Canada, L4K 4V8. ix

Contributors to volume XLI

Gunter Knapp Graz University of Technology, A-8010 Graz, Technikerstrasße 4, Graz, Austria. [email protected] Byron G. Kratochvil Department of Chemistry, University of Alberta, Edmonton, Alberta, Canada T6G 2G2. [email protected] Eva Krupp CNRS, Laboratoire de Chimie Analytique Bio-inorganique et Environnement, He`lioparc, 2 avenue du President Angot, F-64000 Pau, France. Doris Kuehnelt Institute of Chemistry, Analytical Chemistry, Universita¨tsplatz 1, 8010 Graz, Austria. [email protected] Claudia Ponce de Leon Department of Chemistry, University of Cincinnati, P.O. Box 210172, Cincinnati, OH 45221-0172, USA. Maurice Leroy European Commission, JRC-170, P.O. Box 2340, 76125 Karlsruhe, Germany Fuhe Li Air Liquide – Balazs Analytical Services, 46409 Landing Pky, Frement, CA 94538, USA. Ryszard Łobin´ski CNRS UMR 5034 Helioparc 2, av. Pr. Angot F-64053 PAU, France. [email protected] Yolanda Madrid Departamento de Quimica Analitica Facultad de Ciencias Quimicas, Universidad Complutense de Madrid 28040 Madrid, Spain. Lennart Mathiasson Analytical Chemistry, Lund University, P.O. Box 124, S-221 00 Lund, Sweden. [email protected] Henryk Matusiewicz Politechnika Poznan´ska, Department of Analytical Chemistry, 60-965, Poznan´, Poland. [email protected] Zolta´n Mester NRC/INMS, 1500, Montreal Rd, Ottawa, ON, K1A 0R6, Canada. [email protected] x

Contributors to volume XLI

Roberto Morabito ENEA, UTS PROT, SP Anguillarese 301, IT-00060 S. Maria di Galeria (Rome), Italy. [email protected] Angel Morales – Rubio Department of Analytical Chemistry, Faculty of Chemistry, University of Valencia, Dr Moliner St. 50 Burjassot, 46100 –Valencia, Spain. [email protected] Taketoshi Nakahara Department of Applied Chemistry, Graduate School of Engineering, Osaka Prefecture University, Sakai, Osaka 599-8531, Japan. [email protected] Marie-Pierre Pavageau CNRS, Laboratoire de Chimie Analytique Bio-inorganique et Environnement, He`lioparc, 2 avenue du President Angot, F-64000 Pau, France. Christophe Pe´cheyran CNRS, Laboratoire de Chimie Analytique Bio-inorganique et Environnement, He`lioparc, 2 avenue du President Angot, F-64000 Pau, France. Philip J. Potts Department of Earth Sciences, The Open University, Walton Hall, Milton Keynes MK7 6AA, UK. [email protected] Philippe Quevauviller European Commission, rue de la Loi 200, B-1049 Brussels, Belgium. [email protected] Gemma Rauret Departament de Quimica Analitica, Universitat de Barcelona, Martı´ i Franque`s 1-11, 3a Planta, 08028 Barcelona, Spain. [email protected] Philip Robinson School of Earth Sciences-Centre for Ore Deposit Research, University of Tasmania, Hobart, Tasmania 7001, Australia. [email protected] Ilia Rodushkin Analytica AB, Aurorum 10, SE-977 75 Lulea˚, Sweden. [email protected]

xi

Contributors to volume XLI

Richard E. Russo Lawrence Berkeley National Lab, 1 Cyclotron Road, Berkeley, CA 94720, USA. [email protected] Angels Sahuquillo Departament de Quimica Analitica, Universitat de Barcelona, Martı´ i Franque`s 1-11, 3a Planta, 08028 Barcelona, Spain. [email protected] Peter Schramel GSF-Forschungszentrum Institut fu¨r Oekologische Chemie AG, Spurenelementanalytik und Metallspeziation, Postfach 1129 (P.O. Box 1129), D-85758 Neuherberg. [email protected] Ralph Sturgeon NRC/INMS, 1500, Montreal Rd, Ottawa, ON, K1A 0R6, Canada. [email protected] Joanna Szpunar CNRS UMR 5034 He`lioparc 2, av. Pr. Angot F-64053 PAU, France. Anne P. Vonderheide Department of Chemistry, University of Cincinnati, P.O. Box 210172, Cincinnati, OH 45221-0172, USA. Scott Willie NRC/INMS 1500 Montreal Road, Ottawa, ON, K1A 0R6, Canada. [email protected]

xii

WILSON AND WILSON’S

COMPREHENSIVE ANALYTICAL CHEMISTRY VOLUMES IN THE SERIES Vol. IA

Vol. IB Vol. IC Vol. IIA

Vol. IIB

Vol. IIC

Vol. IID Vol. III

Vol. IV

Vol. V

Vol. VI Vol. VII Vol. VIII

Vol. IX

Analytical Processes Gas Analysis Inorganic Qualitative Analysis Organic Qualitative Analysis Inorganic Gravimetric Analysis Inorganic Titrimetric Analysis Organic Quantitative Analysis Analytical Chemistry of the Elements Electrochemical Analysis Electrodeposition Potentiometric Titrations Conductometric Titrations High-Frequency Titrations Liquid Chromatography in Columns Gas Chromatography Ion Exchangers Distillation Paper and Thin Layer Chromatography Radiochemical Methods Nuclear Magnetic Resonance and Electron Spin Resonance Methods X-Ray Spectrometry Coulometric Analysis Elemental Analysis with Minute Sample Standards and Standardization Separation by Liquid Amalgams Vacuum Fusion Analysis of Gases in Metals Electroanalysis in Molten Salts Instrumentation for Spectroscopy Atomic Absorption and Fluorescence Spectroscopy Diffuse Reﬂectance Spectroscopy Emission Spectroscopy Analytical Microwave Spectroscopy Analytical Applications of Electron Microscopy Analytical Infrared Spectroscopy Thermal Methods in Analytical Chemistry Substoichiometric Analytical Methods Enzyme Electrodes in Analytical Chemistry Molecular Fluorescence Spectroscopy Photometric Titrations Analytical Applications of Interferometry Ultraviolet Photoelectron and Photoion Spectroscopy Auger Electron Spectroscopy Plasma Excitation in Spectrochemical Analysis

xiii

Volumes in the series Vol. X Vol. XI

Vol. XII

Vol. XIII

Vol. XIV Vol. XV Vol. XVI Vol. XVII Vol. XVIII Vol. Vol. Vol. Vol. Vol.

XIX XX XXI XXII XXIII

Vol. Vol. Vol. Vol. Vol. Vol. Vol. Vol. Vol. Vol. Vol.

XXIV XXV XXVI XXVII XXVIII XXIX XXX XXXI XXXII XXXIII XXXIV

Vol. Vol. Vol. Vol.

XXXV XXXVI XXXVII XXXVIII

Vol. XXXIX Vol. XL

xiv

Organic Spot Tests Analysis The History of Analytical Chemistry The Application of Mathematical Statistics in Analytical Chemistry Mass Spectrometry Ion Selective Electrodes Thermal Analysis Part A. Simultaneous Thermoanalytical Examination by Means of the Derivatograph Part B. Biochemical and Clinical Applications of Thermometric and Thermal Analysis Part C. Emanation Thermal Analysis and other Radiometric Emanation Methods Part D. Thermophysical Properties of Solids Part E. Pulse Method of Measuring Thermophysical Parameters Analysis of Complex Hydrocarbons Part A. Separation Methods Part B. Group Analysis and Detailed Analysis Ion-Exchangers in Analytical Chemistry Methods of Organic Analysis Chemical Microscopy Thermomicroscopy of Organic Compounds Gas and Liquid Analysers Kinetic Methods in Chemical Analysis Application of Computers in Analytical Chemistry Analytical Visible and Ultraviolet Spectrometry Photometric Methods in Inorganic Trace Analysis New Developments in Conductometric and Oscillometric Analysis Titrimetric Analysis in Organic Solvents Analytical and Biomedical Applications of Ion-Selective Field-Effect Transistors Energy Dispersive X-Ray Fluorescence Analysis Preconcentration of Trace Elements Radionuclide X-Ray Fluorescence Analysis Voltammetry Analysis of Substances in the Gaseous Phase Chemiluminescence Immunoassay Spectrochemical Trace Analysis for Metals and Metalloids Surfactants in Analytical Chemistry Environmental Analytical Chemistry Elemental Speciation – New Approaches for Trace Element Analysis Discrete Sample Introduction Techniques for Inductively Coupled Plasma Mass Spectrometry Modern Fourier Transform Infrared Spectroscopy Chemical Test Methods of Analysis Sampling and Sample Preparation for Field and Laboratory Countercurrent Chromatography: The Support-Free Iiquid Stationary Phase Integrated Analytical Systems Analysis and Fate of Surfactants in the Aquatic Environment

Contents Contributors to Vol XLI Volumes in the Series . Series Editor’s Preface. Preface. . . . . . . . . Acronyms . . . . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

Chapter 1. Sampling and sample preservation for trace element analysis . . . . . . . . . . . . . . . . . . . . . . Byron Kratochvil 1.1 Introduction. . . . . . . . . . . . . . . . . . . . . 1.2 Preliminary considerations . . . . . . . . . . . . . 1.2.1 Sampling variability . . . . . . . . . . . . . . . . . . . 1.2.2 Sampling strategies . . . . . . . . . . . . . 1.2.3 Uncertainties in sampling . . . . . . . . . 1.3 Types of samples . . . . . . . . . . . . . . . . . . 1.3.1 Judgment samples . . . . . . . . . . . . . 1.3.2 Random samples . . . . . . . . . . . . . . 1.3.3 Systematic samples . . . . . . . . . . . . . 1.3.4 Subsamples . . . . . . . . . . . . . . . . . 1.3.5 Composite samples . . . . . . . . . . . . . 1.4 Planning the sampling operation . . . . . . . . . . 1.4.1 Deﬁning goals . . . . . . . . . . . . . . . . 1.4.2 Sampling plans . . . . . . . . . . . . . . . 1.5 Statistical sampling . . . . . . . . . . . . . . . . . 1.5.1 Introduction. . . . . . . . . . . . . . . . . 1.5.2 Minimum number of increments . . . . . . 1.5.3 Minimum size of increments in well-mixed particulate populations . . . . . . . . . . . 1.5.4 Sample increment size in segregated populations . . . . . . . . . . . . . . . . . 1.5.5 From where should increments be taken? . 1.5.6 Model-based sampling . . . . . . . . . . .

. . . . .

. . . . .

vi xiii xliii xliv xlvi

. .

1

. . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 2 2 2 3 3 3 4 4 5 5 5 5 6 8 8 8

. .

9

. . . . . .

10 11 12

xv

Contents

1.5.7

Balancing economic factors and purpose of data collection against sample quality . . . . . . . . 1.6 Sample handling and preservation during collection, transport, and storage . . . . . . . . . . . . . . . . . 1.6.1 Handling and storage of samples . . . . . . . . 1.6.2 Sampling equipment . . . . . . . . . . . . . . 1.6.3 Sample containers . . . . . . . . . . . . . . . 1.7 Quality assurance in sampling [24,25] . . . . . . . . . 1.7.1 Overall objectives . . . . . . . . . . . . . . . . 1.7.2 Quality control . . . . . . . . . . . . . . . . . 1.7.3 Quality assessment . . . . . . . . . . . . . . . 1.8 Glossary. . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . Selected bibliography . . . . . . . . . . . . . . . . . . . . . Chapter 2. Sources of analyte contamination and loss during the analytical process . . . . . . . . . . . . . . . . . . . . . . . Gunter Knapp and Peter Schramel 2.1 Introduction. . . . . . . . . . . . . . . . . . . . . . . 2.2 Contamination . . . . . . . . . . . . . . . . . . . . . 2.2.1 Materials . . . . . . . . . . . . . . . . . . . . 2.2.2 Reagents . . . . . . . . . . . . . . . . . . . . 2.2.3 Airborne particles . . . . . . . . . . . . . . . . 2.3 Losses . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Sampling . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Storage . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Drying and homogenisation. . . . . . . . . . . . . . . 2.7 Dilution, dissolution and digestion . . . . . . . . . . . 2.8 Separation and preconcentration . . . . . . . . . . . . 2.9 Element measurement . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . Chapter 3. Calibration approaches for trace element determination Douglas C. Baxter and Ilia Rodushkin 3.1 Introduction. . . . . . . . . . . . . . . . . . . . . . . 3.2 Basic assumptions and some terminology . . . . . . . 3.3 Selection of the calibration approach . . . . . . . . . . 3.4 Statistical evaluation of recovery data . . . . . . . . .

xvi

13 14 14 16 16 17 17 17 17 18 20 21

23 23 24 24 27 28 29 34 35 37 39 41 42 42 47 47 48 49 52

Contents

3.5

Linear 3.5.1 3.5.2 3.5.3

regression . . . . . . . . . . . . . . . . . . . . Ordinary linear regression . . . . . . . . . . . Weighted linear regression . . . . . . . . . . . Linear regression for data with uncertainties in both variables . . . . . . . . . . . . . . . . . . 3.6 External calibration. . . . . . . . . . . . . . . . . . . 3.6.1 Estimating uncertainty . . . . . . . . . . . . . 3.6.2 Optimizing precision . . . . . . . . . . . . . . 3.6.3 Accounting for non-constant sensitivity . . . . 3.7 Method of standard additions . . . . . . . . . . . . . . 3.7.1 Estimating uncertainty . . . . . . . . . . . . . 3.7.2 Optimizing precision . . . . . . . . . . . . . . 3.7.3 Accounting for non-constant sensitivity . . . . 3.8 Internal standardization . . . . . . . . . . . . . . . . 3.8.1 Estimating uncertainty . . . . . . . . . . . . . 3.8.2 Optimizing precision . . . . . . . . . . . . . . 3.9 Isotope dilution . . . . . . . . . . . . . . . . . . . . . 3.9.1 Mass discrimination and detector dead time . . 3.9.2 Estimating uncertainty . . . . . . . . . . . . . 3.9.3 Optimizing precision . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . .

55 56 59 64 67 68 70 72 74 75 76 78 79 80 81 81 85 86 88 90 90

Chapter 4. Stated references for ensuring traceability of trace element analysis . . . . . . . . . . . . . . . . . . . . . . . . 93 Philippe Quevauviller 4.1 Introduction. . . . . . . . . . . . . . . . . . . . . . . 93 4.2 Meaning of traceability for chemical measurements . . 94 4.3 SI units . . . . . . . . . . . . . . . . . . . . . . . . . 96 4.4 Documented standards . . . . . . . . . . . . . . . . . 96 4.5 Reference methods . . . . . . . . . . . . . . . . . . . 97 4.6 Reference materials . . . . . . . . . . . . . . . . . . . 99 4.6.1 The various categories of materials and related requirements . . . . . . . . . . . . . . . . . . 99 4.6.2 Production . . . . . . . . . . . . . . . . . . . 100 4.6.3 Methods used for material characterisation or certiﬁcation . . . . . . . . . . . . . . . . . . . 102

xvii

Contents

4.6.4 Use of reference materials . . . . . . . . . . 4.6.5 Traceability of reference materials . . . . . . 4.7 Specimen banking . . . . . . . . . . . . . . . . . . 4.8 Proﬁciency testing . . . . . . . . . . . . . . . . . . 4.9 Real-case achievement of traceability of trace element analysis . . . . . . . . . . . . . . . . . . . 4.9.1 Total trace element determinations . . . . . . 4.9.2 Operationally deﬁned trace element determinations . . . . . . . . . . . . . . . . 4.9.3 Determinations of chemical forms of elements 4.10 Conclusions . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . .

. . . .

. 106 . 106 . . . .

Chapter 5. Detection methods for the quantitation of trace elements . . . . . . . . . . . . . . . . . . . . . . . . . . . . Les Ebdon, Andrew S. Fisher, Maria Betti and Maurice Leroy 5.1 Introduction. . . . . . . . . . . . . . . . . . . . . . . 5.2 Classical methods . . . . . . . . . . . . . . . . . . . . 5.3 Flame spectrometry . . . . . . . . . . . . . . . . . . . 5.3.1 Introduction. . . . . . . . . . . . . . . . . . . 5.3.2 Theory . . . . . . . . . . . . . . . . . . . . . 5.3.3 Instrumentation . . . . . . . . . . . . . . . . 5.3.4 Interferences and background correction techniques. . . . . . . . . . . . . . . . . . . . 5.3.5 Conventional nebulisation . . . . . . . . . . . 5.3.6 Alternative methods of sample introduction . . 5.4 Electrothermal AAS. . . . . . . . . . . . . . . . . . . 5.4.1 Introduction. . . . . . . . . . . . . . . . . . . 5.4.2 Conventional ET-AAS . . . . . . . . . . . . . . 5.4.3 Multi-element ET-AAS . . . . . . . . . . . . . 5.4.4 Chemical vapour generation – ET-AAS . . . . . 5.4.5 Speciation . . . . . . . . . . . . . . . . . . . . 5.5 Inductively coupled plasma-atomic emission spectrometry . . . . . . . . . . . . . . . . . . . . . . 5.5.1 Introduction. . . . . . . . . . . . . . . . . . . 5.5.2 Theory and interferences . . . . . . . . . . . .

xviii

102 103 105 105

108 110 113 114

117 117 117 118 118 118 119 121 123 127 135 135 136 139 140 141 142 142 142

Contents

5.5.3 Instrumentation . . . . . . . . . . . . . . . . 5.5.4 Figures of merit . . . . . . . . . . . . . . . . . 5.6 Inductively coupled plasma-mass spectrometry . . . . 5.6.1 Introduction. . . . . . . . . . . . . . . . . . . 5.6.2 Theory . . . . . . . . . . . . . . . . . . . . . 5.6.3 Instrumentation . . . . . . . . . . . . . . . . 5.6.4 Different types of analysis . . . . . . . . . . . 5.6.5 Interferences . . . . . . . . . . . . . . . . . . 5.6.6 Sample introduction techniques . . . . . . . . 5.6.7 Figures of merit . . . . . . . . . . . . . . . . . 5.7 Atomic ﬂuorescence spectrometry. . . . . . . . . . . . 5.7.1 Introduction. . . . . . . . . . . . . . . . . . . 5.7.2 Theory . . . . . . . . . . . . . . . . . . . . . 5.7.3 Instrumentation . . . . . . . . . . . . . . . . 5.7.4 Sample introduction . . . . . . . . . . . . . . 5.7.5 Interferences . . . . . . . . . . . . . . . . . . 5.7.6 Figures of merit . . . . . . . . . . . . . . . . . 5.8 Other atomic absorption, emission and ﬂuorescence methods of detection . . . . . . . . . . . . . . . . . . 5.8.1 Microwave induced plasma . . . . . . . . . . . 5.8.2 Direct current plasma. . . . . . . . . . . . . . 5.9 Secondary ion mass spectrometry. . . . . . . . . . . . 5.9.1 Introduction. . . . . . . . . . . . . . . . . . . 5.9.2 Practical principles . . . . . . . . . . . . . . . 5.9.3 Sensitivity and quantiﬁcation. . . . . . . . . . 5.10 Glow discharge mass spectrometry . . . . . . . . . . . 5.10.1 Introduction. . . . . . . . . . . . . . . . . . . 5.10.2 Glow discharge processes . . . . . . . . . . . . 5.10.3 Applications to trace element analysis . . . . . 5.11 X-ray ﬂuorescence spectrometry . . . . . . . . . . . . 5.11.1 Introduction. . . . . . . . . . . . . . . . . . . 5.11.2 Instrumentation . . . . . . . . . . . . . . . . 5.11.3 Matrix effects . . . . . . . . . . . . . . . . . . 5.11.4 Quantitative and trace analysis . . . . . . . . 5.12 UV/Visible spectrophotometric and chemiluminescence techniques. . . . . . . . . . . . . . . . . . . . . . . . 5.12.1 UV/Visible spectrophotometric techniques . . .

143 152 152 152 153 154 156 156 158 160 160 160 162 162 163 163 163 164 164 165 165 165 167 168 171 171 173 175 176 176 177 177 178 179 179

xix

Contents

5.12.2 Molecular ﬂuorescence and chemiluminescence detection . . . . . . . . . . 5.13 Electrochemical methods . . . . . . . . . . . . . . . . 5.13.1 Differential pulse anodic stripping voltammetry 5.13.2 Cathodic and adsorptive stripping voltammetry 5.13.3 Ion selective electrodes . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . .

181 183 183 184 185 186

DIGESTION AND EXTRACTION APPROACHES

Chapter 6. Wet digestion methods . . . . . . . . . . . . . . . . . Henryk Matusiewicz 6.1 Introduction and brief history . . . . . . . . . . . . . 6.2 Nomenclature . . . . . . . . . . . . . . . . . . . . . . 6.3 Bibliography . . . . . . . . . . . . . . . . . . . . . . 6.4 Reagents and vessel materials for wet digestion procedures . . . . . . . . . . . . . . . . . . . . . . . 6.5 Wet acid digestion (decomposition and dissolution) procedures . . . . . . . . . . . . . . . . . . . . . . . 6.5.1 Open systems . . . . . . . . . . . . . . . . . . 6.5.2 Closed systems . . . . . . . . . . . . . . . . . 6.5.3 Flow systems . . . . . . . . . . . . . . . . . . 6.5.4 Vapor-phase acid digestion (gas-phase reactions) 6.5.5 Efﬁciency of wet digestion (decomposition and dissolution) procedures . . . . . . . . . . . . . 6.5.6 Comparison of wet digestion techniques . . . . 6.5.7 Digestion systems (instrumentation, equipment, automation) . . . . . . . . . . . . 6.5.8 Safety of acid digestions (sample acid digestion safety). . . . . . . . . . . . . . . . . 6.6 Conclusions and future trends . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . Chapter 7. Dry ashing . . . . . . . . . . . . . . . Michel Hoenig 7.1 General considerations . . . . . . . . . 7.2 Why dry ashing? . . . . . . . . . . . . 7.3 Oxidation process and dissolution of the

xx

193 193 194 194 195 199 199 203 210 213 216 219 220 221 224 228

. . . . . . . . 235 . . . . . . . . 235 . . . . . . . . 238 residue . . . . 240

Contents

7.3.1 Particular case of plant matrices . . . . . . . Methodology . . . . . . . . . . . . . . . . . . . . . 7.4.1 Heating devices . . . . . . . . . . . . . . . . 7.4.2 Ashing vessels . . . . . . . . . . . . . . . . 7.4.3 Inﬂuence of the sample composition . . . . . 7.4.4 Operating modes for environmental samples . 7.5 Particular cases of arsenic and selenium . . . . . . . 7.5.1 Ashing aids . . . . . . . . . . . . . . . . . . 7.5.2 What to do? . . . . . . . . . . . . . . . . . . 7.6 Conclusions . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4

Chapter 8. Microwave based extraction . . . . . . . . . Edward E. King and David Barclay 8.1 Introduction. . . . . . . . . . . . . . . . . . 8.2 Brief history of industrial microwave devices. 8.3 Microwave theory . . . . . . . . . . . . . . . 8.4 Microwave laboratory equipment . . . . . . . 8.4.1 Magnetron . . . . . . . . . . . . . . 8.4.2 Power application . . . . . . . . . . . 8.4.3 Waveguide . . . . . . . . . . . . . . 8.4.4 Microwave cavity . . . . . . . . . . . 8.4.5 Reﬂected energy . . . . . . . . . . . 8.4.6 Mode stirrer and turntables . . . . . 8.4.7 Microwave compatible materials . . . 8.5 Vessels . . . . . . . . . . . . . . . . . . . . 8.5.1 Materials . . . . . . . . . . . . . . . 8.5.2 Structural components . . . . . . . . 8.5.3 Safety . . . . . . . . . . . . . . . . . 8.5.4 Closed vessels . . . . . . . . . . . . . 8.5.5 Vent and reseal vessels . . . . . . . . 8.5.6 Open vessels . . . . . . . . . . . . . 8.6 Control systems . . . . . . . . . . . . . . . . 8.6.1 Power/time . . . . . . . . . . . . . . 8.6.2 Pressure. . . . . . . . . . . . . . . . 8.6.3 Temperature . . . . . . . . . . . . .

. . . . . . . . . . .

243 244 244 245 246 246 248 250 251 253 254

. . . . . 257 . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . .

257 257 258 263 264 267 267 267 267 268 269 270 271 272 272 273 274 274 275 275 275 278

xxi

Contents

8.6.4 Power optimization feedback . . . . . Methodology . . . . . . . . . . . . . . . . . 8.7.1 Pressurized closed vessel extractions . 8.7.2 Atmospheric open vessel extractions . 8.8 Sample types . . . . . . . . . . . . . . . . . 8.8.1 Inorganic . . . . . . . . . . . . . . . 8.8.2 Leaches and other partial extractions 8.8.3 Complete dissolutions . . . . . . . . . 8.8.4 High-temperature extractions . . . . 8.8.5 Complex sequential extractions . . . . 8.8.6 Organic . . . . . . . . . . . . . . . . 8.8.7 Carbohydrates . . . . . . . . . . . . 8.8.8 Proteins . . . . . . . . . . . . . . . . 8.8.9 Fats, oils, and waxes . . . . . . . . . 8.9 Advanced applications . . . . . . . . . . . . 8.9.1 Clean chemistry. . . . . . . . . . . . 8.9.2 Concentration/evaporation . . . . . . 8.10 Conclusions . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . 8.7

Chapter 9. Fusion and ﬂuxes . . . . . . . . . . . . . Fernand Claisse 9.1 Introduction. . . . . . . . . . . . . . . . . 9.2 Fusion in lithium borates . . . . . . . . . . 9.2.1 General . . . . . . . . . . . . . . . 9.3 The key to successful fusion beads . . . . . 9.3.1 The concept of “neutrality” . . . . . 9.3.2 The optimal ﬂux and crystallization 9.3.3 Cracking of fused beads . . . . . . . 9.3.4 Loss and retention of sulfur. . . . . 9.4 Application to trace element analysis . . . . 9.4.1 Maximizing X-ray intensities . . . . 9.4.2 Minimizing background . . . . . . . 9.5 Features of fusion for trace elements . . . . References . . . . . . . . . . . . . . . . . . . . .

xxii

. . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . .

280 281 282 283 285 285 286 287 288 290 291 292 293 294 297 297 297 298 299 299

. . . . . . 301 . . . . . . . . . . . . .

. . . . . . . . . . . . .

. . . . . . . . . . . . .

. . . . . . . . . . . . .

. . . . . . . . . . . . .

. . . . . . . . . . . . .

301 301 301 305 305 306 307 308 308 308 309 309 310

Contents

Chapter 10. Supercritical ﬂuid extraction . . . . . . . . . Roberto Alzaga, Sergi Dı´ez and Josep M. Bayona 10.1 Properties of supercritical ﬂuids . . . . . . . . 10.2 Instrumentation . . . . . . . . . . . . . . . . 10.2.1 Experimental solubility measurements . 10.3 SFE of trace elements. . . . . . . . . . . . . . 10.3.1 Ligand solubility in SFs. . . . . . . . . 10.3.2 Complex –SF solubility . . . . . . . . . 10.3.3 SFE process . . . . . . . . . . . . . . . 10.4 Organometallic compounds . . . . . . . . . . . 10.4.1 Organotin compounds. . . . . . . . . . 10.4.2 Organomercury compounds . . . . . . . 10.4.3 Organolead compounds . . . . . . . . . 10.4.4 Arsenic compounds . . . . . . . . . . . 10.5 Conclusion . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . .

. . . . 313 . . . . . . . . . . . . . .

Chapter 11. Accelerated solvent extraction of organometallic and inorganic compounds . . . . . . . . . . . . . . . . John L. Ezzell 11.1 Accelerated solvent extraction as a sample preparation technique . . . . . . . . . . . . . . 11.1.1 Introduction. . . . . . . . . . . . . . . . 11.1.2 Basic principles of ASE operation. . . . . 11.1.3 ASE instrumentation . . . . . . . . . . . 11.1.4 ASE methods development . . . . . . . . 11.1.5 Application areas . . . . . . . . . . . . . 11.1.6 Summary . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . Chapter 12. Sonication as a sample preparation method elemental analysis . . . . . . . . . . . . . . . . . . Kevin Ashley 12.1 Introduction. . . . . . . . . . . . . . . . . . 12.2 Methodological considerations . . . . . . . . 12.3 Historical background . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

. . . . . . . . . . . . . .

313 316 318 318 326 328 328 332 332 338 339 339 339 340

. . . 343

. . . . . . . .

. . . . . . . .

. . . . . . . .

343 343 344 345 346 349 351 352

for . . . . . 353 . . . . . 353 . . . . . 354 . . . . . 357

xxiii

Contents

12.4 Applications—sonication and sample preparation 12.4.1 Environmental analysis. . . . . . . . . . 12.4.2 Industrial hygiene . . . . . . . . . . . . 12.4.3 Biological tissues and ﬂuids. . . . . . . . 12.4.4 Other applications . . . . . . . . . . . . 12.5 Summary . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . .

. . . . . . .

. . . . . . .

Chapter 13. Solid phase microextraction as a tool for trace element determination . . . . . . . . . . . . . . . . . . . . Zolta´n Mester and Ralph Sturgeon 13.1 Introduction. . . . . . . . . . . . . . . . . . . . . . 13.2 General description of solid phase microextraction . . 13.2.1 Extraction modes . . . . . . . . . . . . . . . 13.2.2 Coatings. . . . . . . . . . . . . . . . . . . . 13.3 Solid phase microextraction: step-by-step method development. . . . . . . . . . . . . . . . . . . . . . 13.3.1 Extraction mode selection. . . . . . . . . . . 13.3.2 Fiber coating selection . . . . . . . . . . . . 13.3.3 Derivatization method selection . . . . . . . 13.3.4 Optimization of desorption conditions . . . . 13.3.5 Sample volume optimization . . . . . . . . . 13.3.6 Optimization of the extraction time. . . . . . 13.3.7 Optimization of extraction conditions. . . . . 13.3.8 Determination of the linear dynamic range. . 13.3.9 Selection of the calibration method . . . . . . 13.3.10 Precision of the method . . . . . . . . . . . . 13.3.11 Automation of the method . . . . . . . . . . 13.4 Solid phase microextraction for speciation analysis . 13.4.1 Volatile metal species—gas chromatographic determination . . . . . . . . . . . . . . . . . 13.5 Solid phase microextraction as an investigative tool . 13.6 Limitations of solid phase microextraction . . . . . . 13.7 Isotope dilution calibration in combination with solid phase microextraction. . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . .

xxiv

. . . . . . .

358 358 360 363 364 366 366

. 371 . . . .

371 373 373 375

. . . . . . . . . . . . .

375 375 376 376 376 376 377 377 378 378 379 379 380

. 380 . 388 . 388 . 389 . 390

Contents

Chapter 14. Solid-phase extraction . . . . . . . . . . . . . Vale´rie Camel 14.1 Introduction. . . . . . . . . . . . . . . . . . . . 14.2 Theory . . . . . . . . . . . . . . . . . . . . . . 14.2.1 Presentation of the technique . . . . . . . 14.2.2 Operation . . . . . . . . . . . . . . . . . 14.2.3 Advantages of the technique . . . . . . . 14.3 Step-by-step method development guide . . . . . 14.3.1 Selection of solid sorbent . . . . . . . . . 14.3.2 Inﬂuential parameters . . . . . . . . . . 14.4 Applications of SPE to the determination of some trace elements . . . . . . . . . . . . . . . . . . 14.4.1 Chromium. . . . . . . . . . . . . . . . . 14.4.2 Iron . . . . . . . . . . . . . . . . . . . . 14.4.3 Mercury . . . . . . . . . . . . . . . . . . 14.4.4 Selenium . . . . . . . . . . . . . . . . . 14.4.5 Tin . . . . . . . . . . . . . . . . . . . . 14.5 Conclusion . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 15. Chelation solvent extraction for separation of metal ions . . . . . . . . . . . . . . . . . . . . . . . . Hideyuki Itabashi and Taketoshi Nakahara 15.1 Introduction. . . . . . . . . . . . . . . . . . . . 15.2 Theoretical considerations . . . . . . . . . . . . 15.2.1 General principles . . . . . . . . . . . . 15.2.2 Preconcentration of metal ions . . . . . . 15.2.3 Mutual separation of metal ions . . . . . 15.2.4 Speciation of metal ions in natural water. 15.3 Adsorption of metal ions using chelating resins . 15.3.1 General principles . . . . . . . . . . . . 15.3.2 Features of some chelating resins . . . . 15.4 Application of chelation to sample preparation for trace metal analysis. . . . . . . . . . . . . . . . 15.4.1 Procedure for the extraction of metal ions from natural waters. . . . . . . . . . . .

. . . 393 . . . . . . . .

. . . . . . . .

. . . . . . . .

393 393 394 400 403 410 410 432

. . . . . . . .

. . . . . . . .

. . . . . . . .

439 439 443 445 445 445 450 451

. . . 459 . . . . . . . . .

. . . . . . . . .

. . . . . . . . .

459 460 460 465 467 472 474 475 475

. . . 477 . . . 477

xxv

Contents

15.4.2 Procedure for the extraction of metal ions from high-purity materials and inorganic solid samples . . . . . . . . . . . . . . . . . . 15.4.3 Procedure for the extraction of metal ions from biological samples . . . . . . . . . . . . . 15.4.4 Procedure for the speciation of metal ions in natural waters . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . .

488 490 491 492

Chapter 16. Cryogenic trapping for speciation analysis . . . . . Marie-Pierre Pavageau, Eva Krupp, Alberto de Diego, Christophe Pe´cheyran and Olivier F.X. Donard 16.1 Introduction. . . . . . . . . . . . . . . . . . . . . . 16.2 Deﬁnition of volatile species . . . . . . . . . . . . . 16.3 Physico-chemical principles and processes associated with cryofocusing . . . . . . . . . . . . . . . . . . . 16.4 Analytical constraints. . . . . . . . . . . . . . . . . 16.4.1 Removal of CO2 . . . . . . . . . . . . . . . . 16.4.2 Water removal . . . . . . . . . . . . . . . . 16.5 Sample preservation and stability . . . . . . . . . . 16.6 Instrumentation for cryogenic trapping and selected applications. . . . . . . . . . . . . . . . . . . . . . 16.6.1 Cryosampler for determination of industrial and environmental VMCs . . . . . . . . . . 16.6.2 Cryogenic trapping for speciation analysis . . References . . . . . . . . . . . . . . . . . . . . . . . . . .

. 520 . 523 . 529

Chapter 17. Biotrapping as an alternative to metal preconcentration and speciation . . . . . . . . . . . . Yolanda Madrid and Carmen Ca´mara 17.1 Introduction. . . . . . . . . . . . . . . . . . . 17.2 General characteristics of biological substrates. 17.2.1 Algae . . . . . . . . . . . . . . . . . . 17.2.2 Bacteria . . . . . . . . . . . . . . . . . 17.2.3 Fungi . . . . . . . . . . . . . . . . . .

. . . . .

xxvi

. 495

. 495 . 502 . . . . .

504 509 510 512 517

. 520

. . . . 533 . . . . .

. . . . .

. . . . .

533 535 535 536 537

Contents

17.3 Uptake mechanisms . . . . . . . 17.4 Working procedures . . . . . . . . 17.4.1 Immobilisation . . . . . . 17.5 Applications . . . . . . . . . . . . 17.5.1 Analytical applications . . 17.5.2 Technological applications 17.6 Conclusions . . . . . . . . . . . . References . . . . . . . . . . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

Chapter18 Membrane extraction . . . . . . . . . . . . . . . . . . . . ˚ ke Jo¨nsson and Lennart Mathiasson Jan A 18.1 Introduction. . . . . . . . . . . . . . . . . . . . . . 18.2 Membrane extraction techniques . . . . . . . . . . . 18.2.1 Supported liquid membrane extraction (SLM) 18.2.2 Microporous membrane liquid liquid extraction (MMLLE) . . . . . . . . . . . . . 18.3 Chemical principles for metal extraction . . . . . . . 18.4 Properties of membrane extraction . . . . . . . . . . 18.4.1 Clean-up and selectivity . . . . . . . . . . . 18.4.2 Enrichment . . . . . . . . . . . . . . . . . . 18.4.3 Automation and unattended operation . . . . 18.4.4 Solvent consumption . . . . . . . . . . . . . 18.5 Experimental set-up . . . . . . . . . . . . . . . . . 18.5.1 Flow systems for membrane extraction . . . . 18.5.2 How to set up a membrane extraction experiment for metal ions. . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . Chapter 19. Derivatization and vapor generation trace element analysis and speciation . . . . Yong Cai 19.1 Introduction. . . . . . . . . . . . . . 19.2 Theory . . . . . . . . . . . . . . . . 19.2.1 Grignard reactions . . . . . . 19.2.2 Hydride generation . . . . . .

. . . . . . . .

538 541 541 546 546 556 557 557

. 559 . 559 . 559 . 560 . . . . . . . . .

565 566 566 567 568 569 570 570 570

. 571 . 574 . 574

methods for . . . . . . . . . 577 . . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

577 578 578 579

xxvii

Contents

19.2.3 Aqueous derivatization (aryl)borates . . . . . 19.3 Method development . . . . . 19.3.1 Grignard reactions . . 19.3.2 Aqueous derivatization 19.4 Applications . . . . . . . . . . Acknowledgements . . . . . . . . . References . . . . . . . . . . . . . .

with tetraalkyl . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 20. Laser ablation sampling . . . . Richard E. Russo and David P. Baldwin 20.1 Introduction. . . . . . . . . . . . 20.2 Experimental system . . . . . . . 20.3 Ablation detection systems . . . . 20.4 Calibration . . . . . . . . . . . . 20.5 Fractionation . . . . . . . . . . . 20.6 Conclusion . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . References . . . . . . . . . . . . . . . .

. . . . . . .

. . . . . . .

. . . . . . .

. . . . . . .

580 583 583 584 585 590 590

. . . . . . . . . . . 593 . . . . . . . .

593 594 599 601 603 604 606 606

Chapter 21. Flow injection techniques for sample pretreatment . . Zhao-Lun Fang 21.1 Introduction. . . . . . . . . . . . . . . . . . . . . . . 21.1.1 General . . . . . . . . . . . . . . . . . . . . . 21.1.2 General features of ﬂow injection on-line sample pretreatment systems. . . . . . . . . . 21.1.3 Classiﬁcation of FI sample pretreatment systems . . . . . . . . . . . . . . . . . . . . . 21.1.4 Principles and general guidelines for the development of FI systems . . . . . . . . . . . 21.1.5 Practical hints for manipulation of FI equipment 21.2 FI liquid – liquid extraction systems. . . . . . . . . . . 21.2.1 Introduction. . . . . . . . . . . . . . . . . . . 21.2.2 Apparatus for FI liquid – liquid extraction . . . 21.2.3 Guidelines for the development of FI liquid – liquid extraction systems . . . . . . . . 21.2.4 Typical manifolds for FI liquid –liquid extraction 21.3 FI solid phase extraction systems. . . . . . . . . . . .

611

xxviii

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

611 611 612 612 613 614 615 615 617 623 624 626

Contents

21.3.1 Introduction. . . . . . . . . . . . . . . . . . . 626 21.3.2 Sorption media for FI solid phase extraction . . 627 21.3.3 Guidelines for the development of FI solid phase extraction systems . . . . . . . . . . . . 629 21.3.4 Typical manifolds for FI solid phase extraction . 630 21.4 FI vapor generation systems . . . . . . . . . . . . . . 635 21.4.1 Introduction. . . . . . . . . . . . . . . . . . . 635 21.4.2 Gas – liquid separators for FI vapor generation . 635 21.4.3 Guidelines for development of FI vapor generation systems . . . . . . . . . . . . . . . 636 21.4.4 Typical FI manifolds for VG-AAS . . . . . . . . 638 21.5 FI gas diffusion systems . . . . . . . . . . . . . . . . 641 21.5.1 General . . . . . . . . . . . . . . . . . . . . . 641 21.5.2 Gas-diffusion separators . . . . . . . . . . . . 641 21.5.3 Typical FI manifolds for gas-diffusion separation and preconcentration . . . . . . . . . . . . . . 642 21.6 FI on-line sample digestion . . . . . . . . . . . . . . . 643 21.6.1 Introduction. . . . . . . . . . . . . . . . . . . 643 21.6.2 FI on-line sample digestion systems for AAS . . 644 21.6.3 FI digestion systems coupled to VG-AAS . . . . 644 21.6.4 FI systems for digestion of solid samples in AAS 645 21.6.5 FI pretreatment systems with on-line photo-oxidation by UV irradiation . . . . . . . 646 References . . . . . . . . . . . . . . . . . . . . . . . . . . . 646

Chapter 22. Automation of sample preparation . . . . . . . . . . Maria Dolores Luque de Castro and Jose Luis Luque Garcı´a 22.1 Introduction. . . . . . . . . . . . . . . . . . . . . . . 22.1.1 Generalities . . . . . . . . . . . . . . . . . . . 22.1.2 Principal shortcomings in automating sample preparation . . . . . . . . . . . . . . . . . . . 22.1.3 Batch versus serial approaches to automated sample preparation . . . . . . . . . . . . . . . 22.1.4 Bar codes: a necessary tool in automating routine analyses . . . . . . . . . . . . . . . . 22.2 Automation of liquid sample preparation . . . . . . . .

649 649 649 650 651 652 653

xxix

Contents

22.2.1 Continuous systems. . . . . . . . . . . . . . . 22.2.2 Discontinuous approaches . . . . . . . . . . . 22.3 Automation of solid sample preparation . . . . . . . . 22.3.1 One-step approaches to automation and acceleration of solid sample preparation . . . . 22.3.2 Direct solid sampling . . . . . . . . . . . . . . 22.4 Robotics . . . . . . . . . . . . . . . . . . . . . . . . . 22.4.1 Workstations, robots, modules and peripherals . 22.4.2 The role of robots in the analytical process . . . 22.4.3 Analytical scope of robotics for sample preparation . . . . . . . . . . . . . . . . . . . 22.5 Advantages and disadvantages of automation of sample preparation . . . . . . . . . . . . . . . . . . 22.6 Future prospects . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . .

653 656 659 659 660 660 662 670 674 676 677 678

MATRICES

Chapter 23. Sample preparation for crude oil, petroleum products and polymers . . . . . . . . . . . . . . . . . . . . . . . . . Robert I. Botto 23.1 Introduction. . . . . . . . . . . . . . . . . . . . . . . 23.1.1 Nature of petroleum crude, products and polymers . . . . . . . . . . . . . . . . . . . . 23.1.2 Element context and species in petroleum crude, products and polymers. . . . . . . . . . 23.1.3 Sample preparation challenges for trace element analysis . . . . . . . . . . . . . . . . 23.2 Sample preparation techniques and instrumentation . 23.2.1 Ashing techniques . . . . . . . . . . . . . . . 23.2.2 Acid mineralization techniques . . . . . . . . . 23.2.3 Oxygen combustion . . . . . . . . . . . . . . . 23.2.4 Sample component separations/extractive sample preparation . . . . . . . . . . . . . . . 23.2.5 Organic sample dilutions and dissolutions . . . 23.2.6 Stable emulsions . . . . . . . . . . . . . . . . 23.2.7 Scrubber sampling for C1 – C4 hydrocarbons and gases . . . . . . . . . . . . . . . . . . . .

xxx

683 683 683 686 691 693 693 697 704 706 707 709 711

Contents

23.3 Cleanliness and quality assurance . . . 23.3.1 Equipment cleaning. . . . . . . 23.3.2 Clean techniques and disposable 23.3.3 Quality assurance. . . . . . . . Acknowledgements . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . . equipment . . . . . . . . . . . . . . . . . .

. . . . . .

. . . . . .

713 713 714 715 718 719

Chapter 24. Sample preparation of geological samples, soils and sediments . . . . . . . . . . . . . . . . . . . . . . . . . 723 Philip J. Potts and Philip Robinson 24.1 Introduction. . . . . . . . . . . . . . . . . . . . . . . 723 24.2 Sample preparation . . . . . . . . . . . . . . . . . . . 723 24.2.1 Sample collection . . . . . . . . . . . . . . . . 724 24.2.2 Crushing and grinding . . . . . . . . . . . . . 726 24.2.3 Selecting an appropriate test portion . . . . . . 727 24.3 Choice of approach . . . . . . . . . . . . . . . . . . . 728 24.3.1 Fitness-for-purpose . . . . . . . . . . . . . . . 728 24.3.2 Choice of sample preparation procedure based on choice of technique. . . . . . . . . . . . . . 728 24.3.3 Choice of sample preparation based on the characteristics of elements . . . . . . . . . . . 729 24.4 Methods that do not require any sample digestion— in situ methods of analysis . . . . . . . . . . . . . . . 730 24.4.1 Portable X-ray ﬂuorescence . . . . . . . . . . . 730 24.4.2 Gamma spectrometry . . . . . . . . . . . . . . 732 24.4.3 Laser ablation techniques and other microprobe/ microanalytical techniques . . . . . . . . . . . 732 24.5 Methods based on solid samples . . . . . . . . . . . . 733 24.5.1 Direct determinations on powders . . . . . . . 733 24.5.2 Powder pellet for XRF . . . . . . . . . . . . . 734 24.5.3 Glass disks for XRF trace determinations . . . 734 24.6 Dissolution methods based on acid attack . . . . . . . 735 24.6.1 Properties of acids used in the decomposition of geological materials . . . . . . . . . . . . . . . 736 24.6.2 Open vessel and low-pressure acid digestion . . 738

xxxi

Contents

24.6.3 HF – HNO3 decomposition method in Savillexw screw top vials . . . . . . . . . . . . . . . . . 740 24.6.4 Closed vessel high pressure acid digestion . . . 740 24.6.5 HF/H2SO4 decomposition method in closed, high pressure vessels . . . . . . . . . . . . . . 742 24.6.6 HF/HClO4 – HCl decomposition method in closed, high pressure vessels . . . . . . . . . . . . . . 742 24.6.7 Microwave oven digestion. . . . . . . . . . . . 743 24.6.8 Partial acid attack . . . . . . . . . . . . . . . 744 24.6.9 Difﬁcult minerals . . . . . . . . . . . . . . . . 745 24.7 Decomposition by molten salt fusion . . . . . . . . . . 750 24.7.1 Total fusion . . . . . . . . . . . . . . . . . . . 750 24.7.2 A LiBO2 fusion procedure . . . . . . . . . . . 752 24.7.3 Sintering . . . . . . . . . . . . . . . . . . . . 752 24.7.4 Fire assay . . . . . . . . . . . . . . . . . . . . 752 24.8 Pre-concentration and separation procedures . . . . . 753 24.8.1 Ion exchange . . . . . . . . . . . . . . . . . . 753 24.8.2 Solvent extraction and co-precipitation . . . . . 754 24.8.3 Vapour generation . . . . . . . . . . . . . . . 755 24.9 Sequential extractions and dissolutions . . . . . . . . 755 24.9.1 Procedure of Tessier et al. . . . . . . . . . . . 756 24.9.2 The “BCR” method . . . . . . . . . . . . . . . 757 24.9.3 Selective extractions for geochemical exploration . . . . . . . . . . . . . . . . . . . 757 24.10 Summary and conclusions . . . . . . . . . . . . . . . 758 References . . . . . . . . . . . . . . . . . . . . . . . . . . . 758 Chapter 25. Sample preparation for food analysis. . . . . . . . Milan Ihnat 25.1 Introduction. . . . . . . . . . . . . . . . . . . . . . 25.2 Literature . . . . . . . . . . . . . . . . . . . . . . . 25.2.1 Books on sample treatment, decomposition . . 25.2.2 Books including chapters, sections on or discussing sample treatment, decomposition . 25.2.3 Reviews on sample treatment, decomposition 25.2.4 Other reviews including coverage of sample treatment, decomposition . . . . . . . . . .

xxxii

. 765 . 765 . 766 . 767 . 767 . 767 . 768

Contents

25.2.5 Papers, publications, containing (major) writing on sample treatment . . . . . . . . . . . . . . 25.3 Pretreatment . . . . . . . . . . . . . . . . . . . . . . 25.4 Classiﬁcation of sample treatment methods . . . . . . 25.5 Compilation of sample treatment methods for foods . . 25.5.1 No treatment . . . . . . . . . . . . . . . . . . 25.5.2 Dry ashing . . . . . . . . . . . . . . . . . . . 25.5.3 Wet digestion—conventional . . . . . . . . . . 25.5.4 Wet digestion—microwave-assisted . . . . . . . 25.5.5 Slurry sample preparation . . . . . . . . . . . 25.6 Speciﬁc cases: methods, elements, matrices . . . . . . 25.6.1 Analytical method. . . . . . . . . . . . . . . . 25.6.2 Elements . . . . . . . . . . . . . . . . . . . . 25.6.3 Matrix and constituents . . . . . . . . . . . . 25.7 Examples of speciﬁc, recommended sample treatment procedures . . . . . . . . . . . . . . . . . . . . . . . 25.7.1 Conventional wet digestion with nitric and perchloric acids . . . . . . . . . . . . . . . . . 25.7.2 Dry ashing with or without ashing aid . . . . . 25.7.3 Microwave-assisted wet digestion. . . . . . . . 25.8 Closing remarks. . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 26. The determination of trace elements in water . . Scott N. Willie 26.1 Direct methods of determination . . . . . . . . . . 26.2 Preconcentration techniques—multielement . . . . 26.3 Preconcentration—individual elements . . . . . . . 26.4 Determination of trace elements as volatile species 26.5 Mercury . . . . . . . . . . . . . . . . . . . . . . . 26.6 Luminescence . . . . . . . . . . . . . . . . . . . . 26.7 Voltammetry . . . . . . . . . . . . . . . . . . . . 26.8 Total-reﬂection X-Ray ﬂuorescence spectrometry . . 26.9 Conclusions . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . .

768 768 770 770 833 834 836 838 838 839 839 841 842 843 843 845 845 846 847

. . 857 . . . . . . . . . .

. . . . . . . . . .

857 860 865 865 879 880 885 891 894 894

xxxiii

Contents

Chapter 27. Aerosol sampling and sample preparation for elemental analysis . . . . . . . . . . . . . . . . . . . . . Jo´zsef Hlavay 27.1 Introduction. . . . . . . . . . . . . . . . . . . . . 27.1.1 Objectives of monitoring . . . . . . . . . . 27.2 Sampling of aerosols . . . . . . . . . . . . . . . . 27.2.1 General considerations . . . . . . . . . . . 27.2.2 Sampling of aerosol by impactors . . . . . . 27.2.3 Ambient sampling for the respirable fraction . . . . . . . . . . . . . . . . . . . 27.2.4 High-volume aerosol samplers . . . . . . . 27.2.5 Speciation aerosol sampling system . . . . 27.2.6 Passive samplers . . . . . . . . . . . . . . 27.3 Sequential extraction schemes for aerosol samples . 27.4 Discussion. . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . General terms used in sampling. . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . Chapter 28. Sample preparation for industrial waste analysis Peter Drouin and Ray E. Clement 28.1 Types of industrial waste . . . . . . . . . . . . . . 28.2 Safety considerations for industrial waste analysis. 28.3 Sample characteristics and industrial waste sampling . . . . . . . . . . . . . . . . . . . . . . 28.4 Digestions . . . . . . . . . . . . . . . . . . . . . . 28.4.1 Aqueous sample types—US EPA methods . 28.4.2 Solid sample types – US EPA methods . . . 28.4.3 Reported studies—hot plate digestion . . . 28.4.4 Reported studies—microwave digestion methods . . . . . . . . . . . . . . . . . . . 28.4.5 Ultrasound-assisted extractions . . . . . . 28.4.6 Alkaline digestions . . . . . . . . . . . . . 28.4.7 Laboratory safety . . . . . . . . . . . . . . 28.5 Leach procedures . . . . . . . . . . . . . . . . . . 28.5.1 Toxicity characteristic leaching procedure . 28.5.2 TCLP regulatory limits . . . . . . . . . . .

xxxiv

. . 903 . . . . .

. . . . .

903 904 906 906 909

. . . . . . . . .

. . . . . . . . .

916 918 922 924 924 930 931 931 932

. . 935 . . 935 . . 936 . . . . .

. . . . .

936 938 939 942 943

. . . . . . .

. . . . . . .

945 949 949 950 950 951 951

Contents

28.5.3 TCLP method summary. . . . . . 28.5.4 TCLP applications . . . . . . . . 28.5.5 TCLP and sequential extractions . 28.5.6 TCLP limitations . . . . . . . . . 28.6 Certiﬁed reference materials . . . . . . . 28.7 Summary and future developments. . . . 28.8 Useful World Wide Websites . . . . . . . Acknowledgements . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

Chapter 29. Sample preparation for semiconductor materials. . Katsu Kawabata, Yoko Kishi, Fuhe Li and Scott Anderson 29.1 Introduction. . . . . . . . . . . . . . . . . . . . . . 29.2 Contamination control . . . . . . . . . . . . . . . . 29.2.1 Clean room . . . . . . . . . . . . . . . . . . 29.2.2 Equipments, reagents and standards . . . . . 29.3 Sample preparation . . . . . . . . . . . . . . . . . . 29.3.1 Preparation and analysis of samples . . . . . 29.4 Conclusion . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . .

952 953 954 956 957 957 960 961 961

. 965 . . . . . . . .

965 969 969 971 976 978 986 986

TRACE ELEMENT SPECIATION

Chapter 30. Sampling and sample treatment in the analysis of organotin compounds in environmental samples . . . . . . Roberto Morabito 30.1 Introduction. . . . . . . . . . . . . . . . . . . . . . 30.2 Critical steps in organotin analysis . . . . . . . . . . 30.2.1 Sampling . . . . . . . . . . . . . . . . . . . 30.2.2 Storage . . . . . . . . . . . . . . . . . . . . 30.2.3 Sample treatment . . . . . . . . . . . . . . . 30.3 Improving the quality of organotin measurements in Europe . . . . . . . . . . . . . . . . . . . . . . . 30.4 Detailed procedure for the GC – MS determination of organotin compounds in environmental samples . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . .

. 991 . 991 . 994 . 994 . 998 . 1000 . 1005 . 1017 . 1020 . 1021

xxxv

Contents

Chapter 31. Sample preparation for arsenic speciation . . . . . . 1027 Walter Goessler and Doris Kuehnelt 31.1 Introduction. . . . . . . . . . . . . . . . . . . . . . . 1027 31.2 Occurrence and distribution of arsenic in the environment . . . . . . . . . . . . . . . . . . . 1028 31.2.1 Marine environment . . . . . . . . . . . . . . 1028 31.2.2 Terrestrial environment . . . . . . . . . . . . 1031 31.2.3 Humans . . . . . . . . . . . . . . . . . . . . . 1031 31.3 Stability of arsenic compounds . . . . . . . . . . . . . 1033 31.3.1 Arsenite and arsenate . . . . . . . . . . . . . 1034 31.3.2 Methylarsonous acid and dimethylarsinous acid . . . . . . . . . . . . . . . . . . . . . . . 1035 31.3.3 Methylarsonic acid and dimethylarsinic acid . . 1036 31.3.4 Arsenobetaine, arsenocholine, trimethylarsine oxide, and the tetramethylarsonium ion . . . . 1037 31.3.5 Arsenosugars . . . . . . . . . . . . . . . . . . 1037 31.4 Extraction of arsenic compounds from environmental samples . . . . . . . . . . . . . . . . . . . . . . . . . 1038 31.5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . 1041 References . . . . . . . . . . . . . . . . . . . . . . . . . . . 1041 Chapter 32. Sample preparation for speciation of selenium. . . . . . . . . . . . . . . . . . . . . Claudia Ponce de Leon, Anne P. Vonderheide and Joseph A. Caruso 32.1 Why selenium speciation?. . . . . . . . . . 32.2 General sample preparation . . . . . . . . 32.3 Mammals . . . . . . . . . . . . . . . . . . 32.3.1 Body ﬂuid analysis . . . . . . . . . 32.3.2 Tissue sample analysis . . . . . . . 32.4 Fish/birds . . . . . . . . . . . . . . . . . . 32.5 Plants . . . . . . . . . . . . . . . . . . . . 32.5.1 Leafy plants . . . . . . . . . . . . . 32.5.2 Broccoli . . . . . . . . . . . . . . . 32.5.3 Spices (garlic, onion, white clover) . 32.5.4 Grains. . . . . . . . . . . . . . . . 32.5.5 Nuts. . . . . . . . . . . . . . . . .

xxxvi

. . . . . . 1045

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. 1045 . 1045 . 1046 . 1046 . 1049 . 1050 . 1051 . 1052 . 1053 . 1053 . 1054 . 1054

Contents

32.5.6 Mushrooms . . . . . . . 32.6 Microorganisms . . . . . . . . . 32.7 Environmental . . . . . . . . . 32.7.1 Air. . . . . . . . . . . . 32.7.2 Water . . . . . . . . . . 32.7.3 Soil and sediments (solid References . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . matrices) . . . . . . .

. . . . . . .

. . . . . . .

. . . . . . .

. . . . . . .

. . . . . . .

. 1054 . 1055 . 1055 . 1055 . 1056 . 1058 . 1059

Chapter 33. Sample preparation for mercury speciation . . . . Holger Hintelmann 33.1 Introduction. . . . . . . . . . . . . . . . . . . . . . 33.2 Aqueous solution chemistry of methylmercury . . . . 33.3 Sample collection, preservation and storage . . . . . 33.3.1 Cleaning of sampling and laboratory equipment. . . . . . . . . . . . . . . . . . . 33.3.2 Water sampling . . . . . . . . . . . . . . . . 33.3.3 Preservation and storage of water samples . . 33.3.4 Preservation and storage of tissue and vegetation samples . . . . . . . . . . . . . . 33.3.5 Preservation and storage of soil and sediment samples . . . . . . . . . . . . . . . . . . . . 33.4 Sample preparation . . . . . . . . . . . . . . . . . . 33.4.1 Extraction of methylmercury from water . . . 33.4.2 Extraction of methylmercury from soils, sediments and particles . . . . . . . . . . . . 33.4.3 Extraction of methylmercury from biological tissue . . . . . . . . . . . . . . . . . . . . . 33.4.4 Direct techniques involving no sample preparation . . . . . . . . . . . . . . . . . . 33.4.5 Extraction of mercury species other than methylmercury . . . . . . . . . . . . . . . . 33.5 Quality control . . . . . . . . . . . . . . . . . . . . 33.5.1 Artifactual formation of methylmercury . . . 33.5.2 Spike recoveries. . . . . . . . . . . . . . . . 33.5.3 Reference materials . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . .

. 1063 . 1063 . 1063 . 1065 . 1066 . 1066 . 1067 . 1068 . 1068 . 1069 . 1071 . 1072 . 1073 . 1074 . 1074 . 1075 . 1075 . 1076 . 1077 . 1079 . 1079

xxxvii

Contents

Chapter 34. Sample preparation for speciation of lead . . . . . . 1081 Freddy C. Adams and Monika Heisterkamp 34.1 Introduction. . . . . . . . . . . . . . . . . . . . . . . 1081 34.2 Toxicity of organolead compounds . . . . . . . . . . . 1083 34.3 The history of leaded gasoline . . . . . . . . . . . . . 1085 34.4 Properties of organolead compounds . . . . . . . . . . 1088 34.5 Synthesis of organolead compounds . . . . . . . . . . 1090 34.6 The biogeochemical cycle of lead . . . . . . . . . . . . 1092 34.7 Analytical techniques for speciation analysis of organolead compounds . . . . . . . . . . . . . . . . . 1092 34.7.1 Hyphenated techniques for organometal determinations . . . . . . . . . . . . . . . . . 1093 34.7.2 Sample preparation . . . . . . . . . . . . . . . 1096 34.7.3 Sample pretreatment using gas chromatographic separation . . . . . . . . . . 1097 34.7.4 Extraction recovery . . . . . . . . . . . . . . . 1100 34.7.5 Separation . . . . . . . . . . . . . . . . . . . 1101 34.7.6 Detection of organolead compounds after chromatography. . . . . . . . . . . . . . . . . 1104 34.7.7 Procedures for the determination of organolead compounds in dust material . . . . . . . . . . 1107 34.7.8 Comparison of the different hyphenated systems . . . . . . . . . . . . . . . . . . . . . 1109 References . . . . . . . . . . . . . . . . . . . . . . . . . . . 1111

Chapter 35. Sample preparation for chromium speciation . . . Miguel de la Guardia and Angel Morales-Rubio 35.1 The element and its reactivity . . . . . . . . . . . . 35.2 The presence of Cr in nature and industrial processes 35.3 Chemical species of Cr present in real samples. . . . 35.4 Analytical methodologies available for determination of Cr . . . . . . . . . . . . . . . . . . . . . . . . . 35.5 Analytical methodologies for Cr speciation in water . 35.5.1 Speciation of chromium in aqueous matrices . 35.5.2 Types of samples analyzed . . . . . . . . . . 35.5.3 Pretreatments and techniques applied . . . .

xxxviii

. 1115 . 1115 . 1117 . 1118 . 1120 . 1120 . 1121 . 1121 . 1123

Contents

35.5.4 Speciation chromium using atomic spectrometry and MS-based techniques . . . . . . . . . . . . 1124 35.5.5 Determination of chromium speciation using molecular spectrophotometry . . . . . . . . . . 1130 35.6 Analytical methodologies for Cr speciation in biological ﬂuids . . . . . . . . . . . . . . . . . . . . . . . . . . 1136 35.6.1 Speciation of chromium in biological ﬂuids . . . 1136 35.6.2 Types of samples analyzed . . . . . . . . . . . 1137 35.6.3 Pretreatments and techniques applied . . . . . 1137 35.6.4 Chromium speciation using atomic spectrometry detection . . . . . . . . . . . . . 1138 35.6.5 Chromium speciation using molecular spectrophotometry detection . . . . . . . . . . 1143 35.7 Analytical methodologies for speciation of Cr in solid samples . . . . . . . . . . . . . . . . . . . . . . . . . 1145 35.7.1 Speciation of chromium in solid samples . . . . 1145 35.7.2 Types of samples analyzed . . . . . . . . . . . 1145 35.7.3 Solid sample treatments for speciation of chromium . . . . . . . . . . . . . . . . . . . . 1155 35.8 Final considerations . . . . . . . . . . . . . . . . . . 1157 References . . . . . . . . . . . . . . . . . . . . . . . . . . . 1158

Chapter 36. Sample preparation for metal-based drugs . Ronald R. Barefoot 36.1 Introduction. . . . . . . . . . . . . . . . . . 36.2 Platinum-based drugs. . . . . . . . . . . . . 36.2.1 Preparation of biological samples . . . 36.2.2 Cisplatin . . . . . . . . . . . . . . . 36.2.3 Carboplatin . . . . . . . . . . . . . . 36.2.4 Oxaliplatin . . . . . . . . . . . . . . 36.2.5 Ormaplatin . . . . . . . . . . . . . . 36.2.6 Lobaplatin . . . . . . . . . . . . . . 36.2.7 JM216. . . . . . . . . . . . . . . . . 36.2.8 Validated methods of analysis . . . . 36.3 Gold-based drugs . . . . . . . . . . . . . . . 36.3.1 Sample preparation . . . . . . . . . .

. . . . . 1173 . . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. 1173 . 1173 . 1175 . 1176 . 1177 . 1177 . 1178 . 1178 . 1178 . 1179 . 1180 . 1175

xxxix

Contents

36.4 Mercury . . 36.5 Vanadium . 36.6 Lead. . . . 36.7 Conclusions References . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

. . . . .

Chapter 37. Sample preparation for speciation analysis for metallobiomolecules . . . . . . . . . . . . . . . . . . . . . Joanna Szpunar, Brice Bouyssiere and Ryszard Lobinski 37.1 Introduction. . . . . . . . . . . . . . . . . . . . . . 37.2 Elemental species in biological systems: metallobiomolecules. . . . . . . . . . . . . . . . . . 37.3 Tailoring sample preparation: deﬁnition of the analyte moiety . . . . . . . . . . . . . . . . . . 37.4 Homogenization and analyte recovery using unreactive buffers . . . . . . . . . . . . . . . . . . . 37.5 Analyte recovery through partial degradation of sample matrix . . . . . . . . . . . . . . . . . . . 37.5.1 Extraction with protein denaturating reagents . . . . . . . . . . . . . . . . . . . . 37.5.2 Enzymatic extraction of organometallic compounds . . . . . . . . . . . . . . . . . . 37.5.3 Controlled enzymatic degradation prior to speciation of metal complexes. . . . . . . . . 37.5.4 Sequential enzymatic extractions for the evaluation of the bioaccessibility of metals in foodstuffs . . . . . . . . . . . . . . . . . . . 37.6 Fractionation of metal species according to the molecular weight prior to analytical chromatography or capillary electrophoresis . . . . . . . . . . . . . . 37.6.1 Ultraﬁltration . . . . . . . . . . . . . . . . . 37.6.2 Gel ﬁltration . . . . . . . . . . . . . . . . . 37.7 Multidimensional LC clean-up procedures prior to characterization of metal species by electrospray MS 37.8 Sample preparation prior to speciation analysis of biological ﬂuids . . . . . . . . . . . . . . . . . . . .

xl

. 1181 . 1181 . 1182 . 1182 . 1182

. 1185 . 1185 . 1186 . 1188 . 1188 . 1190 . 1190 . 1190 . 1191

. 1192

. 1193 . 1194 . 1194 . 1195 . 1196

Contents

37.8.1 Arsenic in urine . . . . . . . . . . . . . . . . . 1196 37.8.2 Selenium in urine . . . . . . . . . . . . . . . . 1197 37.8.3 Metal complexes in biological ﬂuids . . . . . . 1198 37.8.4 Metallodrug metabolites in biological ﬂuids . . 1199 37.9 Sample preparation prior to speciation analysis in solid matrices . . . . . . . . . . . . . . . . . . . . . . 1200 37.9.1 Organoarsenic species in marine biota and foodstuffs . . . . . . . . . . . . . . . . . . . . 1200 37.9.2 Low-molecular organoselenium species in yeast and plants. . . . . . . . . . . . . . . . . . . . 1201 37.9.3 High-molecular selenium species in animal tissues and yeast . . . . . . . . . . . . . . . . 1203 37.9.4 Metal complexes with metallothioneins. . . . . 1203 37.10 Sources of error . . . . . . . . . . . . . . . . . . . . . 1205 References . . . . . . . . . . . . . . . . . . . . . . . . . . . 1206 Chapter 38. Sample preparation for the analysis metals species . . . . . . . . . . . . . . . . Jo¨rg Feldmann 38.1 Introduction. . . . . . . . . . . . . . 38.2 Species of interest. . . . . . . . . . . 38.3 Characterization of gas samples . . . 38.4 Sample preparation strategies . . . . 38.4.1 Sampling . . . . . . . . . . . 38.4.2 Preconcentration . . . . . . . 38.5 Speciﬁc procedures . . . . . . . . . . 38.5.1 Cryotrapping methods . . . . 38.5.2 Solid phase micro extraction . 38.5.3 Adsorption method . . . . . . 38.6 Problems and future studies . . . . . References . . . . . . . . . . . . . . . . . .

of volatile . . . . . . . . . 1211 . . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. 1211 . 1212 . 1216 . 1216 . 1217 . 1217 . 1222 . 1223 . 1227 . 1227 . 1229 . 1230

Chapter 39. Sequential extraction . . . . . . . . . . . . . . . . . 1233 Angels Sahuquillo and Gemma Rauret 39.1 Sequential extraction procedures: a special case of sample dissolution . . . . . . . . . . . . . . . . . . . 1233

xli

Contents

39.1.1 A brief historical introduction . . . . . . . . 39.1.2 Deﬁnition of sequential extraction procedures 39.2 Types, uses and limitations of SEPs . . . . . . . . . 39.2.1 Characteristics of the extraction agents . . . 39.2.2 SEPs more widely used . . . . . . . . . . . . 39.2.3 Types of matrices and elements analysed. . . 39.2.4 Use of the information obtained . . . . . . . 39.2.5 Limitations of SEPs . . . . . . . . . . . . . . 39.3 Sample pre-treatment for SEPs. . . . . . . . . . . . 39.3.1 Drying. . . . . . . . . . . . . . . . . . . . . 39.3.2 Grinding and sieving steps . . . . . . . . . . 39.3.3 Use of inert atmosphere . . . . . . . . . . . 39.3.4 Recommendations. . . . . . . . . . . . . . . 39.4 Application of other extraction techniques to SEPs. . 39.4.1 Microwave . . . . . . . . . . . . . . . . . . 39.4.2 Ultrasound . . . . . . . . . . . . . . . . . . 39.4.3 Other alternatives . . . . . . . . . . . . . . 39.4.4 Conclusions . . . . . . . . . . . . . . . . . . 39.5 Quality control for SEPs . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . .

. 1233 . 1234 . 1235 . 1235 . 1235 . 1239 . 1241 . 1242 . 1246 . 1246 . 1248 . 1249 . 1250 . 1250 . 1250 . 1251 . 1251 . 1251 . 1252 . 1253

Index. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1257

xlii

Series Editor’s Preface This book on Sample Preparation for Trace Element Analysis, edited by Zoltan Mester and Ralph Sturgeon, is a useful addition to the Comprehensive Analytical Chemistry series. The impressive number of pages indicates the importance of sample preparation in the area of trace element determination. In a way, it follows the philosophy of a previous book in the series edited by Janusz Pawliszyn (Sampling and Sample Preparation for Field and Laboratory, vol XXXVII), and devoted to organic analysis. In that work, the two editors of this volume contributed a chapter on sample preparation for trace element speciation. It is a pleasure for me to introduce such a comprehensive book with a total of 39 chapters divided in four sections, including several introductory chapters on sampling, calibration, traceability and detection methods. These are followed by 17 chapters dealing with approaches to sample digestion and extraction. This is obviously one of the key issues in sample preparation, and for this reason a variety of chapters that include most of the methods in use – microwaves, solid phase microextraction, membrane extraction, laser ablation, ﬂow injection etc. – are presented. The ﬁnal 10 chapters cover speciﬁc applications to trace element speciation, dealing with different species and matrices, e.g. organotin, mercury, arsenic, metal-based drugs, chromium and also sequential extraction. The book includes a long list of recognised experts. In addition, many of them are previous contributors to books in this series dealing with speciation. In this respect, the present book is complementary to two previous volumes in the series – vol XXXIII on Elemental Speciation edited by Joe Caruso et al. and vol XXXIV on Discrete Sample Introduction Techniques for Inductively Coupled Plasma Mass Spectrometry by Diane Beauchemin and co-authors. With the publication of these three books the Comprehensive Analytical Chemistry series has extensively covered the area of elemental analysis, speciation and the very important bottleneck of sample preparation. I am sure that all three volumes will be a valuable reference for all researchers working in these ﬁelds. Finally I would like to thank not only the editors of the book but also the various authors for their contributions towards such a comprehensive, unique book on sample preparation for trace element analysis. Professor D. Barcelo´ Dept. of Environmental Chemistry IIQAB-CSIC Barcelona, Spain

xliii

Preface Two years ago we were asked to write a short review on sample preparation for trace metal speciation as a contribution to a book dealing with general sample preparation issues. Over the course of this work, we realized that this short review was rather an extended table of contents for a future project. We were also acutely aware that there was no comprehensive book devoted to sample preparation on the market dealing with the analysis of samples for trace elements. The stage was thus set. Following the collection of a sample, every analytical chemist will agree that its subsequent preservation and processing are of paramount importance. The availability of high performance analytical instrumentation has not diminished this need for careful selection of appropriate pretreatment methodologies, intelligently designed to synergistically elicit optimum function from these powerful measurement tools. These were the objectives of this book, to present, in a concise and comprehensive volume, an account of the state-of-the art of this subject matter. When considering the need for publication of a body of work such as this, it is wise to invest time appraising current literature; with the high cost of books, there can be no defense for simply making yet another one available. From our perspective, Sample Preparation for Trace Element Analysis was conceived because we believe there was no modern, comprehensive treatise at hand to satisfy the varied needs of the practicing analytical chemist. Without doubt, many of the subject areas targeted in this book have already received in-depth treatment by appropriate monographs. Assembling this knowledge into a single source proves advantageous to the user only if it is accomplished concisely and comprehensively. We hope the reader will vindicate our conclusions. This book is a multiauthor work, reﬂecting the diverse expertise arising from its highly qualiﬁed contributors. Efforts have been made to maintain a uniformity of style and diction, but readers will agree that the advantages which accrue from the talents of these individuals outweigh that arising from the simple uniformity gained with a single-author treatise. The cooperation of all the contributors in providing material for this book is thus deeply appreciated. The 39 chapters are authored by international leaders of their ﬁelds. The ﬁrst ﬁve chapters deal with general issues related to the determination of trace metals in varied matrices, such as sampling, contamination control, reference materials, calibration and detection techniques. The second part of

xliv

Preface

the book deals with extraction and sampling technologies (totaling 15 chapters), providing theoretical and practical hints for the users on how to perform speciﬁc extractions. Subsequent chapters overview seven major representative matrices and the sample preparation involved in their characterization. This portion of the book is heavily based on the preceding chapters dealing with extraction technologies. The last ten chapters are dedicated to sample preparation for trace element speciation. Dating from the original discussions with the Publisher, this book has been realised in record time, requiring less than two years to advance from concept to fruition, thanks to excellent work of the over 70 contributing authors and the efforts of the Publisher. The editors and authors hope that readers will ﬁnd this book useful and instructive and that it will be consulted frequently as a source of information which will make sample preparation less challenging for both the novice and seasoned expert alike. We wish to acknowledge the support of our home organization: the Institute for National Measurement Standards of the National Research Council of Canada, a stimulating environment and center of excellence for analytical chemistry research. Finally, we wish to thank the contributing authors for the privilege to work with them on this project and our families their patience and love for having forgone our company on many occasions. Zolta´n Mester Ralph E. Sturgeon

xlv

Acronyms 2-MBT 8-HQ AAS ACN ACP AED AFM AFS ANOVA ARC AsB AsC ASE ASTM ASV BEC CCD CCFA CCP CE CEA CGC CL CPG CPX CRM CSV CTF CV CV-AAS CZE-UV DAD DAL DBT DC arc emission

xlvi

2-mercaptobenzothiazole 8-hydroxyquinoline atomic absorption spectrometry acetonitrile alternating current plasma atomic emission detection atomic force microscopy atomic ﬂuorescence spectrometry analysis of variance anti-reﬂective coating arsenobetaine arsenocholine accelerated solvent extraction American Society for Testing and Materials anodic stripping voltammetry background equivalent concentration charge coupled device completely continuous ﬂow analysis capacitively coupled plasma capillary electrophoresis combustion elemental analysis capillary gas chromatography chemiluminescence controlled pore glass complexation certiﬁed reference material cathodic stripping voltammetry centrifugation coefﬁcient of variation cold vapour atomic absorption spectrometry capillary zone electrophoresis ultraviolet spectrophotometry diode array detector dialkyllead dibutyltin direct current arc emission spectrometry

Acronyms

DCP DCP-OES DE DEL DESe DIN DIW DLF-AAS DMA DMA(III) DMDSe DML DMSe DOM DP-ASV DPCSV DPhT DRC-ICP-MS DSI DTA DZ ECD ED-XRF EI-MS EL ESI Et ET-AAS: EtOH ETV EXT F-AAS FAB F-AES FIA FID FLD FPD FT-ICR-MS FTT GC(EC): GCB

direct current plasma direct coupled plasma optical emission spectrometry diatomaceous earth diethyllead diethyl selenide direct injection nebulizer deionized water diode laser ﬂame atomic absorption spectrometry. dimethylarsinic acid dimethylarsinous acid dimethyl diselenide dimethyllead dimethyl selenide dissolved organic material differential pulse anodic stripping voltammetry differential pulse cathodic stripping voltammetry diphenyltin dynamic reaction cell ICP-MS direct sample insertion diethylenetriamine dithizone electron capture detection energy dispersive X ray ﬂuorescence electron impact ionization mass spectrometry ethyl lactate electrospray ionization ethyl electrothermal (graphite furnace) atomic absorption spectrometry ethanol electrothermal vaporization liquid extraction ﬂame atomic absorption spectrometry fast atom bombardment ﬂame atomic emission spectrometry ﬂow injection analysis ﬂame ionization detection ﬂuorimetric detection ﬂame photometric detection fourier transform ion cyclotron resonance mass spectrometry ﬁltration gas chromatography with electron capture detection graphitized carbon black

xlvii

Acronyms

GC-MS GD-MS GF-AAS GLP GTF HEPA HG HMDE HMW HPA HPLC HRGC HR-ICP-MS HSAB HT18C6TO HTA IAEA IBMK IC ICP ICP-MS ICP-OES ICP-QMS ICP-RC-MS ICP-TOF-MS ID-MS ID-ICP-MS ID-TIMS INAA IP IR IRMM ISE ISO ITRS IUPAC KR LAS LC LEAF(S)

xlviii

gas chromatography-mass spectrometry glow discharge mass spectrometry graphite furnace atomic absorption spectrometry good laboratory practice glucose tolerance factor high efﬁciency particulate air hydride generation hanging mercury drop electrode high molecular weight high pressure ashing high performance liquid chromatography high resolution gas chromatography high resolution (sector ﬁeld) ICP-MS hard-soft acid-base hexathia-18-crown-6-tetraone high temperature ash International Atomic Energy Agency isobutyl methyl ketone ion chromatography inductively coupled plasma inductively coupled plasma mass spectrometry inductively coupled plasma (atomic) optical emission spectrometry ICP-quadrupole MS ICP-reaction cell-MS ICP-time of ﬂight-MS isotope dilution mass spectrometry isotope dilution inductively coupled plasma mass spectrometry isotope dilution thermal ionization mass spectrometry instrumental neutron activation analysis (NAA) ion pair infra-red Institute for Reference Materials and Measurements ion selective electrode International Organization for Standardization International Technology Roadmap for Semiconductors International Union of Pure and Applied Chemistry knotted reactor light absorption spectrometry (molecular UV-visible absorption) liquid chromatography laser excited atomic ﬂuorescence (spectrometry)

Acronyms

LIBS LiM LIMS LiT LLE LMW LOD LOV LRM LSASV LSE LTA MA MA(III) MALDI MBE MBT Me MEKC MeOH MIP MLS MMA MMLLE MOCVD MPD MPhT MPT MS MT MW NAA NCH NIES NIOSH NIST Nl NMR NRCC NSOM NWA ODETA OES OLR

laser induced breakdown spectrometry lithium metaborate, LiBO2 laboratory information management system lithium tetraborate, Li2B4O7 liquid-liquid extraction low molecular weight limit of detection lab-on-valve laboratory reference material linear sweep anodic stripping voltammetry liquid-solid extraction low temperature ashing methylarsonic acid methylarsonous acid matrix assisted laser desorption mass spectrometry molecular beam epitaxy monobutyltin methyl micellar electrokinetic chromatography methanol microwave induced plasma master laboratory station monomethyarsonic acid microporous membrane liquid-liquid extraction molecular organic compound vapor deposition microwave-induced plasma detector monophenyltin microwave plasma torch mass spectrometry metallothionein microwave neutron activation analysis neocuproine National Institute for Environmental Studies National Institute of Occupational Safety and Health National Institute of Standards and Technology measured gas volume in liter at 08C. nuclear magnetic resonance National Research Council of Canada near-ﬁeld scanning optical microscopy non-wetting agents 4-(N-octyl)diethylenetriamine optical emission spectrometry ordinary linear regression

xlix

Acronyms

OXI PA PAA PAAM PADMAP PAH PAR PBMS PDMS PE PEC PGC Ph PIXE PP PR PS-MS PTFE PTV PUF PVC P-XRF QA QCM QF-AAS QMS QTA QZ RCC REE RM RNAA ROMP RP RSD RTD SA SDS SEC SE-FLR SEP SF SFE SF-ICP-MS

l

oxidation polyacrylate photon activation analysis piconilic acid amide 2-(2-pyridylazo)-5-dimethylaminophenol polyaromatic hydrocarbon 4-(2-pyridylazo)-porphyrin performance based measurement system polydimethyl siloxane polyethylene power and event controller porous graphitized carbon phenyl proton induced x-ray emission spectrometry polypropylene photoresist plasma source mass spectrometry polytetraﬂuoroethylene programmed temperature vaporization polyurethane foam polyvinylchloride portable XRF quality assurance quality control material quartz furnace atomic absorption spectrometry quadrupole mass ﬁlters (heated) quartz tube atomizer quartz residual carbon content rare earth element reference material radiochemical separation neutron activation analysis ring-opening metathesis polymerization reverse phase relative standard deviation resistance temperature detector salicylic acid sodium dodecyl sulfonate size exclusion chromatography solvent extraction ﬂuorometry (molecular) sequential extraction procedure supercritical ﬂuid supercritical ﬂuid extraction sector ﬁeld ICP-MS

Acronyms

SGBM SI SI SIA SIM SIMS SLM SPE SPME SPS SRM SS-MS STAT T4BPP TAL TBT TCD TCLP TD TeAL TeEL TEL TeML TFA THET-AAS: THF TIMS TMAB TMAO TML TMOS TOF-MS TPB TPhT TprT TS-FF-AAS T-XRF UE ULPA UPW US US EPA UV-VIS

silica gel bound macrocycles Syste´me International sequential injection sequential injection analysis selected ion monitoring secondary ion mass spectrometry supported liquid membrane extraction solid phase extraction solid phase microextraction solid phase spectrophotometry standard reference material spark source mass spectrometry slotted tube atom trap tetra-(4-bromophenyl)-porphyrin trialkyllead tributyltin thermal conductivity toxicity characteristic leaching procedure thermodesorption tetraalkyllead tetraethyllead triethyllead tetramethyllead triﬂuoroacetylacetone transverse heated graphite atomizer ET-AAS (THGA: transverse heated graphite atomizer) tetrahydrofuran thermal ionization mass spectrometry tetramethylammonium bromide trimethylarsine oxide trimethyllead tetramethoxy silane time of ﬂight mass spectrometry tetraphenylborate triphenyltin tripropyltin thermospray ﬂame-furnace AAS total reﬂection XRF ultrasonic extraction ultra low penetration air ultrapure water ultrasound United States Environmental Protection Agency ultraviolet visible spectrometry

li

Acronyms

VG VMC VOCs VOL VPD WHO WLR XRA XRF ZHE

lii

vapor generation volatile metal(loid) compound volatile organic compounds volumetry (titrimetry) vapor phase deposition World Health Organization weighted linear regression X-ray absorption X-ray ﬂuorescence spectrometry zero headspace extraction

Chapter 1

Sampling and sample preservation for trace element analysis Byron Kratochvil

1.1

INTRODUCTION

Modern analytical methods and instrumentation make possible the measurement of increasingly smaller concentrations of even the most complex molecules and species in complex matrices. This has increased the importance of collecting, storing, and processing samples for analysis in a manner that keeps them as unaltered and contamination free as possible. In addition, improved measurement techniques and tools allow, or often require, the use of smaller analytical test portions to determine analyte concentrations. Small test portions mean more difﬁculty in achieving representativeness of the population, especially when analyzing for trace components. First of all, the quality of any analytical result depends on sample representativeness and integrity. Although many sources of error in an analysis can be controlled through use of blanks, standards, or reference samples, neither blank nor standard can repair the damage caused by an invalid sample. Keith [1], in the preface of a book on environmental sampling, says: “The logic is simple. If the right kinds of samples are not collected from the right areas at a site and then preserved, prepared, and analyzed correctly, wrong answers will be obtained. They may be precise and accurate answers, but they will be wrong in that they will not represent the condition of the site with respect to the absence, presence, or representative concentrations of the pollutants of interest.” Keith’s statement applies with equal validity to all analytical sampling operations regardless of analyte, concentration, or matrix. This chapter outlines some general principles of sampling design and sample preservation. Speciﬁc sampling and sample preparation procedures for various matrices and individual elements are treated in subsequent chapters. A brief bibliography and glossary of selected sampling terms are provided at the end of this chapter. Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

1

B. Kratochvil

1.2 1.2.1

PRELIMINARY CONSIDERATIONS Sampling variability

When obtaining an estimate of the uncertainty in an analytical result, the uncertainty in the sampling step is often signiﬁcant, and frequently far larger than the measurement uncertainty. For random errors, the overall standard deviation, so, is related to the standard deviation for the sampling operation, ss, and to that for the remaining analytical operations, sa, by: s2o ¼ s2a þ s2s

ð1:1Þ

Measurements should be designed insofar as possible to allow the separate evaluation of sample and measurement variability. For measurements in a state of statistical control, sa can be determined by the analysis of reference materials or standards. Then, ss can be obtained from Eq. (1.1), because so is obtained by analysis of a series of samples. Alternatively, a set of replicate measurements on samples may be designed to evaluate both sa and ss. Youden [2] noted that further reduction in the analytical uncertainty is unimportant once it is one-third or less of the sampling uncertainty. So, if the sampling uncertainty is large, use of a rapid, approximate analytical method may be faster, simpler, and permit more samples to be tested, thereby reducing overall uncertainty without increasing time or effort. 1.2.2

Sampling strategies

Sampling strategies may be classiﬁed as judgmental (intuitive), statistical, or systematic. Judgment sampling relies on general knowledge gained by experience with the population (or similar ones). Therefore, any conclusions drawn from the resulting data are necessarily intuitive, in part. Statistical sampling is based on all parts of the population having an equal chance of being selected. With a statistical sampling strategy, conclusions may be drawn based on statistical probabilities. In systematic sampling, the sample increments are collected in a regular pattern throughout the population. It has the advantage that execution is usually more straightforward and less expensive. Protocol sampling is a form of sampling speciﬁed in deﬁned circumstances, often by regulatory agencies or by groups, such as the American Society for Testing and Materials (ASTM), as a basis for decision-making in legal and commercial matters. For example, regulations may specify detailed sampling procedures, which, if not followed, could make the sample invalid for the intended purpose. The sampling procedure may be intuitive, statistical, or a combination, but must be followed explicitly.

2

Sampling and sample preservation for trace element analysis

1.2.3

Uncertainties in sampling

Sampling uncertainties may arise either from the properties of the population, and therefore inherent to any sample taken from it, or from the sampling operation itself. These uncertainties may be reduced, but never completely eliminated, by careful execution of a properly designed sampling plan that incorporates identiﬁcation of the population and sampling sites, along with the procedures required to deliver an uncontaminated, representative sample to the analytical laboratory. An important source of sampling uncertainty is systematic, non-random, bias caused by exclusion or inclusion in the sample of some components of the population over others owing to differences in size, mass, location, stickiness, and so on. Another is sample contamination or change during collection, transport, storage, or preparation for analysis (this topic is discussed in Section 1.6). Poor design or improper use of sampling equipment may also introduce bias, as may the omission of collateral measurements, such as ﬂow rate or pressures, that affect results. 1.3 1.3.1

TYPES OF SAMPLES Judgment samples

Judgment samples are samples collected from a population on the basis of experience, intuition, and knowledge of the history or properties of the population (or related ones). Sometimes, the goal is to obtain a single sample that may be termed “representative” to connote that it is expected to exhibit the average properties of the population. Collection of a single sample may have validity in situations where the population is essentially homogeneous or made so prior to sample collection. It may also be legitimate when random sampling is difﬁcult or impossible owing to safety or cost considerations. Under these conditions, however, the shortcomings of the sampling operation and the limitations in data treatment should be clearly stated. Generally, a plan based on at least some elements of random sampling is recommended. Judgment sampling requires assumptions about the degree to which the samples may be considered representative. Because the validity of the assumptions depends on the experience of the one making them, it is difﬁcult to know the degree to which they are acceptable for a given application. A major advantage of judgment sampling is that it is usually less costly than rigorous random sampling. For regulatory or legal purposes, however, personal bias should be reduced or eliminated as much as possible. Often a combination of judgment and random sampling provides the best compromise between unacceptable costs and data quality.

3

B. Kratochvil

1.3.2

Random samples

Analyses are almost always performed to obtain information about a population that is larger than the test portions being measured. If the samples under examination are biased, inferences made from them will be biased. The difference between the target population to which conclusions are applied, and the samples from which the test portions for analyses are drawn, may be minimized by selecting samples in a manner that gives each part of the population an equal chance of selection. This process, called random sampling, allows the user of the resulting analytical data to make statistical generalizations based on mathematical probabilities. Selecting truly random samples is difﬁcult; random in this context does not mean haphazard. A recommended method for a population consisting of units such as pharmaceutical tablets is to use random numbers to select units for analysis. Each unit is assigned a number, and units are selected by use of a random number generator.1 Bulk materials may be divided into a number of real or imaginary segments; the segments may be areas on a two-dimensional surface or volumes for a three-dimensional population. Data obtained by measurements on random samples can be analyzed by statistical methods to identify whether systematic relations among results exist due to trends or biases in the measurements. 1.3.3

Systematic samples

Because of its simplicity, sampling at evenly spaced intervals over a population is often used in place of random sampling. The criterion that all parts of the population have an equal chance of selection may be satisﬁed for evenly spaced sampling by imposing a random start time or sampling location on the process. This allows the application of classical statistical tests to the data. A potential problem with systematic sampling is that results may be biased if the analyte of interest is distributed in a periodic fashion within the population. It is also sometimes useful to collect samples in a systematic manner to reﬂect or test a hypothesis, such as the presence of systematic changes in population composition with time, temperature, or spatial location. Under speciﬁed conditions, each sample may be considered as a separate discrete population but the results may still be statistically tested for the signiﬁcance of apparent differences. 1

Random numbers may be obtained from several sources on the Internet. A good example is http://www.fourmilab.ch/hotbits/, which generates sequences of random numbers based on radioactive decay of krypton-85. A Geiger– Muller tube is interfaced to a computer and the times between successive pairs of radioactive decays measured and provided as bytes. Once the bytes are delivered, they are discarded.

4

Sampling and sample preservation for trace element analysis

1.3.4

Subsamples

Field samples are typically placed in containers and sent to the laboratory for further processing. Sometimes, transport of all the ﬁeld increments to the laboratory is deemed too inefﬁcient or costly. In this case, the increments may be homogenized, after crushing or grinding if needed, and subsampled on site prior to transport. The work needed to reduce particle size, blend, or otherwise to process a bulk ﬁeld sample before withdrawing subsamples for analysis depends on the variability in composition of the material constituting the original sample and on the extent of mixing required. Care must be taken to avoid contamination or loss that may introduce bias. Generally, processing and subsampling in a clean, controlled environment whenever possible provide better quality control. When subsampling is done in the ﬁeld, the sampling plan, discussed in Section 1.4.2, should specify that the sampler have sufﬁcient training and knowledge of sampling theory to subsample properly. Also, the analyst should be provided with all available information on prior subsampling and homogenizing operations. 1.3.5

Composite samples

Sometimes, increments are combined to produce a laboratory sample that is deﬁned as representative. Advantages of compositing include reduced sample handling and analytical effort. It provides an estimate of the average concentration of the analyte, but not of its distribution. A variety of sampling systems and mixing procedures have been developed to produce composites from both liquid and solid materials. Compositing of increments is attractive when costs of analytical measurements are greater than the costs of sampling. But potentially useful information, such as the presence of hot spots, may be lost. Analysis of individual increments allows not only estimation of the distribution of the analyte within the population, but also evaluation of apparent differences within and among samples. Garner et al. [3] discuss the advantages and limitations of composite sampling for environmental monitoring. 1.4 1.4.1

PLANNING THE SAMPLING OPERATION Deﬁning goals

Several key decisions should be made before sampling is initiated. These include deﬁning the population to be studied, the substance(s) to be measured, the precision required in the result, and the extent to which speciation and distribution within the population is needed. Any assumptions about the population should be clearly identiﬁed. Decision-makers should preferably

5

B. Kratochvil

Fig. 1.1. Elements of the overall analytical process.

include the client for the data, sampling personnel, the person responsible for the analytical work, and someone knowledgeable about statistics. Decisions made at this point establish the goals of the work, and are the ﬁrst step in the overall analytical process (Fig. 1.1). With this information in hand, a viable sampling plan can be drafted. 1.4.2

Sampling plans

The wide variety of populations sampled for chemical analysis makes the establishment of a single overall protocol impossible; accordingly, each matrix requires its own sampling plan. Often, regulatory agencies issue documents covering analytical methodologies that include sampling procedures. Examples include the US Environmental Protection Agency (US EPA), the International Organization for Standardization (ISO), and the ASTM. In addition, many specialty groups, such as the American Water Works Association, provide information on sampling protocols, tools, and techniques applicable to speciﬁc matrices. Where the analytical data may involve potential legal issues regarding compliance with environmental regulation, with workplace safety, or with commercial contract agreements, protocols recommended by recognized associations or agencies should be used whenever possible. All valid sampling plans and protocols have basic elements in common. These elements include speciﬁcation of the size, number, and location of sample increments, the extent of compositing where warranted, and steps for subsampling (after particle size reduction, if applicable, and mixing) of the initial increments to produce laboratory samples and test portions. The plan should be in the form of a written protocol that includes procedures for all steps, from initial sample collection to ﬁnal preparation of test portions for analysis. The protocol should tell when, where, and how to collect sample increments. It should include criteria for rejection of material not part of the population, as for example stones above a deﬁned size in a soil sample being analyzed for available trace nutrients. It should also specify who performs the sampling, sample logging and chain of custody procedures, the type and size of containers to be used, cleaning procedures for equipment and containers, preservatives, conditions of sample storage and, as appropriate, auxiliary information such as temperature or ﬂow velocity in a stream. It should also list the qualiﬁcations and training required of the personnel carrying out the operations. A checklist, adapted from Ref. [4], is provided in Table 1.1.

6

Sampling and sample preservation for trace element analysis TABLE 1.1 Checklist for elements of a sampling protocol (after Ref. [4]) Apparatus and equipment checklist Sampling tools and apparatus Sample containers of appropriate type, material, and size Cleaning supplies for tools, equipment, and containers Preservatives, including provision for cooling of samples if necessary Labels, tape, waterproof pens, packaging materials Chain of custody forms, sample seals, log books Safety equipment, including protective clothing

Instructions checklist for presampling Recording of observations at sampling sites Cleaning of apparatus before and after sampling Calibration of apparatus Cleaning and handling of sample containers Safety procedures Procedure if problems prevent strict adherence to protocol

Instructions checklist for sampling Number, type, and size of exploratory, regular, and quality assurance samples Number, type, and size of sample increments Procedure for identifying locations from which increments are to be collected Procedure for operation of apparatus and collection of increments Special sampling precautions or conditions of collection, including criteria for rejection of foreign material Procedure for compositing, if applicable Use of preservatives

Instructions checklist for postsampling Completion of auxiliary information on sample labels and in logbooks Chain of custody forms Sample packaging, transport, and conditions for travel and storage, including maximum holding time for samples prior to analysis

General Information on analytical methods, limits of detection, interferences

Once the sampling plan is drafted, it is worthwhile to have it reviewed by independent experts. This is especially important when assumptions have been made, or when all or part of the plan is based on judgment. For populations whose characteristics are little known, time and effort may be saved by collecting and analyzing a preliminary set of samples, using experience and intuition as a guide to make them as representative as possible. On the basis of this information, a more efﬁcient and cost-effective plan can be prepared.

7

B. Kratochvil

Where feasible, it is useful to have the analyst perform or supervise the sampling operation. Otherwise he or she should, in addition to helping prepare the written protocol, ensure that the sample collectors are well trained and understand the importance of each step so that bias and contamination are minimized. The training should emphasize the importance of accurate sample labeling and logging, and of a chain of custody to ensure sample integrity from collection to measurement. For bulk materials, local homogeneity affects sample size. Increments should be large enough to not be biased with respect to the different sizes and types of particles present in the material. Where available sampling equipment precludes collection of larger increments, two or more smaller ones may be taken adjacent to each other. These may be composited or analyzed separately. (Separate analysis can provide information on the extent of local heterogeneity.) When sampling a material whose properties are unknown, a good approach is to collect a small number of exploratory samples, using experience and judgment to make them as representative as possible, and analyze them for the substance of interest. From this preliminary information, a more reﬁned sampling plan can be developed.

1.5 1.5.1

STATISTICAL SAMPLING Introduction

Statistics provides a number of useful tools to assist in determining how many sample increments to take from a population, how large they should be, and from where they should be taken in order to hold the sampling uncertainty to some speciﬁed level with a given level of conﬁdence. Most statistical sampling theory is based on the population having a normal (Gaussian) distribution, but other distributions, such as lognormal, do occur in nature. 1.5.2

Minimum number of increments

Unless a population is known to be homogeneous, a valid sampling plan requires collection of increments from multiple locations. Assuming, for the moment, negligible measurement uncertainty relative to that for sampling, Provost [5] describes the minimum number of increments, ns, needed to hold the sampling uncertainty, Es, to a given level of conﬁdence by the relation: ns ¼ ðzss =Es Þ2

ð1:2Þ

where z is a stated level of conﬁdence, say 95%. In most applications, ss is either known from past history of the population or can be estimated from measurements on a set of preliminary samples to obtain values of ss and X.

8

Sampling and sample preservation for trace element analysis

(Remember that if measurement uncertainties are not negligible relative to those of the sampling operations, then ss should be calculated by Eq. (1.1).) Since p m ¼ 2X ^ ðts= n 1:3Þ where t is obtained from statistical tables as an estimate of z from n measurements, the maximum acceptable sampling uncertainty, Es, can be deﬁned by: p Es ¼ lm 2 Xl ¼ ts= ns ð1:4Þ Rearranging, ns ¼ ðtss =Es Þ2

ð1:5Þ

Initially, t can be set at 1.96 for 95% conﬁdence limits and a ﬁrst estimate of n can be calculated. The t-value for this n is then substituted and the system iterated to constant n. 1.5.3 Minimum size of increments in well-mixed particulate populations When sampling well-mixed populations of heterogeneous particles, as is often encountered in the subsampling of laboratory samples, Ingamells and Switzer [6] showed the relation: WR2 ¼ Ks

ð1:6Þ

to be applicable. Here W is the weight of sample analyzed, R is the relative standard deviation of sample composition in percent, and Ks is a constant equal to the weight of sample required to limit the sampling uncertainty to 1% relative with 68% conﬁdence. In practice, Ks is determined by estimating ss from a series of samples of weight W. Once Ks is known, the minimum sample weight, W, required for any maximum relative standard deviation can be calculated. For poorly mixed or stratiﬁed materials, the calculated value of Ks increases as W increases. This provides a way of testing the homogeneity of the population. When sampling a mixture of particles, it is important to collect enough of each particle type to ensure representativeness. In some cases, where the element under test is present as only a small fraction of the particles (as in elemental gold or diamond deposits), quite large bulk samples must be taken, and particle size reduction and thorough mixing must be conducted before subsampling. For such populations the sampling standard deviation, s(g1), may be calculated using the Johnson equation [7]: n hX io1=2 fi ð2ri Þ3 ð1:7Þ sðg1 Þ ¼ ðpd1 g1 =6Þ

9

B. Kratochvil

where g1 is the mass and d1 is the density of the sample particles containing the trace component, fi is the fraction by mass of the trace element in particle size class i, and ri is the radius of particles containing the trace element. If the element of interest is present in each of a mixture of two types of particles but the fraction of one type is small, Zheng and Kratochvil [8] have shown that a combination of the Johnson equation with one developed by Bennedetti-Pichler [9] is applicable. Here the standard deviation, sP, expressed in percent, is given by: n hX i o1=2 sP ¼ ½ðP1 2 P2 Þ=g ðpd1 =6Þ fi ð2ri Þ3 g1 ð1:8Þ where P1 and P2 are the percentages of the trace element in each of the two types of particles in the mixture, g is the mass of sample, g1 is the mass of the fraction of type 1 particles, and d1 is the density of the type 1 sample particles. The remaining terms are as deﬁned in Eq. (1.7). Equations (1.7) and (1.8) show that the sampling standard deviation varies as the square root of the sample mass and number of particles. This means that for every 10-fold decrease in the percentage of sought for substance, testportion size must increase 100-fold for a given level of sampling error and particle size. It is therefore especially important that laboratory samples for trace analysis are adequately ground and mixed prior to removal of test portions for trace analysis. The general approach described in this section has been extended by Gao and Kratochvil [10] to the calculation of sampling uncertainty for well-mixed materials containing more than two types of particles. 1.5.4

Sample increment size in segregated populations

Visman [11] demonstrated that for some segregated materials the variance of sampling could be expressed by:

s2s ¼ ðA=wnÞ þ ðB=n

1:9Þ

The constant A is related to Ingamells’ subsampling constant K and the average composition of the analyte, xav, by A ¼ 104xav. The constant B is related to the degree of segregation of the population. Values of A and B must be obtained experimentally from the bulk population. This can be done in two ways. In the ﬁrst, two sets of sample increments are collected, one with w as small as, and the other as large as, feasible. The two sets are analyzed, the sampling variances calculated and substituted into Eq. (1.9) to give values for A and B. In the second, arising out of published discussions by Duncan and Visman [12], Visman proposed collection of a set of increment pairs, each pair of increments being of the same weight and taken from adjacent sites in the population. From the analytical data on the increments, an intraclass correlation coefﬁcient, r, is calculated, either directly or by ANOVA [13]. Values for A and B are then

10

Sampling and sample preservation for trace element analysis

calculated from Eq. (1.9) and the relation r ¼ Bm=A; where m is the average particle mass. Increasing either W or N will reduce uncertainty due to random variability, but only increasing the number of increments, n, will reduce uncertainty due to segregation. All the sampling equations discussed in this section have been derived for normally distributed populations. As mentioned earlier, not all populations follow a Gaussian distribution. Procedures to test data for normality and for dealing with non-normality by data transformation or use of other procedures or distribution functions are available in the statistical literature. Problems may arise when small regions of a population contain analyte in much higher concentrations than elsewhere. This so-called “nugget” or “hot spot” effect is often encountered when sampling populations such as gold ores or contaminated industrial sites, but it can also be a factor in less obvious situations. An example is microanalytical investigation of surfaces using current sophisticated microtechniques. In situ analytical measurements on heterogeneous surfaces with a probe only a few micrometers in diameter may produce signiﬁcant errors if areas of unusually high or low concentration are missed or oversampled. There is also the danger that an unusually high result from a hot spot may be rejected as an anomalous outlier. The sampling plan should take into account the possibility of encountering hot spots and their potential effect on the goals of the sampling program. 1.5.5

From where should increments be taken?

The variety of populations of analytical, and therefore sampling, interest encompasses every part of nature and human activity. To ensure that all parts of a population have an equal chance of being selected for analysis requires a random element in the sampling strategy (see Section 1.3.2). Several strategies have been proposed to meet this requirement. These include, in addition to simple random sampling, systematic grid sampling with a random initial start point or with random sampling within individual grid areas or volumes. To improve sampling efﬁciency, other sampling schemes, including stratiﬁed, cluster, and two-stage sampling, have been developed. In simple random sampling, the target population is divided on paper into a set of units and a deﬁned number of the units are randomly selected for sampling. The units may be one dimensional, as a drill core or objects on a production line; two dimensional, as an agricultural ﬁeld or a surface ﬁlm coating on a manufactured product; or three dimensional, as a lake, railway tank car, or the atmosphere in an industrial plant. In systematic grid sampling, the population is divided into a two- or threedimensional grid and samples are collected from within each grid area or volume. Systematic sampling is often used to increase the probability of locating possible hot spots in a population. It has little inherent bias but may require more samples to be as effective as random sampling.

11

B. Kratochvil

In two-stage sampling, primary blocks or units are randomly selected within the population and two or more sample increments taken from locations within each unit. The locations may be selected systematically or randomly. Stratiﬁed random sampling involves division of the population into sections called strata. The number, size, and shape of strata are important to the design of an efﬁcient and cost-effective sampling plan. If the goal is to estimate more precisely the average analyte concentration in the population, then each stratum should be as uniform in the elements of interest as possible. This reduces the number of sample increments needed to deﬁne analyte distribution within each stratum. If analyte distribution among separate strata is of interest, then the sampling plan may involve judgment as to size and location of the strata. In cluster sampling, a number of increments are collected from one or more small sections of the population. This method is used when speciﬁc sections have been identiﬁed, either through judgment or by previous sampling, to be likely to contain more of the substance of interest. 1.5.6

Model-based sampling

The sampling equations discussed in previous sections are all based on classical sampling theory, such as described by Cochran [14] and others. This approach, sometimes called design-based sampling, makes no assumptions about the population other than that it is ﬁxed. Many sampling methodologies and statistical tools have been developed to handle various population distributions within this classical framework. A second approach, termed model-based sampling, employs one of several types of models to describe variability within a population. This methodology is most developed in the area of geostatistics. Borgman et al. [15] propose that, since the model-based approach views randomness as a property of a population, pure random sampling is no longer required and, in fact, may not be desirable because regularly spaced observations usually provide the best information about the degree of randomness present. A drawback is that the model must include information on expected patterns of variability within the population, though these patterns need not be completely understood to achieve reliable results. The biggest applications of model-based sampling have been for geostatistical estimations of underground ore reserves, but the method has also been applied to environmental studies [16]. A widely used form, called kriging, assumes a linear trend in concentration of the sought-for element. A sampling approach that includes elements of model design has been developed by Gy [17]. Although Gy employs classical random sampling statistics, he systematically considers all possible errors that might be encountered in the collection of a valid sample, including population variability, prior to sampling. In effect, Gy recommends incorporation of all uncertainties

12

Sampling and sample preservation for trace element analysis

that may affect representativeness of samples into the sampling design rather than assuming that randomness is the only source of variability. 1.5.7 Balancing economic factors and purpose of data collection against sample quality Sampling is often costly, especially in terms of time commitment by trained personnel. Therefore, the sampling plan should consider ways of minimizing the cost and variance of the sampling operation. Suppose a stratiﬁed sampling design is formulated consisting of n1 strata with n2 samples taken from each stratum and n3 analytical measurements on each sample. For strata equal in size and variance, the cost of determining a population mean to within a desired variance may be minimized as follows. The total cost of the operation, c, is equal to the sum of the cost of selecting the strata c1, sampling within the strata c2, and performing the analysis c3: c ¼ n1 c1 þ n1 n2 c2 þ n1 n2 n3 c3

ð1:10Þ

The overall variance for the population may be expressed as the sum of the variance contributions from the two stages of sampling and analyses: s2 ¼

s21 s22 s23 þ þ n1 n1 n2 n1 n2 n3

ð1:11Þ

Bennett and Franklin [18] show that to minimize the total cost for a preselected overall variance, the values of n1, n2, and n3 may be found from: qﬃﬃﬃﬃﬃﬃﬃﬃ s21 =c1 qﬃﬃﬃﬃﬃﬃ qﬃﬃﬃﬃﬃﬃ qﬃﬃﬃﬃﬃﬃ ð1:12Þ s21 c1 þ s22 c2 þ s23 c3 n1 ¼ s2 sﬃﬃﬃﬃﬃﬃﬃﬃ s22 c1 ð1:13Þ n2 ¼ s21 c2 sﬃﬃﬃﬃﬃﬃﬃﬃ s23 c2 ð1:14Þ n3 ¼ s22 c3 Note that the optimum allocation of sampling effort after the ﬁrst stage is independent of the desired overall variance. This means that when the goal is reduction in overall variance at minimum cost, one should increase the number of strata sampled and hold the other steps constant. Similarly, for a ﬁxed total cost; it was shown by Marcuse [19] that the optimum value for n1 is given by: qﬃﬃﬃﬃﬃﬃﬃﬃ c s21 =c1 ð1:15Þ n1 ¼ qﬃﬃﬃﬃﬃﬃ qﬃﬃﬃﬃﬃﬃ qﬃﬃﬃﬃﬃﬃ s21 c1 þ s22 c2 þ s23 c3

13

B. Kratochvil

while the optimum values for n2 and n3 continue to be given by Eqs. (1.13) and (1.14). Thus, the optimum allocation beyond the ﬁrst stage is the same for ﬁxed total cost as for ﬁxed total variance. The same principles can be applied to any number of stages in a nested sampling design. If strata are not equal in size or in distribution of the analyte, appropriate weighting factors must be incorporated into these expressions.

1.6

1.6.1

SAMPLE HANDLING AND PRESERVATION DURING COLLECTION, TRANSPORT, AND STORAGE Handling and storage of samples

Samples may undergo a variety of chemical or physical changes during collection, transport, storage, and preparation for analysis. Changes may include loss of sample through volatilization, chemical reactions among components of the sample, or reaction of sample components with sampling tools, sample containers, or transfer lines. Other sources of change include reactions of sample components with external agents such as oxygen, carbon dioxide, or water in the atmosphere, or with sampling equipment or containers. Decomposition during transport or storage may occur as a result of high temperatures or microbial action. Errors from these sources can be minimized by protecting samples from exposure to external agents, and by reducing rates of reaction through addition of preservatives and/or maintaining samples at low temperatures. Preservatives reduce decomposition by altering pH, redox conditions, or solubility; by converting species of interest into more stable forms; by blanketing or coating samples to prevent reaction; or by acting as biocides. Care must be taken that preservatives do not interfere with subsequent analytical measurements. In fact, the best preservation method is storage at temperatures that are as low as possible. Most materials may be stored without change for years at liquid nitrogen temperature (2 1968C), though this method is costly and often difﬁcult to implement. Since samples may begin to change from the time they are taken, analysis should ideally be done immediately after collection. Where the analysis involves digestion or extraction, consideration should be given to implementing this step promptly after collection, then storing the processed sample until measurement can be made. Procedures for sample collection, preservation, and storage are available from a variety of sources, such as the US Environmental Agency, for sampling of the environment, and the ASTM and ISO for industrial and commercial materials. An example of some of the recommendations provided by the US EPA for the evaluation of inland water and sediments is given in Table 1.2.

14

Suggested sample preservation and storage conditions for selected analyses of sediments and water [20] Analyte

Sample

In sediment Metals 100 g In water Metals

500 –2000 ml

Container

Preservation

Storage

Maximum holding time

Precleaned polyethylene jar

Refrigeration: dry ice or freezer for extended storage

#48C

Hg—28 days; others—6 months

pH , 2 with HNO3; refrigeration

28C

Zinc acetate, NaOH to pH . 9; refrigeration

48C

Hg—28 days; others—6 months 7 days

48C

28 days

Sulﬁde

250 ml

Acid-rinsed polyethylene or glass jar Plastic bottle

Fluoride

100 ml

Plastic bottle

Sampling and sample preservation for trace element analysis

TABLE 1.2

15

B. Kratochvil

1.6.2

Sampling equipment

A key component in any sampling operation is the quality of the apparatus used for the collection of sample increments and for splitting, grinding, or otherwise processing samples to obtain representative test portions for analysis. Koerner [21] discusses the selection and composition of equipment for environmental sampling. Good sampling equipment should be as simple as possible for ease of sample removal, maintenance, and use by different operators. It should protect the integrity of the sample by minimizing bias or loss of components during collection. The material of construction should be non-contaminating, durable, and readily cleaned. Plastics, especially Teﬂon or polyvinyl chlorides (PVCs), are often preferred over glass or metal for durability, ease of cleaning, and lower cost, but when strength and resistance to abrasion are needed, stainless steel is a good alternative. Several different stainless steel alloys are available. For example, type 316, which contains chromium, nickel, and molybdenum, has good resistance to corrosion and to sulfur compounds such as H2SO4. It will corrode with time when exposed to water containing iron-oxidizing microorganisms, however, especially at welds. Under these conditions, it is better to replace welds with threaded joints where possible. When sampling aqueous solutions for trace element determinations, sorption of analyte onto container walls must be considered. Parker et al. [22], in a study of the uptake of chromium, lead, and arsenic from groundwater by various materials, found that sorption was highest on types 304 and 316 stainless steel, followed by PVC. Teﬂon sorbed the least. 1.6.3

Sample containers

The purpose of a sample container is to protect the sample from interaction with its surroundings during transport and storage. Changes that may occur include loss of components to or contamination from the surroundings, as well as reaction with atmospheric components, especially oxygen, water, or carbon dioxide. Therefore, containers should have closures that seal completely, and do not introduce contamination into the sample. A study by Moody and Lindstrom [23] showed that, for most trace element analysis, containers of Teﬂon or linear polyethylene introduce the least contamination, but should be carefully cleaned with HCl and HNO3 before use. Of course, careful work requires that the purity of the acids and water used for cleaning and rinsing be considered too. Borosilicate glass is widely used for sample containers because it is relatively inexpensive, non-reactive to most organic compounds, and impermeable to gases. Though it sorbs a variety of molecules and ions, this is normally a problem only when the analyte is present in very low concentrations. Closures for borosilicate containers are usually screw caps with liners of Teﬂon or other inert plastic.

16

Sampling and sample preservation for trace element analysis

Most conventional plastics are sufﬁciently permeable to gases that they should not be used for the collection of gas samples or samples in which gas diffusion is undesirable. Fluorocarbons such as Teﬂon or Tedlar, however, have quite low permeability to gases. For this reason, and because of low sorption properties, they are often used as sample container materials when trace components are to be determined. 1.7

QUALITY ASSURANCE IN SAMPLING [24,25]

1.7.1

Overall objectives

The purpose of a quality assurance program in chemical measurements is to identify the nature and sources of errors in the overall analytical process, from sampling to data treatment, and to set up ways of assessing and minimizing those errors. The principal steps in reaching these goals are to: (1) assess the limits of error in the analytical data obtained from a sampled population; (2) reduce these errors to acceptable levels; (3) reduce the work required to obtain reliable data; and (4) provide, to the extent possible, a statistical basis for use of the data in decision-making. These objectives may be achieved through application of two concepts, quality control and quality assessment, which together comprise a quality assurance program. 1.7.2

Quality control

The goal of quality control is to attain a level of data quality that is adequate for the purpose, dependable, and economical. Quality control involves a system of testing and corrective actions that allows, through inspection, an estimate of the quality of the results. The system should specify whether changes are needed and, if they are, what measures should be taken to maintain a predetermined level of quality. Factors important to improved quality control in sampling include: † † † †

trained, knowledgeable samplers (who preferably have been involved in planning the sampling operations); a clear, complete, and detailed sampling protocol (including well-deﬁned criteria for rejecting foreign material); clean, well-maintained, and appropriate sampling tools and sampling containers; a sample management system that protects sample quality and integrity from collection through analysis.

1.7.3

Quality assessment

Quality assessment is a system of activities designed to ensure that the quality control job is being done effectively. It involves continuing evaluation of

17

B. Kratochvil

the quality control program. For analytical measurements, this may be done through test samples, interlaboratory comparisons, control charts, and so on, but for sampling it is not so simple. A sampling program may require that multiple increments be collected from both adjacent and widely spaced sites across the population to ensure representativeness and provide backup in the event of sample loss. It may also require the use of ﬁeld blanks and spiking of samples in the ﬁeld to detect bias from sample contamination, loss, or alteration. Sample integrity must be maintained through appropriate use of preservatives, containers, storage conditions, labeling, and logging. A good quality assessment program could include the following activities on a scheduled basis: † † † †

collection and comparison of data on replicate samples; external audit of sampling procedures and their execution in the ﬁeld, including appropriate safety precautions; review of sampling protocols, sample documentation procedures, and record keeping; thorough and objective feedback to all involved in sampling operations.

The overall aims of quality assurance in sampling are to provide a mechanism to reduce sampling errors to acceptable limits, the means to assure that the mechanism is operative, and the means to assure that the samples have a high probability of acceptable quality. Achieving these aims requires constant attention and maintenance, but with regular monitoring and review, a well-designed and implemented QA program can ensure quality of sampling operations indeﬁnitely.

1.8

GLOSSARY

Bulk sampling: sampling of a population that does not consist of discrete, identiﬁable, constant units, but rather of arbitrary, irregular units. Bulk sample: (also called gross sample, lot sample) one or more increments of material taken from a population for analysis or record purposes. Composite sample: a sample composed of two or more increments collected from different locations within a population or from the same location more than one time. Grab sample: (also called discrete sample) a single increment collected from a population at a speciﬁc time and location. Homogeneity: the degree to which a property or substance is randomly distributed throughout a material. Homogeneity depends on the size of the units under consideration. Thus, a mixture of two minerals may be inhomogeneous at the molecular or atomic level, but homogeneous at the particulate level.

18

Sampling and sample preservation for trace element analysis

Hot spot: a localized part of the population in which the analyte is present in signiﬁcantly higher concentration than elsewhere. Increment: a portion of material, collected by a single operation of a sampling device, from parts of a lot or population separated in time or space. Increments may be either analyzed individually or combined and tested as a composite sample. Individuals: conceivable constituent parts of the population. Laboratory sample: a sample, intended for testing or analysis, usually prepared from a bulk sample in one or more subsampling steps. The laboratory sample must retain the composition of the bulk sample. Reduction in particle size and mixing is typically necessary during its preparation. Lot: a quantity of units or bulk material of similar composition whose properties are under study. Population: a generic term denoting a collection of bulk material, individual items or events in the broadest concept; an aggregate determined by some property that distinguishes individuals that do and do not belong. Protocol: a detailed written description of the steps and procedures to be followed for the collection of valid samples. Reduction: the process of preparing one or more subsamples from a sample. Replicate samples: two or more samples collected from a population in an identical manner at the same time and place. Representative sample: a sample collected from a population in a manner that ensures, to the extent possible, that it accurately represents the population, or subset of the population, from which it was taken. Sample: a portion of a population or lot. It may consist of an individual or groups of individuals. Segment: a speciﬁcally demarked portion of a population, either actual or hypothetical. Spiked sample: a sample to which has been added a known quantity of the analyte to test the extent of interference by the matrix with the analytical measurement. Split sample: a sample divided into two or more representative parts for independent analysis. Strata: segments of a population that may vary with respect to the property under study. Subsample: a portion taken from a sample. A laboratory sample may be a subsample of a bulk sample; similarly, a test portion may be a subsample of a laboratory sample. Test portion: (also called specimen, test specimen, test unit, aliquot) that quantity of a material of proper size for measurement of the property of interest. Test portions may be taken from the bulk sample directly, but often preliminary operations such as mixing or further reduction in particle size are necessary.

19

B. Kratochvil

REFERENCES 1 2 3

4 5 6 7 8 9 10 11 12 13 14 15

16 17 18 19 20 21 22 23 24 25

20

L.H. Keith, in: L.H. Keith (Ed.), Principles of Environmental Sampling, 2nd ed., American Chemical Society, Washington, DC, 1996, p. xxvii. W.J. Youden, J. Assoc. Off. Anal. Chem., 50 (1967) 1007. F.C. Garner, M.A. Stapanian and L.R. Williams, in: L.H. Keith (Ed.), Principles of Environmental Sampling, 2nd ed., American Chemical Society, Washington, DC, 1996, p. 679. L.H. Keith, Environmental Sampling and Analysis, A Practical Guide. Lewis Publishers, Chelsea, MD, 1991, p. 14. L.P. Provost, Environmental Sampling for Hazardous Wastes, ACS Symposium Series 267, American Chemical Society, Washington, DC, 1984, p. 67. C.O. Ingamells and P. Switzer, Talanta, 20 (1973) 547; C.O. Ingamells and P. Switzer, Talanta, 23 (1974) 263. M.C.R. Johnson, Pharm. Acta Helv., 47 (1972) 546. L. Zheng and B. Kratochvil, Analyst, 121 (1996) 163. A. Benedetti-Pichler, in: W.M. Berl (Ed.), Physical Methods of Chemical Analysis, Vol. 3. Academic Press, New York, 1956, p. 183. Z. Gao and B. Kratochvil, Analyst, 126 (2001) 943 (see also p. 947). J. Visman, Mat. Res. Stds. 1969, November, pp. 8, 51, 62. J. Visman, A.J. Duncan and M. Lerner, Mat. Res. Stds., 11 (1971) 32; J. Visman, J. Mat., 7 (1972) 345. G.W. Snedecor and W.G. Cochran, Statistical Methods, 6th ed., Iowa State University Press, Ames, IA, 1967, pp. 294– 295. W.G. Cochran, Sampling Techniques. Wiley, New York, 1977. L.E. Borgman, J.W. Kern, R. Anderson-Sprecher and G.T. Flatman, in: L.H. Keith (Ed.), Principles of Environmental Sampling, 2nd ed., American Chemical Society, Washington, DC, 1996, p. 204. G.T. Flatman and A.A. Yfantis, in: L.H. Keith (Ed.), Principles of Environmental Sampling, 2nd ed., American Chemical Society, Washington, DC, 1996, p. 779. P.M. Gy, Sampling of Particulate Materials, Theory and Practice. Elsevier, New York, 1982. C.A. Bennett and N.L. Franklin, Statistical Analysis in Chemistry and the Chemical Industry. Wiley, New York, 1954, p. 490. S. Marcuse, Biometrics, 5 (1949) 189. “Required Containers, Preservation Techniques, and Holding Times”, Code of Federal Regulations, Title 40, Part 136, 1984, 49 FR 43260; Part 264, 40 CFR 264. C.E. Koerner, in: L.H. Keith (Ed.), Principles of Environmental Sampling, 2nd ed., American Chemical Society, Washington, DC, 1996, p. 155. L.V. Parker, A.D. Hewitt and T.E. Jenkins, Ground Water Monit. Rev., 10 (1990) 146. J.R. Moody and R. Lindstrom, Anal. Chem., 49 (1977) 2264. J.K. Taylor, Quality Assurance of Chemical Measurements. Lewis Publishers, Chelsea, MI, 1987. S.V. Kulkarni and M.J. Bertoni, in: L.H. Keith (Ed.), Principles of Environmental Sampling, 2nd ed., American Chemical Society, Washington, DC, 1996.

Sampling and sample preservation for trace element analysis

SELECTED BIBLIOGRAPHY G.E. Schweitzer and J.A. Santolucito (Eds.), Environmental Sampling for Hazardous Wastes, ACS Symposium Series 267, American Chemical Society, Washington, DC, 1984. A set of articles ranging from general aspects of sampling statistics and quality assurance to speciﬁc sampling problems in industry and in the environment. L.H. Keith (Ed.), Principles of Environmental Sampling, 2nd ed., American Chemical Society, Washington, DC, 1996. An excellent collection of chapters covering planning and sample design, statistical sampling, and quality assurance and quality control. Also included are chapters on speciﬁc elements to consider in sampling of air, water, biota, solids, and hazardous wastes. L.H. Keith, Environmental Sampling and Analysis, A Practical Guide. Lewis Publishers, Chelsea, MD, 1991. A guidance manual that focuses on those aspects of the sampling and analysis process required to produce data of a known quality. OSHA Technical Manual (TED 1-0.15A), U.S. Occupational Safety and Health Administration, U.S. Department of Labor, January 1999. Provides information relating to various aspects of sampling survey design and sampling practice. Section II of the manual provides guidance on sampling procedures. OSHA Chemical Sampling Information, U.S. Occupational Safety and Health Administration, U.S. Department of Labor. Contains a summary of sample collection parameters along with information on chemical properties and exposure limits. Available on-line and on CD-ROM. The on-line version is regularly updated. http://www.osha-sic.gov/SLTC/samplinganalysis/sampling.html. F.F. Pitard, Pierre Gy’s Sampling Theory and Sampling Practice, Heterogeneity and Sampling, Vol. I. CRC Press, Boca Raton, FL, 1989. J.K. Taylor, Quality Assurance of Chemical Measurements. Lewis Publishers, Chelsea, MI, 1987. A highly readable treatment of the basic concepts of quality assurance and how to plan and implement a quality assurance program for chemical analyses, including the sampling component.

21

This page intentionally left blank

Chapter 2

Sources of analyte contamination and loss during the analytical process G. Knapp and P. Schramel

2.1

INTRODUCTION

Contamination of the sample with the analyte and/or losses of the analyte from the sample are the most important systematic errors that can occur during preparation steps such as sampling, storage and preparation of the sample, decomposition, separation and analyte preconcentration and the ﬁnal determination of the elements. These problems are highlighted in many papers [1 –8]. Direct analytical procedures—such as neutron activation analysis (NAA) or X-ray ﬂuorescence spectrometry (XRF)—minimise these systematic errors, but often cannot be applied for many reasons, but most frequently because these methods require suitable standard reference materials for calibration. Unfortunately, there is a lack of such materials at low concentration levels and for a whole variety of matrices. Therefore, in most cases, combined multistep procedures have to be applied, which can lead to a variety of possible systematic errors. However, the most important advantage is the ease of calibration associated with wet chemical procedures based on aqueous standard solutions, by which the problem of the lack of reliable standard reference materials is overcome. Sampling has been a largely neglected area in trace element research for a long time. The research was mainly focused on the development of trace analytical methods for increasing sensitivity and selectivity. During the last 20 years, the analysts recognised more and more that the majority of systematic errors might be introduced during the analytical steps at the beginning of a combined analytical procedure and not with the ﬁnal measurement. A good analytical strategy also includes a sampling procedure free of contamination and losses, and proper stabilisation and storage of the samples. As analytical chemistry is a discipline that helps other disciplines to solve their problems, a close co-operation is necessary. In praxis, the analytical chemist is often not involved in the sampling procedures, the analyst is mostly Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

23

G. Knapp and P. Schramel

not even informed of the origin of the sample. Therefore, severe systematic errors in these ﬁrst steps of an analytical procedure are the consequence. Such considerations have been discussed by Kratochvil in Chapter 1 of this volume. The inﬂuence of contamination and losses on the analytical results becomes increasingly important with decreasing concentrations of the analyte, today down to the mg/kg(l) or even to the ng/kg(l) range. These effects depend not only on the concentration range but also on the nature of the analyte. In fact, elements that occur ubiquitously in higher concentrations create more difﬁculties in low concentration analysis than the other ones (a typical example is Al in blood, serum or tissues). On the other hand, losses during the analytical procedure depend on the chemical nature of the analyte (volatile compounds, etc.). Accuracy can never be directly measured, meaning that systematic errors cannot be detected easily. One indication of the presence of systematic errors is an unstable blank value. This may result from contaminations or losses during different analytical steps. Only contamination by the reagents can be taken into account, because it is reproducible, at least for one lot of a reagent. This chapter deals with sources of systematic errors caused by contamination and losses of elements during the single steps of a combined analytical procedure and highlights techniques for reducing of the inﬂuence of these errors on the analytical results. One should keep in mind that contamination and losses can never be completely eliminated, but they must be reduced to an acceptably low extent. It must be pointed out that this contribution cannot be comprehensive, but will illustrate the ways to minimise these systematic errors for special analytical procedures. Most of the examples given for the single analytical steps are for biological and clinical samples, but the ideas behind how to reduce contamination and losses in trace element analysis can easily be transferred to other sample matrices.

2.2

CONTAMINATION

Sources of contamination include materials from which tools and vessels are made reagents and laboratory air. These are individually discussed below. 2.2.1

Materials

Materials of vessels and tools are of great importance in trace element analysis. Contaminants can be leached out of the materials or desorbed from impurities at the surface of vessels and tools. Table 2.1 shows preferred materials and their use in trace element analysis. Vitreous silica, especially artiﬁcial quartz, is available in high purity and is one of the best materials for vessels in trace element analysis. The main

24

Preferred materials and their use for trace element analysis

Storage containers Beakers and ﬂasks Separatory vessels Vessels for wet digestion Vessels for bomb digestion Vessels for microwave digestion Crucibles Boats

PE and PP

FEP (#2008C)

þ þ þ

þ þ þ þ

PFA (#2508C)

TFM (#2508C)

PTFE (#2508C)

þ

þ þ

þ

þ þ þ

Vitreous silica (.5008C) þ þ þ þ þ þ þ

Glassy carbon (.5008C)

þ þ þ þ

Sources of analyte contamination and loss during the analytical process

TABLE 2.1

25

G. Knapp and P. Schramel

advantages include low risk of contamination, low wall adsorption due to a very smooth surface, temperature resistance up to 12008C and high resistance to most inorganic acids with exception of HF and concentrated phosphoric acid. Most microwave digestion devices have quartz vessels available. PTFE (Teﬂonw), PFA (Teﬂonw) and TFM (Hostaﬂonw) are ﬂuorinated polymers that are preferably used for digestion vessels. The non-polar surface of these materials minimises adsorption of polar ions and, therefore, contamination and losses by adsorption and desorption are low. The maximum digestion temperature for such vessels is about 2508C. Classical Teﬂon bombs made for use with conductive heating normally allow lower temperatures than Teﬂon vessels used for microwave heating. The reason for this is the cooling of microwave-heated vessels from the outside. The advantage of these materials is their resistance against nearly all acids and they can therefore be used for sample digestion with HF. Contamination from the vessel material is not a problem as long as vessels from reputable companies are used. Otherwise it can happen that recycled PTFE is used for vessel production and enhanced contamination risk is the consequence. In general, PTFE is not as good as the other two materials because of its porous structure, which arises from the sintering process used for the vessel production. TFM is a chemically modiﬁed PTFE and does not suffer from the porous structure of PTFE. FEP is an excellent material for storage containers because of its dense and non-polar surface. Losses of polar ions via adsorption effects can mostly be neglected. It is also used for liners of digestion bombs, but must not be heated above 2008C. PE and PP are successfully used for storage containers, beakers and ﬂasks, but they are not as good as FEP, although they are less expensive. Glassy carbon is used for high temperature digestion vessels. It is also resistant against most acids. Unfortunately, it is not resistant to oxidation. Therefore, it must be heated within an inert atmosphere. Additionally, oxidising reagents such as nitric acid signiﬁcantly attack the surface at elevated temperatures. Another disadvantage is a comparatively high risk of contamination; glassy carbon is not as clean as quartz glass or ﬂuorinated polymers. As already noted, one source of contamination by vessels and tools is the material itself. The other contamination source is impurities at the surface. To minimise or eliminate such impurities, proper cleaning procedures are necessary. Tables 2.2 and 2.3 summarise cleaning procedures used for polymers (PE and PP) and for Teﬂon [9]. The most efﬁcient cleaning procedure for quartz vessels is steaming with acid vapour [2]. This method can also be applied to vessels and tools made of borosilicate glass, PTFE, TFM, PFA and glassy carbon. For puriﬁcation, the vessels are continuously exposed to the hot vapour of the purifying liquid, frequently nitric acid. The principle of this apparatus is shown in Fig. 2.1.

26

Sources of analyte contamination and loss during the analytical process TABLE 2.2 Polymer cleaning procedure

Clean any residue from the polymer Rinse with DDI water Place in or ﬁll with 1:4 HCl for at least 1 week Rinse with DDI water Place in or ﬁll with 1:4 HNO3 for at least 1 week Rinse with DDI water Dry in clean air environment

A partly automated and very convenient device for steaming out vessels is produced by Milestone SLR, Sorisole, Italy. The suggested method for steaming vessels is given in Table 2.4. 2.2.2

Reagents

Signiﬁcant sources of contamination are the reagents. Gaseous reagents can easily be cleaned, but there are not so many methods where gaseous reagents can be used. Solid reagents on the other hand are difﬁcult to clean and result in comparatively high blanks. Liquid reagents are most important for trace analysis and they are commercially available in high purity grade (Suprapure grade reagents, Merck, Darmstadt, Germany; Optima grade reagents, Fisher Scientiﬁc; Double distilled grade reagents, Aldrich, Milwaukee, WI, USA; Ultrex II grade reagents, J.T. Baker Inc., Phillipsburgh, NJ, USA). The highest level of puriﬁcation can be obtained by sub-boiling distillation by means of stills constructed of quartz or PTFE [10]. Figure 2.2 shows the schematic of a subboiling still. Sub-boiling distillation can be used to purify the following reagents: H2O, HCl, HNO3, HF, H2SO4, NH4OH and organic solvents such as alcohols, chloroform, ketones, etc. A simple and inexpensive version of a subboiling still apparatus is shown in Fig. 2.3 (Savillex Corp., Minnetonka, MN, USA). Table 2.5 summarises the residual impurities in water and different acids. TABLE 2.3 Teﬂon cleaning procedure

Clean any residue from the Teﬂon Rinse with DDI water Place in 1:1 HCl acid bath (80–908C) for at least 4 h Rinse with DDI water Place in 1:1 HNO3 acid bath (80–908C) for at least 4 h Rinse with DDI water Dry in clean air environment

27

G. Knapp and P. Schramel

Fig. 2.1. Scheme of steaming apparatus for vessel puriﬁcation in acid vapour.

It should be noted that sub-boiling distillation ensures the separation of impurities of low vapour pressure such as metal ions, but it does not eliminate impurities having high vapour pressure such as organic compounds or some anions [10]. 2.2.3

Airborne particles

Contamination by laboratory air is a severe source of rising blank levels. There are three stages for reducing the inﬂuence of airborne particles on the TABLE 2.4 Procedure for steaming vessels

28

Steaming with HNO3 for at least 6 h (over night) Rinse with DDI water Dry in clean air environment

Sources of analyte contamination and loss during the analytical process

Fig. 2.2. Scheme of a sub-boiling distillation apparatus. 1: distillation chamber, 2: heating ﬁlament, 3: cooling device, 4: inlet funnel, 5: bottle with puriﬁed reagent, 6: reagent to be distilled.

analytical blank. The ﬁrst and least expensive step is to run the particular analytical step in a closed system. Figure 2.4 shows the scheme of a simple but effective device for evaporation of solutions. The next level of clean working area is a laminar ﬂow bench (Fig. 2.5). The most efﬁcient but also most expensive equipment to reduce contamination with airborne particles is a clean room (Fig. 2.6). Table 2.6 summarises the particulate concentrations in laboratory air in an ordinary laboratory, a clean room and a clean hood [11,12]. 2.3

LOSSES

Losses of elements are caused by volatilisation, chemical reactions or by reactions with the material of vessels and tools and, ﬁnally, by adsorption. In general, volatilisation can be prevented by application of closed systems (evaporation, closed vessel digestion, etc.). When closed systems are not suitable, volatilisation can be reduced or prevented by reducing the temperature (storage, freeze-drying, low temperature ashing, etc.). Reaction with the material of vessels and tools can be minimised by proper selection of the material and again by reducing the temperature (e.g. comparison of dry ashing and low temperature ashing). Chemical reactions can lead to precipitation of the analyte but frequently the addition of stabilising reagents (e.g. oxidising or chelating reagents) can prevent such unwanted effects. As described earlier, adsorption and desorption effects lead to losses or contamination. This physico-chemical phenomenon cannot be prevented, as

29

G. Knapp and P. Schramel

Fig. 2.3. IR-heated Teﬂon sub-boiling still in class-100 clean air hood constructed from modular PFA segments. TABLE 2.5 Blank levels in water and different acids in p.a. and suprapure grade from Merck, and after sub-boiling distillation [2] Element concentration (mg/l)

H2O HCl HCl HCl HNO3 HNO3 HNO3 HF HF HF

30

10 M 10 M 12 M 15 M 15 M 15 M 54% 40% 54%

Sub-boiling Sub-boiling Suprapure p.a. Sub-boiling Suprapure p.a. Sub-boiling Suprapure p.a.

Cd

Cu

Fe

Al

Pb

Mg

Zn

0.01 0.01 0.03 0.1 0.001 0.06 0.1 0.01 0.01 0.06

0.04 0.07 0.2 1.0 0.25 3.0 2.0 0.5 0.1 2.0

0.32 0.6 11 100 0.2 14 25 1.2 3.0 100

,0.05 0.07 0.8 10 ,0.002 0.7 0.5 0.5 3.0 4.0

0.02 ,0.05 0.13 0.5 ,0.002 0.7 0.5 0.5 3.0 4.0

,0.02 0.2 0.5 14 0.15 1.5 22 1.5 2.0 3.0

,0.04 0.2 0.3 8.0 0.04 5.0 3.0 1.0 1.3 5.0

Sources of analyte contamination and loss during the analytical process

Fig. 2.4. Evaporation chamber.

there is always a dynamic equilibrium. However, it is possible to signiﬁcantly reduce this effect to the extent that correct analytical results can be obtained. The lower the concentration range of the analysis, the more difﬁcult it is to reduce adsorption/desorption to an acceptable value. The extent of adsorption/desorption processes can be reduced by application of the following steps: † † † † †

choice of proper vessel materials; treatment of the vessel material; area of the vessel surface; single vessel principle; equilibration of the vessel surface.

The characteristics of vessel materials have been discussed earlier. For storage and analysis of trace metal ions, FEP, PFA and TFM are ideal materials because of their non-polar surface. The situation changes rapidly when organic complexes are formed, e.g. for separation and preconcentration methods. These complexes may possibly be adsorbed to the non-polar surfaces to a signiﬁcant extent. Artiﬁcial quartz is another excellent material for storage and analysis of metal ions, although the material is not as non-polar as the ﬂuorinated

31

G. Knapp and P. Schramel

Fig. 2.5. Laminar ﬂow bench.

polymers. Therefore, special treatment of the quartz surface is strongly recommended to deactivate some active sites at the surface and to reduce adsorption of metal ions. This happens during the steaming process with nitric acid [2]. Therefore, steaming of quartz vessels is essential because decontamination and deactivation of the quartz surface occur simultaneously. Another important rule in trace element analysis is to keep the surface of vessels as small as possible. In this manner, adsorption/desorption processes can also be reduced. Simply by taking a look at an analytical instrument, one can predict whether it is useful in trace analysis or not. Large vessels equipped with a large condenser, e.g., will lead to erroneous analytical results with high probability. In this same vein, there is another important rule, called the “single vessel principle”, which states: whenever possible keep the sample solution in one and the same vessel during all analytical steps. Each transfer to another vessel raises the probability of additional adsorption/desorption effects. As adsorption/desorption can never be completely eliminated, one can use dynamic equilibrium to keep the concentration of dilute solutions constant, at least for a certain time. This is important for calibration solutions having very low concentration levels. It can easily be observed, by repeated measurements

32

Sources of analyte contamination and loss during the analytical process

Fig. 2.6. Cross section of a clean room with a clean bench.

33

G. Knapp and P. Schramel TABLE 2.6 Element concentrations in laboratory air dust in an ordinary laboratory, in a clean room and in a clean hood Element concentration (mg/m3)

Ordinary laboratory Clean room Clean hood

Fe

Cu

Pb

Cd

0.2 0.001 0.0009

0.02 0.002 0.007

0.4 0.0002 0.0003

0.002 n.d. 0.0002

at intervals of several minutes, how quickly element concentrations can decrease when at the low mg/l level. Therefore, it is advisable to always use the same vessels for such highly diluted calibration solutions to maintain the dynamic equilibrium at that exact desired concentration level.

2.4

SAMPLING

Sources of contamination include all instruments and containers that come into contact with the samples, as they can cause contaminations to be introduced into the samples, either by mechanical abrasion or by leaching from the surfaces. Particularly in the case of tissues or body ﬂuids, it is especially difﬁcult to eliminate this kind of contamination when samples are taken from a living organism. Generally, it is not possible to intervene in routine clinical procedures where standardised equipment is used. Such possible contaminations of tissues and blood due to sampling have been investigated [13]. Elements such as Co, Cr, Cu, Fe, Mn and Ni, which are present in the material of surgical blades, biopsy needles and metal cannulas, can contribute to contamination. For Co, Mn and Ni, the errors are of the same order of magnitude as the element content in blood or serum. In the case of Cr, the analysis may be completely erroneous because contamination may be one order of magnitude higher than its content in blood or serum. Plastic containers, for example those used for sampling urine or water, may contain some plasticizers that can highly contaminate samples, especially when they have been acidiﬁed for stabilisation. A typical example is Cd. Each material used in the sampling procedure should be checked for a contamination risk under worst conditions, e.g. with dilute acid. In addition to contamination by vessels and tools, samples can also be contaminated by reagents, which have to be added as stabilisers, anticoagulants and preservatives [14 –19], especially because they are often added in excessive amounts.

34

Sources of analyte contamination and loss during the analytical process

A further type of contamination may occur by using the frequently applied “Vacutainer” in combination with a thin needle for taking blood samples. The vacuum inside the tube produces a high velocity of ﬂow through the needle. This will lead to a partial destruction of the erythrocytes, which produces haemolysis and with that contaminations of the serum or plasma with some of the elements that are present in higher concentrations in the whole blood, especially Fe, Zn and many others [20–22]. Contamination of blood can be prevented using the so-called “Braunu¨len” for taking blood samples. They consist of a steel needle in a plastic tube for penetrating the skin, which can be removed. Blood is then only in contact with the plastic tube. After rejecting the ﬁrst 5–10 ml of blood, one can obtain a blood sample nearly free from contamination. Because of the relatively large diameter (approximately 2 mm) of this sampling tool, these needles are not very popular among the patients. In general, for trace analysis, sampling tools made from materials other than stainless steel, such as titanium and ceramics for blades, or highly pure materials like nickel for needles should be used. In this case, there is only one element that cannot be analysed due to high blanks. Many other examples of such systematic errors can be found in Ref. [23]. Finally, errors may be the result of analytes being introduced into the sample via air dust or via the operating personnel, with particles of skin, perspiration, cosmetics, tobacco smoke, particles of clothing, washing powders, etc. being entrained in the material. These examples clearly demonstrate that adequate sampling is one of the most critical steps for an accurate analysis. Contamination sources arising from sampling procedures are most difﬁcult to identify and often remain unnoticed for a long time. One means of discovering the source of contamination is the application of different, independent sampling procedures. As already stressed, one principle to reduce or avoid contamination during sampling is to use tools or storage vessels made of materials which do not contain the analyte elements or contain them in very low concentration ranges. However, one cannot completely exclude contamination and therefore the principle of “controlled contamination” has to be applied appropriately for the actual analytical problem. Systematic errors caused by element losses due to volatilisation, adsorption or precipitation generally do not occur during the sampling stage. Mercury can be lost by precipitation on metal tools or by volatilisation at high temperatures, e.g. during sampling by drilling of solid materials. 2.5

STORAGE

Liquid samples, such as water, body ﬂuids, fruit juices, etc., are in a dynamic equilibrium state at the time of their collection. During storage, chemical, physical and/or microbiological processes affect the samples and can lead to

35

G. Knapp and P. Schramel

signiﬁcant changes in elemental concentrations. The factors responsible for changing the element concentration include adsorption and desorption of elements at or from container walls, chemical interactions with the container material, precipitation and losses by volatilisation. Important points that have to be considered in this context are the type of the container material, container pretreatment, temperature and duration of storage and the addition of stabilising reagents [24,25]. Leaching surfaces with acid presents the worst case for possible contaminants, but cannot be used as “blank” because it does not really reﬂect the conditions of the sample. This permits only a raw estimation of the possible dangers of contamination. As noted earlier, a wide variety of sample container materials for bottles, ﬂasks, tubes and vials can be used [24,26], the most common being polyﬂuorocarbons (PTFE and PFA) and other commercial products, polyethylene (PE), polypropylene (PP), polyvinyl chloride (PVC), polystyrene (PS), polycarbonate (PC), high purity quartz and borosilicate glasses. Synthetic polymeric materials can be recommended for achieving minimal contamination or losses. The purest materials were found to be conventional polyethylene, the various Teﬂonsw and polystyrene [27]. As a rule, cleaning procedures must be adopted to meet the standards required for trace element analytical work [27–32]. The procedures used should minimise the blanks but should not be too drastic so as to damage the surface of the container material and thus expose fresh and increased areas for chemical interactions, absorption and leaching [33]. Many cleaning procedures have been recommended [31], including use of different acids and their concentrations (mainly HNO3, HCl or HClO4), combinations of them but also non-ionic detergents and only distilled water. Again it depends on the problem at hand which method would be the most effective. Various papers [4,34 –42] describe observations on trace element exchanges between dilute aqueous solutions and their container materials. These results cannot be simply transposed to the biological ﬁeld. Serum, for example, has a high content of proteins, which bind numerous elements. Other problems may arise when serum samples are to be stored. Prolonged storage of small samples in plastic containers may lead to losses of water by diffusion through the wall [43], or by sublimation in the case of an inadequate seal [44]. The best way to store biological material is to maintain it at temperatures around 2 208C until analysis. But again, all such recommendations are very dependent on the element and its species. Every problem requires its own investigation. Urine and water samples have to be acidiﬁed (about 2–5 v/v % in HNO3) before storage to avoid losses due to wall adsorptions. However, it is very important, especially for frozen body ﬂuids, to thoroughly mix them (homogenise) after warming to room temperature in order to eliminate concentration gradients formed during the freezing process (Table 2.7) [45,46].

36

Sources of analyte contamination and loss during the analytical process TABLE 2.7 Concentration gradient for calcium, copper and sodium levels in serum, sweat and urine samples frozen and subsequently thawed without shaking [49] Position in tube

T M B C

Serum (meq/l)

Sweat (meq/l)

Urine (meq/l)

Ca

Cu

Na

Ca

Cu

Na

Ca

Na

0.25 1.10 3.60 2.20

17 90 127 89

26 126 270 142

0.09 0.17 0.41 0.31

63 143 364 250

11 23 71 42

4.6 9.8 15.9 10.7

69 152 204 152

Notes: T, top; M, middle; B, bottom; C, control (after shaking).

Drying has also often been used for stabilisation. There is no standardised method and all of these procedures are subject to losses of certain elements and must be again carefully studied for the particular element under consideration. Freeze-drying or lyophilization is the method considered least susceptible to element loss [47–50]. The risk of contaminations persists during the entire storage period. Thus, if samples cannot be analysed immediately, they should be kept in an adequately controlled environment or hermetically sealed in proper (plastic) material. To prevent element losses and/or transformation of the species, samples should be stored at low temperatures, sometimes at 2170 to 2 1908C (specimen banking). Losses of elements by volatilisation mainly occur at high temperatures. However, for very volatile elements such as Hg, As, Se, Sb and some others, this effect may be remarkably high even at room temperature [51,52]. As an example Hg may be lost within a few hours from an acidic solution [53]. Additionally, the use of plastic containers such as polyethylene or polypropylene cannot be recommended because Hg quickly penetrates such sample containers. Some general rules to prevent element losses or contamination during storage can be given. Samples should be stored at low temperatures in cleaned containers made of proper materials. Acidiﬁcation to pH , 2 with ultra-pure acids is a useful method for preservation of aqueous samples. The optimal conditions for sample storage ultimately depend strongly on the sample material and the analytical problem and must be investigated for every new analytical task. 2.6

DRYING AND HOMOGENISATION

Three methods are commonly used for drying different sample materials—oven drying, microwave drying and freeze-drying. Oven drying of biological material

37

G. Knapp and P. Schramel

is performed at temperatures around 1008C. It is very important to control the temperature because the biological matrix may decompose, depending upon the nature of the sample [54]. This decomposition results in a loss of residual dry matter and intrinsically volatile elements such as Hg, Sb, Se and others, may be lost. In an experiment described in Ref. [55], it was shown that the loss of dry residue was most pronounced for urine (31%), compared to blood or other soft tissues, where it was found in the range between 4 and 7%. Drying at lower temperatures will reduce volatilisation losses but also exposes the sample to the ambient environment for longer periods of time, which serves to increase the risk of contamination. Microwave drying, a very fast drying procedure, exhibits the same problems as oven drying. An exact control of the microwave energy is necessary to prevent overheating of the sample and losses of some elements [56]. Freeze-drying is most commonly used for trace element analysis of biological materials. It is also known as lyophilization or vacuum drying. To avoid losses of volatile elements, especially Hg or Se, it is recommended that the sample be cooled during lyophilization to ,108C. Without cooling, the temperature of the sample will increase to room temperature after sublimation of the moisture and this may again lead to losses. Modern freeze-drying instruments have an additional device to cool the sampler holder. More details can be found in the literature (e.g. [57]). To avoid systematic errors during the drying process, it is recommended that the original moist sample material be analysed whenever possible and to correct the analytical result with the factor obtained by the separate determination of the dry mass. In this case, losses of volatile elements or contamination by dust during the drying process is inconsequential. The next analytical step, i.e. homogenisation of solid materials, easily leads to contamination and/or losses of elements. Homogenisation of samples is, in many cases, necessary to provide a representative sample. The methods and tools used for this step are very dependent on the sample material. Containers and tools can lead to contaminations and losses as in the other analytical steps. For soft tissues, grinding and milling in vibrating ball mills applying PTFE or PFA containers and PTFE coated balls (made from stainless steel or tungsten, etc.), eventually under cooling with water or liquid nitrogen, are the preferred means. For hard materials, such as bone, teeth, etc., other container materials like Zr, Ti or W and cooling under liquid nitrogen are necessary. The use of mixers with stainless steel blades should be avoided in trace element analysis. On the one hand, there is a potential danger of contaminations and, on the other hand, the warming up of the sample may result in losses of volatile elements or compounds. For these instruments other materials for the blades, such as Ti, W or special ceramic materials, and water-cooling of the containers are commercially available. As mentioned earlier, the strategy of “controlled contamination” can be used. In every case, dependent on the given analytical problem, it is necessary and important to investigate systematically

38

Sources of analyte contamination and loss during the analytical process

possible errors in this step. It may be difﬁcult to verify, because there is nothing like a “blank”. The only way may be to analyse a piece of the sample before and after the homogenisation step. However, in this case, one has to keep in mind possible inhomogeneities in the original larger sample. All procedures that are commonly used in the production of CRMs cannot be used for trace element analysis in unknown samples. The ﬁnal CRM always shows traces of contaminations caused by the different production steps. It is only necessary that contaminations for such a material are homogeneously distributed. Most importantly for a CRM, it is the matrix that does not change and corresponds to the original material, but not the original concentration of some trace elements. At this stage it should be pointed out that, for all the analytical steps mentioned before, the use of a CRM for quality control is not permitted. The reason is that a CRM has passed all these steps during the production cycle, meaning that volatile elements were lost during the sample preparation steps such as drying and homogenisation, and are no longer available in the CRM for quality control of these analytical steps. For all subsequent analytical steps the use of an adequate CRM is strongly recommended for quality assurance and for method development. 2.7

DILUTION, DISSOLUTION AND DIGESTION

Dilution is important for direct measurement of aqueous samples such as serum, plasma, urine, milk, etc., especially for serum and plasma as they have a high salt (approximately 0.95%) and protein content (approximately 7%) and a high viscosity. This may create difﬁculties when a nebuliser is used as a sample introduction device because of changing nebulisation and transport efﬁciency [58]. Much attention must be given to the trace element impurities of the dilution reagent. Water, acids [59–62], Triton X-100 [63–69] or TMAH [70,71] are mainly used. Contamination control by running blank determination of the diluents is strongly recommended. Dissolution and digestion of samples are other important steps of the combined analytical techniques, which are sources of contamination and losses. Table 2.8 presents a survey of currently used dissolution and digestion techniques used in trace element analysis [72–77]. Fusion techniques should be avoided for trace element analysis whenever possible. Solid reagents can never be puriﬁed to such a high level as liquid reagents via sub-boiling distillation. Contamination by dust is also possible but can be normally neglected in comparison with contamination by fusion reagents. Due to the high fusion temperatures, losses of volatile compounds and reaction with the vessel material can lead to low analytical results. Fusion is not a viable sample decomposition technique for ultra-trace analysis.

39

G. Knapp and P. Schramel TABLE 2.8 Decomposition methods for trace element analysis

Fusion Combustion + In open systems + In closed systems Wet decomposition + In open systems + In closed systems + In ﬂow systems

The next family of sample decomposition techniques is combustion of organic materials with oxygen in open and closed systems. In principle, these techniques have the great advantage that gaseous reagents can easily be puriﬁed and no contamination from the reagent takes place. There is one exception—the dry ashing technique and unfortunately this technique is often still used in trace analytical laboratories despite its numerous sources of systematic errors. Continuous contamination by dust passing across the sample, together with the air necessary for oxidation, can occur. Another source of contamination is the mufﬂe oven itself. Volatile element compounds are evaporated from the hot surfaces. On the other hand, there are many possibilities for losses via evaporation of volatile compounds, like the halogens. The extent of volatilisation depends not only on the combustion temperature but also to a high degree of the sample matrix. The conclusion of a consideration of all these sources of systematic errors is that it is possible to generate correct analytical results using dry ashing, but the probability of erroneous analytical data is high and in these days there are much better sample digestion methods available. Combustion in closed vessels, the so-called oxygen ﬂasks, or in pressurised oxygen bombs is useful for the determination of volatile elements such as the halogens [78,79]. Contamination from the glass surface of the oxygen ﬂask or the stainless steel surface of the combustion bomb can be a problem. A general adsorption and desorption of elements must be taken into consideration because the relation of surface area to sample weight is very unfavourable with the oxygen ﬂask combustion. Only about 50 mg sample material can be combusted in a 500 ml ﬂask. There are many other combustion techniques described in the literature [80], but they are not signiﬁcant for trace element analysis, with one exception. The decomposition of organic samples in an oxygen plasma can be useful for ultra-trace analysis because of its extremely low contamination risk and practically no adsorption and desorption effects [81]. Only losses of volatile elements, such as Hg, As and Se, can be detected. Up to 2 g organic material can

40

Sources of analyte contamination and loss during the analytical process

be oxidised and the remaining ash dissolved in only 2 ml of diluted acid, preferably nitric acid. The most important family of sample digestion techniques is wet digestion. It is used for digestion in open vessels, in closed pressurised vessels and with automated ﬂow systems. All of these techniques can be equipped with conventional conductive heating or with microwave heating. With regard to sources of systematic errors, one erroneous comment must be eliminated, namely the comment that microwave techniques help to reduce systematic errors and therefore improve analytical results. Microwave irradiation does not reduce losses of elements and microwaves do not prevent or reduce contamination. All these systematic errors depend only on the reagents and vessel materials used and on the digestion method (open, closed, temperature, etc.). The only parameter that is signiﬁcantly inﬂuenced by microwaves is the reaction rate. Therefore, the digestion time is reduced dramatically, which is of great importance for routine analysis. Concerning systematic errors there is a big difference between the frequently used techniques of open and closed vessel wet digestion. Losses of elements, well known in open vessel wet digestion, cannot occur intrinsically with closed vessel digestion techniques when the vessel is tightly sealed. Only Hg can get lost when PTFE vessels are used. Mercury, in elemental form, diffuses into the porous structure of the PTFE surface. Ions are rejected from the non-polar surface. Elements can be volatilised in open vessel wet digestion as the element (Hg), as halogen compounds (As, B, Cr, Ge, Sn, Te, Ti, Zn, Zr), under oxidising conditions (Os, Pb, Rh, Ru) and under reducing conditions (Se, W). Contamination by the vessel material and by impurities of the surface can be minimised as described in Section 2.2.1. Vessel materials used in state-of the-art digestion methods include TFM and PFA as well as quartz glass. It must be kept in mind that the surface structure deteriorates with the number of digestion cycles and therefore adsorption and desorption effects increase [82]. By comparing open and closed vessel wet digestion in the context of contamination by reagents, it becomes obvious that closed vessel wet digestion is the method of choice for ultra-trace analysis. For the same amount of sample, much less digestion reagent is needed with closed vessel techniques. In addition, open vessel digestion usually requires sulphuric acid, which is disadvantageous for many measurement techniques.

2.8

SEPARATION AND PRECONCENTRATION

Analyte matrix separation and preconcentration should be avoided whenever possible, because these techniques are severe sources of losses and contamination. In principle, chemical separation techniques will be adopted for a number of different reasons: to concentrate analyte elements due to insufﬁcient

41

G. Knapp and P. Schramel

detection limits of the ﬁnal analytical method [83–85], to remove interfering species [78,86–88] or to eliminate matrix problems [89–99]. For the analysis of low-level trace elements, especially in human body ﬂuids, these procedures must be rigidly controlled because of their extreme susceptibility to contamination problems. Reagents other than those discussed earlier, such as, e.g., ion exchange resins, organic solvents, etc., have to be applied, and are thus also potential contaminants. It must be remembered that current manufacturing processes for many of these materials and reagents are not yet capable of yielding products containing only mg/kg levels of trace element impurities [84,95–103]. The other problem that has to be under control is the recovery of analyte during the procedure. It is not necessary that it is 100%, but it has to be constant. Problems with varying recoveries can be overcome by application of ID-MS calibration techniques. 2.9

ELEMENT MEASUREMENT

Instrumental analysis of the sample solution may also introduce severe problems with contaminations and losses. In most cases, an auto sampler is used and the sample solution has to be transferred into special auto sampler cups made of different materials. This is an additional source of contamination and losses. Contamination by the vessel material due to the manufacturing processes and adsorption and desorption effects due to the properties of the material may occur. All the cleaning and conditioning procedures mentioned before for vessels must also be applied to auto sampler vessels. A further source of contamination in this context is dust from the laboratory air. The samples, carefully prepared under clean conditions, stay open sometimes for many hours in the auto sampler. The atmosphere of the laboratory is loaded with particulate matter from different sources, which contaminates the sample. If the whole analytical procedure, including the ﬁnal measurement, cannot be done in a clean room, it is necessary to put at least the sample changer in a special clean box or in a laminar ﬂow box or zone. In the case of graphite furnace AAS, it may be necessary to keep the whole instrument in such a device, at least for the analysis of elements such as Al, Mn and others having a high environmental background level. On the other hand, losses may occur due to an insufﬁcient stabilisation of the sample solution. Elements such as Al, Cr or Sb need to be complexed with HF to avoid losses due to wall adsorption effects.

REFERENCES 1

42

E.J. Maienthal and D.A. Becker, National Bureau of Standards U.S. Technical Note No. 929. Government Printing Ofﬁce, Washington, DC, (1976).

Sources of analyte contamination and loss during the analytical process 2 3 4 5 6 7 8 9

10 11 12 13 14

15 16

17 18 19 20 21 22 23 24 25 26 27 28

29

30

P. Tscho¨pel, L. Kotz, W. Schulz, M. Veber and G. To¨lg, Fresenius Z. Anal. Chem., 302 (1980) 256. P. Tscho¨pel, Pure Appl. Chem., 54 (1982) 913. P. Tscho¨pel and G. To¨lg, J. Trace Microprobe Technol., 1 (1982) 1. J. Versieck, D. Barbier, R. Cornelis and J. Hoste, Talanta, 29 (1982) 973. J.R. Moody, Anal. Chem., 54 (1982) 1358A. J.R. Moody and E.S. Beary, Talanta, 29 (1982) 1003. J.R. Moody, Tr. Anal. Chem., 2 (1983) 116. H.M. Kingston, Quantitative Ultratrace Transition Metal Analysis of High Salinity Waters Utilizing Resin Separation. National Bureau of Standards and the Environmental Protection Agency, (1979). E.C. Kuehner, R. Alvarez, P.J. Paulson and T.J. Murphy, Anal. Chem., 44 (1972) 2050. E.J. Maienthal, U.S. National Bureau of Standards Technical Note No. 545, 1970 T.J. Murphy, The role of the analytical blank in accurate trace analysis, National Bureau of Standards Special Publication 4222, 1976. J.M.J. Versieck and A.B.H. Speecke, Nuclear Activation Techniques in the Life Sciences 1972. IAEA, Vienna, 1972, p. 39. N.W. Alcock, in: McC. Howell, J.M. Gawthorne and C.L. White (Eds.), Trace Element Metabolism in Man and Animals, Vol. 4. Australian Academy of Science, Canberra, 1981, p. 678. A. Astrug, V. Kukeva, I. Kuleff, Z. Kiriakov, A. Tomov and R. Djingova, Trace Elements Med., 1 (1984) 65. W.F. Bethard, D.A. Olehy and R.A. Schmitt, in: D. Comar (Ed.), L’Analyse par Radioactivation et ses Applications aux Sciences Biologiques. Presses Universitaires de France, Paris, 1964, p. 379. H.J.M. Bowen, Trace Elements in Biochemistry. Academic Press, New York, 1966, Ch. 5. G.V. Iyengar, H. Borberg, K. Kasperek, J. Kiem, M. Siegers, L.E. Feinendegen and R. Gross, Clin. Chem., 25 (1979) 699. E.W. Reimold and D.J. Besch, Clin. Chem., 24 (1978) 675. D.J. Kosman and R.I. Henkin, Lancet, 1 (1979) 1410. R.T. Lofberg and E.A. Levri, Anal. Lett., 7 (1974) 775. D. Mahanand and J.C. Houck, Clin. Chem., 14 (1968) 6. J. Versieck and R. Cornelis, Trace Elements in Human Plasma or Serum. CRC Press, Boca Raton, FL, 1989. T.D. Spittler and J.B. Bourke, in: L.H. Keith (Ed.), Principles of Environmental Sampling. American Chemical Society, Washington, DC, 1988, p. 375. V.D. Anand, J.A. White and H.V. Nino, Clin. Chem., 21 (1975) 595. B. Sansoni and G.V. Iyengar, IAEA Tech. Rep. Ser. No. 197, Vienna, 1980, p. 57. J.R. Moody and R.M. Lindstrom, Anal. Chem., 49 (1977) 2264. A. Speecke, J. Hoste and J. Versieck, in: P.D. LaFleur (Ed.), Accuracy in Trace Analysis: Sampling, Sample Handling, Analysis, Vol. 1. National Bureau of Standards, U.S. Department of Commerce, Washington, DC, 1976, p. 299. T.J. Murphy, in: P.D. LaFleur (Ed.), Accuracy in Trace Analysis: Sampling, Sample Handling, Analysis, Vol. 1. National Bureau of Standards, U.S. Department of Commerce, Washington, DC, 1976, p. 509. C.C. Patterson and D.M. Settle, in: P.D. LaFleur (Ed.), Accuracy in Trace Analysis: Sampling, Sample Handling, Analysis, Vol. 1. National Bureau of Standards, U.S. Department of Commerce, Washington, DC, 1976, p. 321.

43

G. Knapp and P. Schramel 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48

49 50 51 52 53 54 55

56 57 58 59 60 61 62 63 64 65 66

44

D.P.H. Laxen and R.M. Harrison, Anal. Chem., 53 (1981) 345. M. Zief and J.W. Mitchell, Contamination Control in Trace Element Analysis. Wiley, New York, 1976, Ch. 4. G. To¨lg, Z. Anal. Chem., 283 (1977) 257. A.W. Struempler, Anal. Chem., 45 (1973) 2251. G.G. Eichholz, A.E. Nagel and R.B. Hughes, Anal. Chem., 37 (1965) 863. E.S. Gladney and W.E. Goode, Anal. Chim. Acta, 91 (1977) 411. R. Massee, F.J.M.J. Maessen and J.J.M. de Goeij, Anal. Chim. Acta, 127 (1981) 181. R.V. Moore and G.W. Leddicotte, in: D.D. Hemphill (Ed.), Trace Substances in Environmental Health, Vol. 2. University of Missouri, Columbia, MO, 1968, p. 243. D.E. Robertson, Anal. Chim. Acta, 42 (1968) 533. A.D. Shendrikar and P.W. West, Anal. Chim. Acta, 72 (1974) 91. A.D. Shendrikar, V. Dharmarajan, H. Walker-Merrick and P.W. West, Anal. Chim. Acta, 84 (1976) 409. A.D. Shendrikar and P.W. West, Anal. Chim. Acta, 74 (1975) 189. K. Heydorn, In: Neutron Activation Analysis for Clinical Trace Element Research, Vol. I. CRC Press, Boca Raton, FL, 1984, Ch. 2. E.W. Reimold and D.J. Besch, Clin. Chem., 24 (1978) 675. D.H. McGlory, Clin. Chem., 17 (1971) 1074. S.H. Omang and O.D. Vellar, Fresenius J. Anal. Chem., 269 (1974) 177. J.J.M. de Goeij, K.J. Volkers and P.S. Tjioe, Anal. Chim. Acta, 109 (1979) 139. D. Shapcott, In: Chromium in Nutrition and Metabolism. D. Shapcott and J. Hubert (Eds.), Elsevier/North-Holland Biochemical Press, Amsterdam, 1979, p. 43. D. Shapcott, K. Khoury, P.-P. Demers, J. Vobecky and J. Vobecky, Clin. Biochem., 10 (1977) 178. P.D. LaFleur, Anal. Chem., 45 (1973) 1534. K. Ba¨chmann and J. Rudolph, J. Radioanal. Chem., 32 (1976) 243. P. Reh and J. Gaede, Fresenius J. Anal. Chem., 343 (1992) 715. G. Kaiser, D. Go¨tz, P. Schoch and G. To¨lg, Talanta, 22 (1975) 889. H.J.M. Bowen, J. Radioanal. Chem., 19 (1974) 215. G.V. Iyengar, K. Kasperek and L.E. Feinendegen, 3rd Int. Conf. Nuclear Methods in Environmental and Energy Research, University of Missouri, Columbia, Oct. 10– 13, 1977. B. Maichin, P. Kettisch and G. Knapp, Fresenius J. Anal. Chem., 366 (2000) 26. International Atomic Energy Agency, Technical Reports Series No. 197, 1980 P. Schramel and J. Ovcar-Pavlu, Fresenius Z. Anal. Chem., 298 (1979) 28. J.W. Mitchell, J. Radioanal. Chem., 69 (1982) 47. J.W. Mitchell, Anal. Chem., 45 (1973) 492A. V.C. Smith, in: M. Zief and R. Speights (Eds.), Ultrapurity, Methods and Techniques. Marcel Dekker, New York, 1972, p. 173. M. Zief and J.W. Mitchell, Contamination Control in Trace Element Analysis. Wiley, New York, 1976, Ch. 5. A.J. Schermaier, L.H. O’Connor and K.H. Pearson, Clin. Chim. Acta, 152 (1985) 123. M. Drazniowsky, I.S. Parkinson, M.K. Ward, S.M. Channon and D.N.S. Kerr, Clin. Chim. Acta, 145 (1985) 219. G. Brodie and M.W. Routh, Clin. Biochem., 17 (1984) 19. L.J. Hinks, B.E. Clayton and R.S. Lloyd, J. Clin. Pathol., 36 (1983) 1016.

Sources of analyte contamination and loss during the analytical process 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103

Y. Pegon, Anal. Chim. Acta, 172 (1985) 147. P.A. Pleban and K.H. Pearson, Clin. Chem., 25 (1979) 1915. D.A. Thompson, Ann. Clin. Biochem., 17 (1980) 144. P. Schramel and S. Hasse, Mikrochim. Acta, 116 (1994) 205. G. Knapp, B. Maichin, P. Fecher, S. Hasse and P. Schramel, Fresenius J. Anal. Chem., 362 (1998) 508. R. Bock, Handbuch der Analytisch-Chemischen Aufschlussmethoden. Wiley-VCH Verlag GmbH, Weinheim, 2001. T.T. Gorsuch, The Destruction of Organic Matter. Pergamon Press, Oxford, 1970. J. Dolezal, P. Povondra and Z. Sulcek, Decomposition Techniques in Inorganic Analysis. Elsevier, New York, 1968. H.M. Kingston and S.J. Haswell, Am. Chem. Soc., Washington, DC, 1997. G. Knapp, Mikrochim. Acta, (1991) 445. G. Knapp, Anal. Proc., 27 (1990) 112. M. Reverchon, Chim. Anal. Paris, 44 (1962) 340. A.L. Conrad, Mikrochemie, 38 (1951) 514. R. Bock, A Handbook of Decomposition Methods in Analytical Chemistry. International Textbook Comp. Ltd., London, 1979. R.T. White and C.W. Lawrence, J. AOAC Int., 78 (1995) 99. H.M. Ortner, S. Sterkel, G. Knapp, B. Maichin, P. Kettisch, L. Kocsis, J. Mihaly and J. Mink, Mikrochim. Acta, 137 (2001) 229. E.M. Bem, Environ. Health Perspect., 37 (1981) 183. M. Alt and H. Massman, Z. Anal. Chem., 279 (1976) 100. H.T. Delves, G. Shepherd and P. Vinter, Analyst, 96 (1971) 260. J. Kumpulainen, A.M. Raittila, J. Letho and P. Koivistoinen, J. Assoc. Off. Anal. Chem., 66 (1983) 1129. K. Heydorn and H.R. Lukens, Danish Atomic Energy Commission, Research Establishment Risø, Roskilde, (1966). H. Bem, J. Holzbecher and D.E. Ryan, Anal. Chim. Acta, 152 (1983) 247. R.O. Allen and E. Steinnes, Anal. Chem., 50 (1978) 1553. I. Andersen and A.C. Høgetveit, Fresenius Z. Anal. Chem., 318 (1984) 41. I. Andersen, W. Torjussen and H. Zachariasen, Clin. Chem., 24 (1978) 1198. R.A. Barfoot and J.G. Pritchard, Analyst, 105 (1980) 551. D.J. Halls, G.S. Fell and P.M. Dunbar, Clin. Chim. Acta, 114 (1981) 21. R. Heinrich and J. Angerer, Fresenius Z. Anal. Chem., 315 (1983) 528. E. Kamata, R. Nakashima, K. Goto, M. Furakawa and S. Shibata, Anal. Chim. Acta, 144 (1982) 197. N. Lekehal and M. Hanocq, Anal. Chim. Acta, 83 (1976) 93. Ne`ve, M. Hanocq and L. Molle, J. Pharm. Belg., 35 (1980) 345. M. Suzuki, K. Hoyashi and W.E.C. Wacker, Anal. Chim. Acta, 104 (1979) 389. M. Suzuki and W.E.C. Wacker, Anal. Biochem., 57 (1974) 605. R. Cornelis, J. Versieck, L. Mees, J. Hoste and F. Barbier, J. Radioanal. Chem., 55 (1980) 35. H.T. Delves, Clin. Chim. Acta, 71 (1976) 495. P.E. Gardiner, J.M. Ottaway, G.S. Fell and R.R. Burns, Anal. Chim. Acta, 124 (1981) 281. R. Pietra, E. Sabbioni, A. Springer and L. Ubertalli, J. Radioanal. Chem., 69 (1982) 365.

45

This page intentionally left blank

Chapter 3

Calibration approaches for trace element determination Douglas C. Baxter and Ilia Rodushkin

3.1

INTRODUCTION

Perhaps 95% of all errors in analytical measurements occur during sample preparation. Avoiding or accounting for these errors is a major theme of this chapter. If it is assumed that the samples have been correctly prepared, then it becomes necessary to ensure that appropriate steps are taken to avoid the remaining sources of errors that are most likely to arise during calibration. Although the primary goal of calibration is to facilitate accurate calculation of analyte concentrations or absolute amounts in the samples, it is no less important to be able to provide a meaningful assessment of the associated uncertainties. The latter allows statistical evaluation of the data, for example, to test whether the measured quantity meets product speciﬁcations or conforms to legislative limits [1]. Note that the general use of the term analyte concentration in the following does not exclude the possibility that the absolute amount of analyte may actually be the quantity of interest. In this respect, the two terms can be considered interchangeable in most contexts below. The quantity to be derived, from calibration of the instrument and subsequent analysis of a test portion of the sample material, is more generally denoted as the measurand. This chapter will ﬁrst introduce the basic assumptions and terminology (Section 3.2) used in the context of calibration. Next, some criteria for selecting the calibration approach are presented in Section 3.3, one of these being recovery, the statistical evaluation of which is summarized in Section 3.4. This is followed by a description of linear regression techniques (Section 3.5), which provide the analytical chemist with some of the most important, yet poorly understood, tools of the trade. Their application to external calibration (Section 3.6), the method of standard additions (Section 3.7) and internal standardization (Section 3.8) is then detailed. For each of these calibration approaches, as well as isotope dilution (Section 3.9), the mathematical models for calculating analyte concentrations and estimating uncertainties are derived, and strategies for optimizing data quality are provided. In the cases of external Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

47

Douglas C. Baxter and Ilia Rodushkin

calibration and standard additions, regression-free alternatives are also included, which may be used when the response of the instrument varies temporally, or with some manipulation, when the number of standards used is limited, e.g. for single point calibration.

3.2

BASIC ASSUMPTIONS AND SOME TERMINOLOGY

It must be emphasized that all calibration approaches are based on certain assumptions, and here it is surmised that there exists a ﬁrst-order, linear relationship between the measured response and the analyte concentration, as illustrated in Fig. 3.1. This may require suitable transformation of one of the variables, e.g., taking the logarithms of the analyte concentrations or activities when employing ion-selective electrodes [2] or the square root of the response when detecting sulfur as excited dimers with a conventional ﬂame photometric detector [3]. Transformations will not be considered further, but an example of linearization is included in Section 3.9, i.e. correction for detector dead time effects. Perhaps, the most important assumption concerns the selectivity of the measurement. Selectivity may be deﬁned as the extent to which the method can be used to determine a particular species in a mixture or a matrix, without spectral (additive) interferences from other components [4]. The occurrence of spectral interferences implies that the method is not completely selective, and thus the detector will respond to one or more concomitant species, as well as the

Fig. 3.1. Idealized calibration curves in the absence (circles, solid line) and presence of spectral (triangles, dotted line) or non-spectral (squares, dashed line) interferents. Note that spectral interferences always cause a positive bias although over-correction for this effect will result in a negative offset. In either case, additive interferences are manifested by a change in the intercept of the calibration curve. A depressive non-spectral interference is illustrated here, but enhancement effects can also be observed. Such multiplicative interferences change the slope of the calibration curve.

48

Calibration approaches for trace element determination

analyte. For a constant spectral interferent concentration, the result will be a positive offset in the calibration curve, as depicted by the dotted line in Fig. 3.1. Non-spectral (multiplicative) interferences, on the other hand, cause changes in the sensitivity, i.e. variations in the response per unit analyte concentration in the presence of matrix components. The dashed line in Fig. 3.1 exempliﬁes a depressive, non-spectral interference. The terms non-spectral and spectral interferences used here are in accordance with the classiﬁcation ramiﬁed by the IUPAC Commission on Spectrochemical and Other Optical Procedures for Analysis in 1978 [5]. In the following, it is implicitly assumed that spectral interferences are absent or have been appropriately dealt with. It must be emphasized that the calibration techniques considered here cannot directly correct for spectral interferences. However, appropriate selection of the standardization approach can eliminate errors originating from non-spectral interferences. The term “errors” will only be utilized to describe situations where the measured quantity systematically deviates from the true value. Random errors will be exclusively denoted as uncertainties, to avoid ambiguities and to conform to current recommendations on terminology [5 –8]. Two further important terms are accuracy and precision that, despite having been deﬁned in IUPAC nomenclature recommendations in 1976 [9], have been frequently misused and confused in the literature. Accuracy refers to the closeness between a measured and expected, consensus or true value. Precision, on the other hand, describes how well repeated measurements of the same quantity in the same sample agree with each other. The major goals of calibration in analytical chemistry are thus to generate results that are both as accurate and as precise as possible.

3.3

SELECTION OF THE CALIBRATION APPROACH

Figure 3.2 summarizes the factors that must be considered in order to make the correct choice of calibration technique. The ﬁrst question to answer concerns whether or not the instrumental response is stable as a function of time, the two possible outcomes leading to different branches of the family of calibration techniques. If the short-term stability of the instrumental response is poor, then it will be necessary to alternate measurements of unknowns and calibrants at regular intervals to facilitate correction for drifting sensitivity. During analytical method development or validation, work will be required to determine whether interference effects are present. This may be achieved by recovery studies, using a range of matrices and analyte concentrations for which the method will be applied [10]. The recovery is the proportion of the amount of analyte, either initially present in or added to the analytical portion of the test material, which is actually measured. Losses of analyte or contamination during the process of sample preparation will obviously cause the recovery to deviate from 100%. (Note that contamination can be considered

49

Douglas C. Baxter and Ilia Rodushkin

Fig. 3.2. Flowchart for the selection of a suitable calibration approach. The question of whether an internal standard can be used depends on whether the instrument is capable of determining several different species or elements in a single run. If it is possible to perform isotope-speciﬁc measurements, then isotope dilution is a viable calibration option, favored for its potential to provide very accurate and precise results.

50

Calibration approaches for trace element determination

as a form of spectral interference [11], requiring additional measurements for its correction, as discussed in Chapter 2.) Further complication arises because concomitant species may depress or enhance the response of the measurement system. Thus, recovery experiments should be designed to facilitate identiﬁcation of the stage at which errors are introduced. Further discussion of recovery is deferred to Section 3.4, although it must be emphasized that this is one of the most important factors in calibration approach selection. If the sensitivity is constant and the recoveries are quantitative, external calibration is the obvious choice. External calibration, also referred to as the calibration curve technique, is the most widely employed approach. The popularity of external calibration is due to the fact that, once calibrated, the measurement system can be used to analyze as many samples as required, at least in principle. In practice, re-calibration may be necessary at appropriate intervals, depending on the temporal stability of the instrumental response. Thus, a crucial prerequisite is that the sensitivity is constant for the duration of the instrument calibration and the analysis of the unknowns. A second prerequisite for employing external calibration is the presumption of a complete lack of interferences; in other words, the sensitivity must be independent of the matrix composition. To ameliorate the stringency of this requirement, some form of matrix matching may be adopted. In certain situations, this may be as straightforward as preparing the unknown test portions and calibrants in the same solvent to ensure a constant sample introduction rate, e.g. for analytical techniques utilizing solution nebulization, such as ICP-OES, ICP-MS or ﬂame AAS [11 – 13]. For potentiometric measurements, the so-called total ionic strength-adjusting buffer is added to sample test portions and standard solutions to ﬁx both the ionic strength and the pH [2]. Whatever the approach chosen, the constancy of the sensitivity can, and should, be conﬁrmed by preliminary method validation work, such as using recovery studies [10]. If recoveries are inadequate, however, then employing internal standardization should be contemplated. Internal standardization provides a means to correct for drift in the sensitivity of the instrument and for non-spectral interferences [14–18]. Instead of using the analyte itself as the calibrant, a chemically distinct species is employed. The internal standard may be a different chemical form (species) of the analyte or another element altogether. In the former case, detection must be preceded by on line separation of the analyte into its native and added forms (e.g. by chromatography, electrophoresis, etc.); in the latter case, the instrument should be capable of detecting both the analyte and the internal standard element simultaneously (or at least in rapid sequence, see Chapter 5). In either case, the internal standard is added at a predetermined concentration, although in favorable circumstances, a species present in all unknowns at a known and preferably constant level may prove suitable. Isotope dilution is considered to represent the ideal variant of internal standardization and involves spiking the unknown with isotopically enriched

51

Douglas C. Baxter and Ilia Rodushkin

analyte [19–24]. As for internal standardization, it is necessary to be able to measure two components in each unknown, i.e. the enriched isotope of the spike and an additional, naturally occurring isotope of the analyte. This requirement obviously dictates the use of analytical instrumentation facilitating isotope speciﬁc measurements. Mass spectrometry is generally the technique of choice, allowing stable isotopes to be employed, and will be treated exclusively here. Like internal standardization, isotope dilution will correct for both drifting sensitivity and non-spectral interferences. The advantage is that the same element is used, avoiding the difﬁculties encountered with internal standardization of ﬁnding chemically distinct species that behave identically with respect to non-spectral interferences [14,16]. Nevertheless, it should be remembered that isotopes of the same element differ with respect to bond energy [25], vapor pressures [26], etc. This is particularly important for lighter elements, which display notable mass effects in their physical and chemical characteristics. However, such effects are of negligible consequence for most applications of isotope dilution. When the sensitivity is constant but neither internal standardization nor isotope dilution is a viable option, the method of standard additions may be adopted. As the name implies, known amounts of standard are added to portions of the sample to derive a calibration function speciﬁc to that sample. The fact that calibration is performed individually on each unknown is both a blessing and a curse. Variable degrees of non-spectral interferences are admirably dealt with, but at the cost of expending much more valuable instrument time per sample. For this reason, applying the method of standard additions is often seen as a last resort. Note that, although correction for recovery is mentioned in two of the calibration approaches included in Fig. 3.2, this operation is not always speciﬁed in descriptions of analytical methods. On the one hand, the accuracy of the reported result is of prime concern in analytical chemistry. On the other hand, some widely employed analytical methods do not provide complete recoveries. If results reported by some, but not all, laboratories were corrected for recovery, the comparability of the data would be compromised and could lead to disputes. Thus, it is important to provide complete documentation of how the results were actually computed [10].

3.4

STATISTICAL EVALUATION OF RECOVERY DATA

Fortunately, the question as to whether the sensitivity is dependent on the matrix can be easily resolved, without controversy, by simple spiking experiments. The recovery, R; is calculated from the difference in the instrumental response to the analytical portion, before ðInative Þ and after ðIobs Þ spiking, and the signal obtained for a calibrant containing the same analyte

52

Calibration approaches for trace element determination

concentration as that added to the spiked material ðIspike Þ: R ¼ ðIobs 2 Inative Þ=Ispike

ð3:1Þ

Ideally, the analytical method will provide recoveries of 100% (i.e. R ¼ 1) for all tested matrices and analyte concentrations. In reality, this is an unlikely situation, due to either the presence of interferences or random variations in the measured responses. This raises the question as to which of these two possibilities actually explain the deviations. To answer this question, some basic statistical concepts will be introduced at this juncture. Here, the discussion will be conﬁned to estimating the uncertainty in the example of Eq. (3.1). The approach can readily be extended to consider all situations of interest. It should be mentioned that a detailed account of evaluating uncertainties associated with recovery experiments has been provided by Barwick and Ellison [27]. If the measurements included in Eq. (3.1) are repeated, such that the uncertainties in the signals can be estimated, it is possible to determine whether the recovery is statistically signiﬁcantly different from unity, i.e. 100%. This requires calculation of the combined standard uncertainty, uc ðRÞ; of the recovery value. First, the uncertainties must be derived for each of the variables, these being termed standard uncertainties if expressed as standard deviations for q replicates: "

q X ðIx;i 2 Ix Þ2 uðIx Þ ¼ q i¼1

#0:5 ,

where the mean is Ix ¼

q X Ix;i q i¼1

ð3:2Þ

Equation (3.2) assumes that the measurements are normally distributed about the mean value, Ix ; which is true for most experimental data providing that q $ 4 [28]. For estimating standard uncertainties for variables following other distributions, reference can be made to a variety of sources [6 –8]. Armed with standard uncertainties for the three variables in Eq. (3.1), the rules for the propagation of random uncertainties can be applied [6 –8,29 –31], enabling an expression for uc ðRÞ to be formulated. This universally applicable approach involves evaluating the partial derivatives of the computed quantity, R in this case, with respect to each of the variables: !2 2 ›R 2 2 ›R ›R 2 uc ðRÞ ¼ u ðIobs Þ þu ðInative Þ þu ðIspike Þ ›Iobs ›Inative ›Ispike 2

0:5

ð3:3Þ

Note that the partial derivative terms may also be denoted sensitivity coefﬁcients, being measures of the extent to which individual uncertainties are ampliﬁed by the relationships of the variables to the calculated

53

Douglas C. Baxter and Ilia Rodushkin

quantity [8,23]. Evaluating the partial derivatives yields: 1

2

uc ðRÞ ¼ u ðIobs Þ

Ispike

!2 2

þu ðInative Þ 2

1 Ispike

!2

I 2I þu ðIspike Þ 2 obs 2 native Ispike

!2

0:5

2

ð3:4Þ The penultimate stage in the process is to calculate the expanded uncertainty, U; by applying an appropriate multiplier to uc ðRÞ: This multiplier is termed the coverage factor, k; with k ¼ 2 being generally recommended as this provides a conﬁdence level of approximately 95%. However, should uc ðRÞ be associated with less than six effective degrees of freedom, n, then k should be equated with the corresponding two-tailed value of Student’s t-statistic at the desired conﬁdence level (generally 95%). One degree of freedom is lost for each parameter, p; calculated from the data set. For example, when the mean was calculated in Eq. (3.2), one degree of freedom was lost, n ¼ ðq 2 pÞ ¼ ðq 2 1Þ: The partial derivative terms are also instrumental in approximating the effective degrees of freedom for the combined standard uncertainty, using Satterthwaite’s formula [28,32]. Again using Eq. (3.1) as an example, nðRÞ is calculated as: 8 2 2 32 2 3 2 > ›R 2 ›R > 2 2 > u ðI Þ u ðI Þ obs native < 1 6 1 6 ›Iobs 7 ›Inative 7 7 7 6 6 nðRÞ ¼ 7 þ 7 6 6 2 2 5 5 4 4 > nIobs nInative u ðRÞ u ðRÞ > c c > : 2

›R 6 u ðIspike Þ ›Ispike 1 6 6 þ 6 nIspike 6 u2c ðRÞ 4 2

21 !2 32 9 > > > > 7 > 7 = 7 7 7 > 5 > > > > ;

ð3:5Þ

the result being rounded down to the nearest integer. Summarizing the aforementioned considerations, the recovery and its expanded uncertainty can be expressed as: R ^ U ¼ R ^ kuc ðR

3:6Þ

or for the more general case: x ^ U ¼ x ^ kuc ðx

3:7Þ

where k ¼ 2 for an approximately 95% conﬁdence level and n . 6; or k ¼ again assuming a 95% conﬁdence level. If the inequality ðR 2 UÞ , R , ðR þ UÞ encompasses the value of one, then it may ﬁnally be concluded that the recovery is not statistically different from 100%. Thus, any deviations from 100% recovery in the data set can be

tna¼0:05,

54

Calibration approaches for trace element determination

attributed to random variations rather than to non-spectral interferences. If, on the other hand, this inequality does not contain the value of 1, then the possibility that the instrumental response is subject to non-spectral interferences cannot be excluded and therefore it can be concluded that the matrix alters the sensitivity. Although this might appear discouraging, it will certainly provide a rational basis on which to select the means to calibrate. However, before embarking on any further exposition of calibration, the most relevant variants of linear regression analysis will be introduced, as these constitute the foundation for several of the approaches to be discussed. It is important to note that, in the following, the term standard deviation of the mean will be required, a quantity that is often referred to as the standard error or the standard error of the mean [8] and is symbolized here by sðxÞ: If a quantity is determined repeatedly such that there are m estimates of the mean and its standard uncertainty, x ^ uðxÞ; then: " #0:5 m m X X ðxi 2 x Þ2 x i sðxÞ ¼ where x ¼ ð3:8Þ m m i¼1 i¼1 and x is the mean of means. There is an exact relationship between the standard deviation of the mean and the standard uncertainty, i.e., uðxÞ sðxÞ ¼ pﬃﬃﬃ m

ð3:9Þ

showing that, as m increases, the spread of the means decreases. Note that calculation of the expanded uncertainty in x is given by: x ^ U ¼ x ^ ksðx

3:10Þ

subject to the same conditions as those noted directly following Eq. (3.7), pﬃﬃﬃ despite the inclusion of the additional divisor m [8]. 3.5

LINEAR REGRESSION

Before embarking on our discussion of calibration approaches, we will attempt to review the linear regression techniques that may be applied to situations of the type illustrated in Fig. 3.1. For ﬁtting straight lines to experimental data, ordinary (or unweighted) linear regression (OLR), as described in Section 3.5.1, is undoubtedly the most popular statistical technique. This is hardly surprising, given that it is the only form of regression analysis included in general analytical textbooks. Application of OLR requires that several assumptions are fulﬁlled, the two most important being that (a) the variation in measured response is constant, irrespective of concentration and (b) the concentrations of analyte in the calibrants are known exactly, with negligible error. The former prerequisite can be readily assessed by examining a residuals plot (see below and Fig. 3.3), which necessitates replicate measurements at each

55

Douglas C. Baxter and Ilia Rodushkin

Fig. 3.3. Illustration of the application of OLR to measured data. The ﬁtted line in (a) has the equation I^i ¼ ð0:010 ^ 0:026Þ þ ð0:920 ^ 0:043ÞCi with r ¼ 0:9993: Uncertainties are 95% conﬁdence limits. A residuals plot is shown in (b), clearly indicating curvature that cannot be modeled by a ﬁrst-order linear regression equation.

concentration [33]. If condition (a) is not met, then weighted linear regression should be used (Section 3.5.2). Linear regression for situations where there are uncertainties along both variable axes is considered in Section 3.5.3. 3.5.1

Ordinary linear regression

The estimation procedure for computing the OLR equation of the line relating measured responses, I, to concentrations, C; Ii ¼ ao þ bo Ci þ 1i

ð3:11Þ

is that of least squares [34]. Here the subscript “o” is used to indicate that OLR is being used to estimate the parameters describing the relationship between I and C: Obviously, in a calibration experiment, the magnitude of the response is dependent on the concentration. Therefore, the terms dependent and independent variables can be applied to I and C; respectively. In Eq. (3.11), ao and bo are the regression coefﬁcients obtained by minimizing the sum of squares (SS) of the deviations ð1i Þ from the line, i.e., SSo ¼

n X i¼1

12i ¼

n X

ðIi 2 ao 2 bo Ci Þ2

ð3:12Þ

i¼1

These deviations, reﬂecting the distances of the measured data points from the ﬁtted equation at Ci ; are usually referred to as residuals. The regression coefﬁcients, ao and bo ; represent the intercept of the line with the y-axis (with the same units as the response) and the slope (units of response per unit analyte concentration, which equates with the sensitivity), respectively. Differentiating Eq. (3.12) with respect to ao and then to bo ; and setting

56

Calibration approaches for trace element determination

the results equal to zero, yields: X X Ii 2 nao 2 bo Ci ¼ 0

ð3:13aÞ

X X X Ci Ii 2 ao Ci 2 bo C2i ¼ 0

ð3:13bÞ

where all summations are from i ¼ 1 to n: These are called the normal equations, with the solution: ao ¼ I 2 bo C P P P P Ci Ii 2 Ci Ii =n ðCi 2 C ÞðIi 2 IÞ ¼ bo ¼ P P 2 P 2 2 ðCi 2 CÞ Ci 2 Ci =n

ð3:14aÞ ð3:14bÞ

Additional statistics are needed for the purposes of assessing the quality of the ﬁtted regression line and estimating the uncertainty in the determined analyte concentration. The most important of these is actually that given by Eq. (3.12), divided by n ¼ n 2 2: P P ðIi 2 ao 2 bo Ci Þ2 ðIi 2 I^i Þ2 S2o ¼ ¼ ð3:15Þ n22 n22 where two degrees of freedom have been lost through calculation of two regression coefﬁcients, ao and bo ; from the data. S2o is called the mean square about regression, elsewhere symbolized in various other ways, such as s2yx [33] or s2y=x [35], and S the residual standard deviation. I^i denotes the predicted or ﬁtted response calculated from ao ; bo and Ci : Knowing S2o ; the standard deviations of the mean values of the estimated slope and intercept, as well as the covariance between ao and bo ; can be calculated, i.e., " #0:5 P 2 Ci sðao Þ ¼ So ð3:16aÞ P 2 n ðCi 2 CÞ " #0:5 S2o sðbo Þ ¼ P ð3:16bÞ ðCi 2 C Þ2 S2 C ð3:16cÞ covðao ; bo Þ ¼ 2 P o 2 ðCi 2 CÞ It is worth noting that the standard deviations of the mean values of the intercept and slope are equivalent to their standard uncertainties [33]. Frequently, the correlation coefﬁcient, r, is used as a measure of the quality of the regression: "P P #0:5 ðCi 2 C ÞðIi 2 IÞ ðCi 2 C Þ2 r ¼ P ¼ bo P ð3:17Þ P 2 2 0:5 ðIi 2 IÞ ðC 2 C Þ2 ðI 2 IÞ i

i

57

Douglas C. Baxter and Ilia Rodushkin

As 21 , r , 1; the correlation coefﬁcient is, in fact, a scaled version of the estimated slope [34], as is clearly demonstrated by Eq. (3.17), implying only that there is a dependence between C and I: Were this not true, then the instrument could not be used for calibration purposes at all. Most important is the fact that a value for the correlation coefﬁcient close to 1 does not actually prove that the calibration is linear over the concentration region considered [33,35,36]. An example of this problem is shown in Fig. 3.3. Despite a correlation coefﬁcient of 0.9993, the ﬁrst-order linear model (Fig. 3.3a) is clearly demonstrated to be inadequate by the appearance of the associated residuals plot of Fig. 3.3b. The latter plot nevertheless indicates that the use of OLR is appropriate, since the range of residuals is independent of concentration, i.e. 1i ðmaximumÞ 2 1i ðminimumÞ < constant: A further possibility to check for curvature is provided by examining the conﬁdence interval for the intercept. Conﬁdence limits for both model parameters, ao and bo ; are given by: ao ^ tsðao

3:18aÞ

bo ^ tsðbo

3:18bÞ

where t is the Student’s t-value for a two-tailed distribution at the selected conﬁdence level and with n ¼ n 2 2: If n . 6; t can be replaced by k ¼ 2 for an approximately 95% conﬁdence interval, as mentioned previously (Section 3.4). If the conﬁdence interval for ao includes the value of the measured response at zero concentration, then there is no evidence for curvature. Unfortunately, this test is not particularly robust, as OLR tends to yield rather broad conﬁdence intervals for the intercept (as shown in the example described in Fig. 3.3, where the response at Ci ¼ 0 was zero and thus within the conﬁdence interval for ao ). Therefore, residuals plots should always be constructed to test for limitations of the ﬁtted equation. An additional beneﬁt is that outlying points may reveal themselves, as suggested by the results for a calibration graph containing a deviating data point (Fig. 3.4a). The corresponding residuals plot (Fig. 3.4b) identiﬁes the responses at the central calibration point as outliers, because they are not randomly distributed about the 1i ¼ 0 line. The ability to discern outliers is highly dependent on the number of concentration levels and the measurement precision, and the use of robust regression techniques may be preferable to ascertain whether outlying data are present [37,38]. If no evidence for curvature is detected, and if the conﬁdence interval for ao includes zero (assuming, of course, that the response at C ¼ 0 really is zero), the data can be better ﬁtted using an alternative, simpler version of Eq. (3.11): Ii ¼ b0o Ci þ 1i

ð3:19Þ

This results from the fact that the intercept term is not signiﬁcantly different from zero and can thus be eliminated. Equations (3.12)– (3.16) can be rewritten

58

Calibration approaches for trace element determination

Fig. 3.4. Effect of an outlying data point (at C ¼ 0:5) on OLR. The ﬁtted line in (a) is described by the equation I^i ¼ ð0:010 ^ 0:063Þ þ ð1:00 ^ 0:10ÞCi with r ¼ 0:997; with no sign of any outlier. The residuals plot (b) is, however, efﬁcient at revealing the deviating behavior of this point.

as: SS0o ¼

n X i¼1

12i ¼

n X

ðIi 2 b0o Ci Þ2

ð3:20Þ

i¼1

X X Ci Ii 2 b0o C2i ¼ 0 P CI b0o ¼ P i 2 i Ci P P ðIi 2 b0o Ci Þ2 ðIi 2 I^i Þ2 ¼ S02 o ¼ n21 n21 hX i 2 0:5 0 0 sðbo Þ ¼ So = ðCi 2 C Þ

ð3:21Þ ð3:22Þ ð3:23Þ ð3:24Þ

with the prime indicating that the relevant term applies to the zero intercept, one parameter OLR model. Apart from computational simplicity, this model offers the advantages of using up only one degree of freedom and avoiding the need to account for covariance between regression parameters, as will become apparent below (Sections 3.6 –3.8). This can lead to considerable reductions in the magnitudes of the uncertainties associated with predicted concentrations. Figure 3.5 is an example of a spreadsheet that could be used for performing the calculations required by OLR. Harris [18] has also presented a similar spreadsheet solution for the case of OLR with a non-zero intercept term. 3.5.2

Weighted linear regression

The choice between ordinary and weighted linear regression (WLR) methods falls on the uncertainty structure. In the residuals plot of Fig. 3.6a, all values fall within a region bounded by two parallel lines, symmetrically displaced from

59

60 Douglas C. Baxter and Ilia Rodushkin Fig. 3.5. Example of a spreadsheet for OLR analysis of calibration data. Note that the ﬁgures in bold in row 10 are all the sums of data contained in the cells above. Also included are data for ﬁve “unknowns”, and the analytical results generated by external calibration (see Section 3.6) using OLR models with and without the intercept, Eqs. (3.11) and (3.19), respectively. Uncertainties in the response values for the unknowns were arbitrarily kept constant at all signal levels.

Calibration approaches for trace element determination

Fig. 3.6. The residuals plots depicted in (a) and (b) are for cases where ﬁrst-order models are adequate, i.e., neither outliers nor curvature is present. However, in (b) the range of the residuals is obviously a function of concentration, indicating that application of WLR is necessary.

the line deﬁned by 1i ¼ 0: This is characteristic of a constant level of uncertainty in the measurements; in other words, uðIÞ is independent of response and hence concentration, and the uncertainty is said to exhibit homoscedasticity. For such cases, OLR is the method of choice. In the case of Fig. 3.6b, the occurrence of heteroscedasticity is evident, i.e., the uncertainty of measurement is clearly a function of concentration and modeling the data will beneﬁt from the use of WLR. The need for WLR follows quite logically from a simple analysis of the major noise contributions common to all instrumental methods of analysis [39]. In the simplest cases, the uncertainty may be dominated by white noise, uðIÞ < ðIÞ0:5 ; ﬂicker noise, uðIÞ / I or a combination of both. Irrespective of which, response variations increase with signal level. This leads to the conclusion that the use of OLR will only be valid in a few, exceptional circumstances. Despite the fact that the beneﬁts of WLR have been advocated quite frequently over the last quarter of a century or so [33,35,40–44], its use in current analytical practice would appear to have remained rather limited. This is perhaps due to the perceived complexity of the calculations involved. Actually, implementation of WLR is relatively straightforward using modern spreadsheet software (Fig. 3.7). Previously highlighted problems concerning the computational intensity and introduction of rounding errors [42] are no longer limitations. Recently, a software package facilitating OLR and WLR analyses of calibration data was made available on the World Wide Web [44], and hopefully this initiative will stimulate further use of the technique. The underlying idea behind WLR is that the ﬁtting procedure should be performed in such a way as to ensure that the calculated line passes, as close as possible, the points with the smallest uncertainties. In OLR, on the other hand, the derived line best ﬁts the points with largest responses. In order to force

61

62 Douglas C. Baxter and Ilia Rodushkin Fig. 3.7. Example of a spreadsheet for WLR analysis of calibration data. Note that the ﬁgures in bold in row 10 are all the sums of data contained in the cells above. Also included are data for ﬁve “unknowns”, and the analytical results generated by external calibration (see Section 3.6) using WLR models with and without the intercept, Eqs. (3.27) and (3.28), respectively. Uncertainties in the response values for the unknowns were arbitrarily kept constant at all signal levels. The uncertainties in the response data for the standards were modeled as uðIi Þ ¼ 2 þ 0:02 £ Ii : In some versions of Excel, the arguments to the IF statement should be separated by semi-colons instead of commas.

Calibration approaches for trace element determination

the line through the most precisely measured points, generally those at lower concentrations, all data are weighted, typically using: wi ¼ u22 ðIi

3:25Þ

It should be mentioned that using experimentally measured response uncertainties to calculate weights has its limitations. For example, in the admittedly unlikely event that uðIi Þ ¼ 0 at some response level, the weight at that point would be inﬁnite. Uncharacteristically large, or small, uncertainties at any given response will result in an abnormal distribution of weights along the regression line. To avoid potential problems in the weighting process, the uncertainties can be modeled as a function of response level [42,43], using a polynomial of the form: u2 ðIi Þ ¼ f þ gIi þ hIi2 þ 1i

ð3:26Þ

The regression parameters may be considered to represent the variance at zero response ðf Þ; and the coefﬁcients of white ðgÞ or ﬂicker ðhÞ noise contribution to signal variability. A comparison of WLR using experimental and modeled uncertainties for ICP-OES calibration indicated that there was little to choose between them [43]. Modeling the uncertainty has a smoothing effect that will ﬁlter out noise to some extent, potentially resulting in artiﬁcially improved precision in the regression parameter estimates. After careful deliberation, we have decided not to pursue the use of modeled uncertainties any further. Depending on whether or not the intercept is signiﬁcantly different from zero, as can be tested as described above, the linear models are: Ii ¼ aw þ bw Ci þ 1i Ii ¼

b0w Ci

ð3:27Þ

þ 1i

ð3:28Þ

where the subscript “w” indicates weighted regression coefﬁcients and the prime denotes the use of the zero intercept assumption. The weights are ﬁrst introduced in the expressions describing the sums of squares to be minimized, with respect to the weighted residuals: SSw ¼

n X

wi 12i ¼

i¼1

SS0w ¼

n X i¼1

n X

wi ðIi 2 aw 2 bi Ci Þ2

ð3:29Þ

wi ðIi 2 b0 Ci Þ2

ð3:30Þ

i¼1

wi 12i ¼

n X i¼1

Differentiating with respect to aw ; if appropriate, and to the slope, the normal equations are obtained: X X X wi Ii 2 aw wi 2 bw wi Ci ¼ 0 ð3:31aÞ X X X 2 wi Ci Ii 2 aw wi Ci 2 bw wi Ci ¼ 0 ð3:31bÞ

63

Douglas C. Baxter and Ilia Rodushkin

X X wi Ci Ii 2 b0w wi C2i ¼ 0 with the solutions: X X X X w ¼ wi Ii = wi 2 bw wi Ci = wi aw ¼ Iw 2 bw C P P P P P wi ðCi 2 C w ÞðIi 2 Iw Þ wi Ci Ii 2 wi Ci wi Ii = wi bw ¼ ¼ P P P P wi ðCi 2 C w Þ2 wi C2i 2 wi Ci 2 = wi

ð3:32Þ

ð3:33aÞ ð3:33bÞ

and

P wCI b0w ¼ P i i 2 i wi Ci

ð3:34Þ

The necessary statistics for describing the uncertainties in the WLR models of Eqs. (3.27) and (3.28) are given by: P P wi ðIi 2 aw 2 bw Ci Þ2 wi ðIi 2 I^i Þ2 ¼ ð3:35Þ S2w ¼ n22 n22 P P 0 wi ðIi 2 b0w Ci Þ2 wi ðIi 2 I^i Þ2 ¼ ð3:36Þ Sw2 ¼ n21 n21 " #0:5 " #0:5 P P P wi C2i wi C2i = wi ¼ S ð3:37aÞ sðaw Þ ¼ Sw P P P P P w wi wi ðCi 2 C w Þ2 wi C2i 2 wi Ci 2 = wi Sw Sw sðbw Þ ¼ P P P i0:5 2 0:5 ¼ hP 2 wi ðCi 2 C w Þ wi Ci 2 wi Ci 2 = wi w S2w C w S2w C covðaw ; bw Þ ¼ 2 P P P 2 ¼2P wi ðCi 2 C w Þ wi C2i 2 wi Ci 2 = wi hX i0:5 wi ðCi 2 C w Þ2 sðb0w Þ ¼ S0w =

ð3:37bÞ

ð3:37cÞ ð3:38Þ

Figure 3.7 displays an example of a spreadsheet that could be used to perform WLR of calibration data, based on the models with and without an intercept described above. 3.5.3

Linear regression for data with uncertainties in both variables

The third important variant of linear regression is that used for data sets where both sets of variables are subject to uncertainties. This is probably most applicable when concentrations, covering a wide range, are determined using two independent analytical methods and the results are to be compared. As such, each data point will consist of two estimated concentrations and their associated uncertainties. It is common practice to assign the results determined using a standard or reference method to the x-axis, implying that

64

Calibration approaches for trace element determination

the concentrations so obtained have negligible uncertainties. This leads to the unintentional inference that the new method must exhibit much greater uncertainties, hardly a desirable conclusion. Another possibility is that the calibrants employed are reference materials, containing experimentally derived concentrations of one or more analytes. Such materials obviously have uncertainties associated with the speciﬁed measurands, which in many situations cannot be justiﬁably neglected, and therefore one of the basic assumptions of OLR and WLR is violated. In either case, it is necessary to account for uncertainties in both the x- and y-directions, and weight the data accordingly. MacTaggart and Farwell [45] have critically evaluated the pertinent methods available in the literature and, on the basis of their ﬁndings, only the most robust version will be considered further, comprising a ﬁrst-order linear model with an intercept: Ii ¼ aW þ bW þ 1i

ð3:39Þ

where the subscript “W” indicates that weighting is now applied to both the y- and x-variables. (The use of responses, I; and concentrations, C; as variables is only for reasons of consistency. For method comparison studies, the responses should be replaced by concentration estimates from the second method.) The weighted residuals sum of squares are given by: SSW ¼

n X i¼1

Wi 12i ¼

n X

Wi ðIi 2 aW 2 bW Ci Þ2

ð3:40Þ

i¼1

where Wi ¼ ½u2 ðIi Þ þ b2W u2 ðCi Þ21

ð3:41Þ

Unfortunately, as shown by Eq. (3.41), the weighting factors are dependent on the slope, and therefore, Wi and bW must either be determined iteratively or additional restraints placed on the weights, such as a constant uncertainty ratio, u2 ðIi Þ=u2 ðCi Þ: For the iterative solution, attributable to Williamson [46], the slope and intercept are estimated as: X X X X W ¼ Wi Ii = Wi 2 bW Wi Ci = Wi ð3:42aÞ aW ¼ IW 2 bW C P WzV ð3:42bÞ bW ¼ P i i i Wi zi Ui where

X X W ¼ Ci 2 Wi Ci = Wi Ui ¼ Ci 2 C X X Vi ¼ Ii 2 IW ¼ Ii 2 Wi Ii = Wi zi ¼ Wi ½u2 ðIi ÞUi þ bW u2 ðCi ÞVi

ð3:43aÞ ð3:43bÞ 3:43cÞ

Standard uncertainties for the slope, as well as the intercept and its

65

66 Douglas C. Baxter and Ilia Rodushkin Fig. 3.8. Example of a spreadsheet for linear regression analysis of calibration data with uncertainties in both variables. Note that this spreadsheet is an extension of that shown in Fig. 3.5, requiring an initial estimate of the slope as provided by OLR. Uncertainties in the response values for the unknowns were arbitrarily kept constant at all signal levels. The uncertainties in the response data for the standards were modeled as uðIi Þ ¼ 2 þ 0:02 £ Ii ; whereas those for the concentrations were random numbers generated by the spreadsheet software. To iterate, cells (O2:Y19) should be copied and pasted into cells (AA2:AK19), (AM2:AW19), etc. The results of the fourth and ﬁfth iterations are identical, those for the latter being included. In some versions of Excel, the arguments to the IF statement should be separated by semi-colons instead of commas.

Calibration approaches for trace element determination

covariance, can also be calculated: X sðaW Þ ¼ 1= Wi þ 2ðC W þ 2zÞzQ þ ðC W þ 2zÞ2 s2 ðbW Þ

0:5

nX o0:5 Wi2 ½u2 ðIi ÞUi2 þ u2 ðCi ÞVi2 sðbW Þ ¼ Q W s2 ðbW covðaW ; bW Þ ¼ 2C

ð3:44aÞ ð3:44bÞ 3:44cÞ

where nX o21 Q¼ Wi ½Ui Vi =bW þ 4z0i ðzi 2 Ui Þ X X z ¼ Wi zi = Wi

ð3:45aÞ

z0i

ð3:45cÞ

¼ zi 2 z

ð3:45bÞ

Solution of Eq. (3.42) requires an initial estimate of the slope, which can most simply be provided by OLR analysis of the data. Although these equations appear rather intimidating, they can be readily solved using commercial spreadsheet programs, as exempliﬁed in Fig. 3.8. MacTaggart and Farwell [45] have conﬁrmed that the slope estimate converges rapidly, generally no more than 20 iterations sufﬁcing to reach a stable value. It should be noted that this model provides an estimate of the slope that is equal to the reciprocal of that obtained upon switching axes. This is particularly important when using linear regression to compare data obtained using two independent methods applied to the same set of samples. In that case, the conﬁdence intervals for the slope and the intercept should contain the expected values for perfect agreement of 1 and 0, respectively. OLR will yield two different sets of values that may provide contradictory conclusions regarding the equivalency of the two methods, clearly an inconsistent and unsatisfactory result. Another attractive feature of Williamson’s [46] approach is that sðbW Þ=bW remains constant on switching axes, which is further proof of the models’ quality.

3.6

EXTERNAL CALIBRATION

To use linear regression, a sufﬁcient number, n; of calibration points must be measured; this requires that n $ 3 or 2 for the models with and without the intercept, respectively. The concentration of analyte in the unknown sample, Cu ; is then given by inserting the measured response, Iu ; in the appropriate variant of the regression equation, Eq. (3.11), (3.19), (3.27), (3.28) or (3.39), and solving for Cu (or C0u ): Cu ¼ ðIu 2 aÞ=b C0u ¼ Iu =b0

where a [ ðao ; aw ; aW Þ and b [ ðbo ; bw ; bW Þ

where b0 [ ðb0o ; b0w ; b0W

ð3:46Þ 3:47Þ

67

Douglas C. Baxter and Ilia Rodushkin

As far as we are aware, linear regression with uncertainties in both variables has never been utilized for calibration purposes, but then again, it has not found much application at all to date. 3.6.1

Estimating uncertainty

The next step is to calculate the combined uncertainty in the estimated concentration sc ðCu Þ or sc ðC0u Þ with reference to Eqs. (3.46) and (3.47), respectively. (Perhaps, the term “combined standard deviation of the mean estimated concentration” would be more consistent with the quantity actually being calculated, although such terminology is not used in the current literature [8].) This includes uncertainties in the measured response and the slope of the calibration line, and, where applicable, the intercepts’ uncertainty and covariance with the slope. The addition of a covariance term accounts for the fact that the estimates of a and b are not independent (unless the data are ﬁrst mean-centered [28]), both being computed from the same calibration data set: sc ðCu Þ ¼ s2 ðIu Þ " sc ðC0u Þ ¼ s2 ðIu Þ

›Cu ›Iu ›Cu ›Iu

2

2

þs2a þs2b

›Cu ›a ›Cu ›b

2

þs2b

›Cu ›b

2 þ2 covða; bÞ

›Cu ›Cu ›a ›b

0:5

ð3:48Þ

2 #0:5 ð3:49Þ

Evaluating the partial derivatives and simplifying yields: sc ðCu Þ ¼

1 2 ½s ðIu Þ þ s2a þ s2b C2u þ 2 covða; bÞCu 0:5 b

ð3:50Þ

sc ðC0u Þ ¼

0 1 2 ½s ðIu Þ þ s2b0 Cu2 0:5 b0

ð3:51Þ

showing how the combined uncertainty in the estimated concentration is related to the parameters described in previous sections. Note that, when using Eqs. (3.50) and (3.51), s2 ðIu Þ ¼ u2 ðIu Þ=m; where m is the number of independently prepared and analyzed unknowns, not the number of replicate measurements of the same test portion of a single sample. The standard uncertainty of replicate measurements of a single unknown, i.e. uðIu Þ; should only be used in the situation that no other estimate of the variation is available, in which case m ¼ 1: This point is further developed at the end of Section 3.6.2. As Eqs. (3.50) and (3.51) are applicable to all types of linear regression with uncertainties in both variables, they are certainly the most ﬂexible forms of the uncertainty expressions for estimated concentrations. However, valuable insight into the uncertainty structure, for the cases of OLR and WLR, can be

68

Calibration approaches for trace element determination

gained by some manipulation of these equations, i.e., " #)0:5 ( 2 1 u2 ðIu Þ ðIu 2 IÞ 2 1 sc ðCu Þo ¼ þ So þ 2P bo m n bo ðCi 2 C Þ2 sc ðC 0u Þo

1 ¼ 0 bo

(

" #)0:5 u2 ðIu Þ Iu2 02 þ So 0 P m bo2 ðCi 2 C Þ2 " # u2 ðIu Þ 1 ðIu 2 Iw Þ2 þ Sw P þ 2P m wi bw wi ðCi 2 C w Þ2

1 sc ðCu Þw ¼ bw sc ðC 0u Þw

1 ¼ 0 bw

ð3:52Þ

(

" #)0:5 u2 ðIu Þ Iu2 0 þ S w 02 P m bw wi ðCi 2 C w Þ2

ð3:53Þ 0:5

ð3:54Þ

ð3:55Þ

Unfortunately, due to the complexity of the equations, little insight is gained by similar manipulation of Eqs. (3.50) and (3.51) for linear regression with uncertainties in both variables. Thus, we will conﬁne ourselves to the OLR and WLR cases when considering how to optimize precision for the analyte concentration estimates in Section 3.6.2. The same strategies are, of course, also applicable to data sets with uncertainties along both axes. Calculation of the expanded uncertainty for the estimated analyte concentration in a single unknown is performed on the basis of Eq. (3.10). Here, the approximate nature of the resultant “conﬁdence interval” should be recognized. In reality, the upper and lower conﬁdence limits for a concentration estimated using any of the linear regression techniques will be asymmetrically distributed about the mean value [33,34,41]. Use of Eq. (3.10), on the other hand, is simple and in line with current recommendations [6,8]. Clearly, application of Eq. (3.10) requires knowledge of the number of degrees of freedom, simply given by [8,33 –35]: n¼n22

ð3:56Þ

or n¼n21

ð3:57Þ

for calibration lines calculated with and without the intercept, respectively. However, once the calibration equation has been calculated, it will normally be used to estimate the analyte concentrations in a larger number of unknowns. As the same regression parameters will be used for all unknowns, Eq. (3.46) or (3.47), the resulting concentration estimates will all be correlated. This particular complication is rarely addressed in the analytical literature, notable exceptions including the work of Kragten [28], MacTaggart and Farwell [33] and Watters et al. [43]. Adequately accounting for this additional uncertainty remains a source of dissatisfaction for statisticians. A simpliﬁed approach is to

69

Douglas C. Baxter and Ilia Rodushkin

make the following substitution in Eq. (3.10): u k ¼ ta=2n n

ð3:58Þ

where a is the desired probability level (e.g. a ¼ 0:05 for a 95% conﬁdence level), nu is the number of unknowns that will be calibrated and n is the appropriate value from Eq. (3.56) or (3.57). The two-tailed t-statistic demanded by Eq. (3.58) can be readily calculated using modern spreadsheet software. More details are provided by MacTaggart and Farwell [33] and the literature cited therein. It must be emphasized at this juncture that multiple uses of calibration lines is a topic not considered in current ofﬁcial publications discussing the treatment of uncertainties in analytical measurements. Therefore, it is probably advisable to ignore this complication at the present time. Returning to the formulae for uncertainty calculations, it is worth noting that the ﬁrst term within the brackets in Eq. (3.52) is often represented as S2o =m: The implication is then that repeated analyses of the unknown will yield equivalent precision to that obtained for any given standard. (To be more speciﬁc, sðIu Þ must be insigniﬁcantly different from So in Eq. (3.52) or S0o in Eq. (3.53). The validity of this assumption can be tested by calculation of the F-statistic: Fn1 ;n2 ¼ s2 ðIu Þ=S2o F 0n1 ;n2

¼s

2

0

ðIu Þ=So2

where n1 ¼ ðm 2 1Þ; n2 ¼ ðn 2 2Þ

ð3:59aÞ

where n1 ¼ ðm 2 1Þ; n2 ¼ ðn 2 1Þ

ð3:59bÞ

If the calculated F-statistic is greater than the tabulated critical value at the selected conﬁdence level, then the use of this alternative form is unjustiﬁed and 0 Eq. (3.52) or (3.53) should be used as written. For OLR, S2o or So2 is a pooled estimate of the spreads of responses over the range of concentrations [33]. 3.6.2

Optimizing precision

Considering OLR with a non-zero intercept, Eq. (3.52) suggests that there are several possibilities to minimize the uncertainty in the estimated concentration. In the vast majority of cases, the ﬁrst term on the right-hand side of each of the Eqs. (3.52) –(3.55) is the major contributor to the uncertainty in the estimated concentration, as m will probably be close to one. Thus, increasing m; or indeed in Eq. (3.52); n; will have the desired effect, as will decreasing the term ðIu 2 IÞ obviously, as the measured response for the unknown approaches the mean, the third term in the brackets in Eq. (3.52) tends toward zero. Consequently, the importance of this term increases toward both extremes of the calibration line, as exempliﬁed by the plot of the expanded (and scaled) uncertainty of the concentration estimate versus unknown response in Fig. 3.9a. For this reason, unknown samples should be prepared in such a way as to ensure that the IÞ; of the calibration line as measured responses are as close to the centroid, ðC; possible. Increasing the sum in the denominator can also reduce the contribution of the third term. One approach is to extend the concentration

70

Calibration approaches for trace element determination

Fig. 3.9. Effect of the type of external calibration model, (a) OLR or (b) WLR, on concentration estimates, Cu (solid lines), and expanded scaled uncertainties as a function of response for unknown. The dashed and dotted lines bound the expanded uncertainty bands for calibration lines with and without an intercept term, respectively. The data for OLR and WLR are those included in Figs. 3.5 and 3.7, respectively. As n ¼ 5 ðn , 6Þ; we ¼ 3:18 or 2.78 for the models with and without an intercept, respectively, used k ¼ ta¼0:05 n rather than k ¼ 2 (Section 3.4). To emphasize the differences between models, the expanded uncertainty terms were also scaled by a factor of 5.

range covered by the calibration line. The other is to prepare standards with concentrations focused at the extremes of, rather than evenly distributed over, the calibration range [47]. However, care should be exercised to avoid violating the assumption of a ﬁrst-order linear relationship between response and concentration when attempting to apply the latter approaches. Similar considerations are also applicable to the use of WLR with a non-zero intercept, Eq. (3.54). For the latter, Fig. 3.9b shows that the expanded (and scaled) uncertainty of the concentration estimate is a generally increasing function of the unknown response. The most striking feature in comparing panels (a) and (b) in Fig. 3.9 is that the use of weighting results in smaller absolute uncertainties at lower concentrations. From Fig. 3.9b it is apparent that the uncertainties of the concentration estimates are proportional to the responses; this is consistent with the fact that most analytical instruments provide readings with larger absolute uncertainties at higher concentrations. Generally, WLR with a non-zero intercept will not provide a pronounced w ; Iw Þ: This results from the minimum in uncertainty close to the centroid, ðC fact that the ﬁrst term on the right-hand side of Eq. (3.54) increases monotonically with response, whereas the corresponding term in Eq. (3.52) should be constant, i.e., consistent with the homoscedasticity assumption. Examples of expanded (and scaled) uncertainty plots derived on the basis of zero intercept OLR and WLR are included in Fig. 3.9. The most obvious effect of eliminating the intercept is that of providing smaller uncertainties in estimated concentrations at lower responses, close to the origin, for OLR. This is as expected from Eq. (3.53), because the ﬁrst and second terms on

71

Douglas C. Baxter and Ilia Rodushkin

the right-hand side are constant and quadratic in the response, respectively. The latter term tends to zero on approaching the origin, whereas the corresponding term in Eq. (3.52) is minimized at the centroid. For the zero intercept WLR model, Fig. 3.9b clearly indicates the advantage gained, compared to the variant including the intercept, in terms of the smaller uncertainties in estimated concentrations over almost the entire calibration range. Obviously, one of the key considerations is the number of standards to use in constructing the calibration function. Preparing a large number of calibrants is tedious, particularly for the multi-elemental measurements that are becoming the status quo in analytical chemistry, and demands valuable instrument time for their analysis. It might, therefore, be tempting to include all replicate measurements in the OLR analysis of the calibration data. There is a precedent for this provided in one of the examples included in the current EURACHEM/CITAC guide to the evaluation of measurement uncertainty [8]. Furthermore, this practice is both pragmatic and logically defensible because standards are usually prepared from solutions; thus, replicate measurements of a single standard should yield an uncertainty that is statistically indistinguishable from that for single measurements of several, independently prepared calibrants of the same nominal concentration. However, the same argument is not applicable to most sample types. Substantial sample preparation may precede analysis, the material under study may be inhomogeneous with respect to analyte distributions, and corrections for spectral interferences may be required. Thus, replicated measurements of a single test portion will often yield an overly optimistic estimate of the uncertainties involved. On the other hand, the costs of preparing series of independent replicate test portions from every laboratory sample are likely to be prohibitive. During method validation, the analytical method should be thoroughly evaluated using the range of matrices and concentrations to which it will be applied [10], as noted previously in Section 3.3. At this stage, sufﬁcient data should be collected so that a reasonable estimate can be made of the expected reproducibility for any future sample. Such data can then be used to provide estimates of response uncertainties for Eqs. (3.50) –(3.55). Unless several test portions of the same laboratory sample are prepared for analysis and subjected to the measurement process, the divisor in the ﬁrst term of Eqs. (3.52)– (3.55) must be set to one, irrespective of the regression technique being used. 3.6.3

Accounting for non-constant sensitivity

If the sensitivity were not constant during the period over which unknowns are analyzed, external calibration would have to be repeated at appropriate intervals. In practice, this is likely to be achieved by measurement of a single standard to correct for changes in the slope of the calibration function. This is,

72

Calibration approaches for trace element determination

statistically speaking, an example of a single point calibration for which linear regression is unsuitable because of insufﬁcient degrees of freedom, as shown by Eqs. (3.56) and (3.57). An alternative form of data treatment is therefore required. Considering the sequence of measurements shown in Fig. 3.10, consecutive pairs of blank and standard responses can be used to estimate the slope and intercept parameters required for calibration: aj ¼ I0 2 bj ¼

j ðI 2 I5 nu þ 2 0

1 j21 ðI1 2 I6 Þ 2 aj I1 2 Cs nu þ 2

3:60aÞ ð3:60bÞ

where Cs is the analyte concentration in the standard, j ¼ 0; 1; …; 6 is the index of the measurement beginning with the blank at j ¼ 0; nu is the number of unknowns and ðnu þ 2Þ is the total number of measurements made before the blank and standard are next analyzed (assuming one blank and one standard per recalibration interval). Here it is assumed that the sensitivity drifts linearly between recalibrations. The concentration of analyte in the unknown can then be estimated as: Cu;j ¼

Iu; j 2 aj bj

ð3:61Þ

Assuming that the uncertainty in the concentration of analyte in the standard is negligible, as will generally be the case, propagating the uncertainties in

Fig. 3.10. Experimental design for external calibration in the presence of non-constant sensitivity. The number above each response bar is the index, j; of the measurement. Note that, if more unknowns had to be analyzed after the second measurement of the standard, j ¼ 6; then indices 5 and 6 would be reset to 0 and 1, respectively, and the cycle continued.

73

Douglas C. Baxter and Ilia Rodushkin

Eqs. (3.60) and (3.61) leads to: (

2

2 )0:5 j j 2 2 þs ðI5 Þ sðaj Þ ¼ s ðI0 Þ 1 2 nu þ 2 nu þ 2 sðbj Þ ¼

1 j21 2 2 j21 2 2 s2 ðI1 Þ 1 2 þs ðI6 Þ þs ðaj Þ b j Cs nu þ 2 nu þ 2

" #2 )0:5 ( Iu; j 2 aj 1 2 2 2 s ðIu; j Þ þ s ðaj Þ þ s ðbj Þ sc ðCu; j Þ ¼ bj bj

ð3:62aÞ 0:5

ð3:62bÞ

ð3:63Þ

a series of expressions rather more complicated than might be anticipated for single point calibration. Of course, this complexity arises out of the need to correct for the drifting blank and standard signal levels, requiring ﬁve separate measurements to calculate one concentration. (As an aside, it can be mentioned that all concentrations computed from consecutive recalibration cycles are correlated, as at least one pair of blank and standard measurements will be common to all the calculations; see discussion in Section 3.6.1.)

3.7

METHOD OF STANDARD ADDITIONS

When the matrix exerts non-spectral interferences, calibration must be performed with the sample itself [11,17,18,35]. The most common solution to this calibration problem is application of the method of standard additions. This typically involves adding either (a) variable volumetric increments ðVs;i Þ of a standard solution ðCs Þ or (b) a ﬁxed volume ðVs Þ of varying concentration ðCs;i Þ standards to constant-volume sample aliquots ðVu Þ; and diluting to a predetermined volume ðVT Þ before measurement. Both cases, illustrated in Fig. 3.11, correspond to spiking the sample at known concentration levels. Depending on the units of concentration employed, volumes may be replaced by masses, as the situation requires. A crucial aspect of this mode of calibration is that the matrix exerts the same effect on the added analyte as it does on the analyte originally present in the unknown. Failure to conform to this requirement will result in the generation of analytical data subject to systematic errors. For trace element determinations, it is therefore good analytical practice to add analyte in the same chemical form as that present in the unknown [11]. It should also be emphasized that this mode of calibration can correct for matrix-induced, but not temporal, variations in sensitivity when carefully applied. For cases (a) and (b), the total analyte concentration in any given spiked sample is expressed by Eq. (3.64a) or (3.64b), respectively, CT;i ¼ C0u Vu =VT þ Cs Vs;i =VT ¼ Cdu þ Cds;i

74

ð3:64aÞ

Calibration approaches for trace element determination

Fig. 3.11. Illustration of the experimental design for the method of standard additions using (a) constant concentration and variable volume additions, or (b) constant volume and variable concentration additions.

CT;i ¼ C0u Vu =VT þ Cs;i Vs =VT ¼ Cdu þ Cds;i

ð3:64bÞ

the superscript “d” indicating the dilution operation. Implicitly assuming a ﬁrst-order linear relationship between response and concentration, the recorded analytical signals for the series of solutions can be described by an expression of familiar form: Ii ¼ bSA CT;i ¼ bSA ðCdu þ Cds;i Þ ¼ aSA þ bSA Cds;i

ð3:65Þ

where aSA ¼ bSA Cdu and hence: Cdu ¼ aSA =bSA

ð3:66Þ

Thus, the analyte concentration in the diluted unknown is given by the ratio of the intercept to the slope for the method of standard additions (distinguished by the subscript “SA”). Taking account of Eq. (3.64), the concentration in the initial unknown is: C0u ¼ Cdu VT =Vu

ð3:67Þ

The least-squares techniques, OLR or WLR, as described in Sections 3.5.1 and 3.5.2, respectively, may be used to determine the intercept and slope. 3.7.1

Estimating uncertainty

Calculation of the combined uncertainty for the method of standard additions is normally based on Eq. (3.66), thus neglecting uncertainties in volume

75

Douglas C. Baxter and Ilia Rodushkin

measurement [17,18,35]: " #0:5 SSA;o 1 I2 d þ 2 P d sc ðCu Þo ¼ ds Þ2 bSA;o n bSA;o ðCs;i 2 C

ð3:68Þ

where the subscripts on the mean square about regression ðSÞ and slope terms are reminders that the method of standard additions and OLR are being employed. Calculation of these parameters is as described in Section 3.5.1, Eqs. (3.14) and (3.15). Similarly, for the use of WLR, the combined uncertainty can be deﬁned as: " #0:5 SSA;w I2w 1 P sc ðCdu Þw ¼ þ 2 P ð3:69Þ ds;w Þ2 bSA;w wi bSA;w wi ðCds;i 2 C Again, computation of the slope and mean square about regression terms have been detailed previously, in Section 3.5.2, Eqs. (3.33) and (3.35), respectively. Having established estimates of the concentration and its uncertainty in the diluted unknown, using Eq. (3.67) gives: sc ðC0u Þ sc ðCdu Þ < C0u Cdu

ð3:70Þ

under the condition that the uncertainties in VT and Vu are negligible, as will generally be true. (The subscripts “o” and “w” denoting OLR and WLR have been dropped, as the equation is equally applicable in either case.) Otherwise, it is a simple matter to account for all three sources of uncertainty: " #0:5 2 d s2 ðVT Þ s2 ðVu Þ 0 0 sc ðCu Þ sc ðCu Þ ¼ Cu þ þ ð3:71Þ VT2 Vu2 ðCdu Þ2 Expanded uncertainties for concentrations estimated using either OLR or WLR, as appropriate, can then be calculated according to Eq. (3.10), noting that there are n 2 2 degrees of freedom. However, examination of Fig. 3.12 shows that the uncertainty bounds for the estimated concentration are not actually symmetric, as was also the case for external calibration. Thus, it should be recognized that the application of Eq. (3.10) would again imply some degree of approximation by inherently providing a symmetric estimate of the expanded uncertainty for the method of standard additions. 3.7.2

Optimizing precision

It would appear that OLR has been used almost exclusively in combination with the method of standard additions, although Gardner and Gunn [48] have demonstrated the precision advantage gained using WLR. Recalling Fig. 3.9, it was apparent that the uncertainties associated with concentrations estimated using non-zero intercept OLR increase considerably at the extremes of the

76

Calibration approaches for trace element determination

Fig. 3.12. Comparison of expanded uncertainty bands when applying OLR (dashed lines) and WLR (dotted lines) to analytical data acquired using the method of standard additions. The uncertainty bars are 95% conﬁdence limits for the measured responses, and are indicative of heteroscedasticity.

calibration region. As shown graphically in Fig. 3.12, the method of standard additions requires extrapolation of the ﬁtted line beyond the lower extreme of the experimental response range. As such, the width of the uncertainty band around the estimated concentration increases rapidly with the extent of the extrapolation required [35,49]. This effect is particularly acute with OLR, because the centroid of the ﬁtted line is further from the concentration-axis intercept than with WLR. From the 95% conﬁdence intervals for the responses included in Fig. 3.12, it is clear that uncertainty increases with response and hence the application of WLR is justiﬁed. For the example shown, including weighting factors in the calculations approximately halves the expanded uncertainty. One important point to bear in mind is that the apparent advantage of WLR, indicated in Fig. 3.12, is dependent on the uncertainty structure. As noted above (Section 3.5.2), analytical instruments have a tendency to generate increasing measurement uncertainties with response. If measurement uncertainty is truly independent of signal level, then the weighting factors will be constant and the WLR solution to the normal equations reduces to that of OLR. If the correct regression technique is used, the uncertainty in the concentration estimated using the method of standard additions would always decrease monotonically with increasing level of added analyte [48,50]. This is analogous

77

Douglas C. Baxter and Ilia Rodushkin

to one of the considerations made for optimizing precision using external calibration, speciﬁcally that increasing the calibration range reduces the uncertainties. An expanded calibration range maximizes the sum in the denominator of the second term on the right-hand side of Eq. (3.68) or (3.69) and decreases the extent of extrapolation. AsPfor external calibration, uncertainties are also minimized by increasing n (or wi in the case of WLR). 3.7.3

Accounting for non-constant sensitivity

If the sensitivity is not constant, the method of standard additions will tend to yield erroneous results. The magnitude of this bias will depend on the rate of change of the sensitivity over the time interval required to complete the analyses of the unspiked and spiked unknowns. Such errors can be minimized by the experimental design shown in Fig. 3.13, where each unknown (or blank) is analyzed before and after a single spiked test portion. The analyte concentration is then estimated using: bu ¼

I1 2 ðI0 þ I2 Þ=2 Cds

ð3:72Þ

Cu ¼

I0 þ I2 2bu

ð3:73Þ

where the slope is calculated individually for each unknown (or blank), u, on the basis of the three measurements. Correction for dilution is made, as before, using Eq. (3.67) and ﬁnally, blank subtraction is implemented, if required.

Fig. 3.13. Experimental design for standard additions in the presence of non-constant sensitivity. The number above each response bar is the index, j; of the measurement.

78

Calibration approaches for trace element determination

The uncertainties associated with the computation of the slope parameter, Eq. (3.72), and the estimated concentration, Eq. (3.73), are evaluated as: 1 ½s2 ðI0 Þ þ 4s2 ðI1 Þ þ s2 ðI2 Þ0:5 2Cs (

)0:5 1 I0 þ I2 2 2 2 2 sc ðCu Þ ¼ s ðI0 Þ þ s ðI2 Þ þ s ðbu Þ bu 2bu sðbu Þ ¼

ð3:74Þ ð3:75Þ

assuming negligible uncertainty in the standard used to spike the unknowns (or blank). 3.8

INTERNAL STANDARDIZATION

As will become apparent, the concentration of internal standard added to the unknown is the basis of this calibration technique. Consequently, the internal standard species should be absent from the unknowns. Alternatively, the species selected should occur at such low concentrations that the result is not signiﬁcantly affected. A useful rule of thumb is that the internal standard is added in a 100-fold excess over the highest concentration originally present in the unknowns. The resulting uncertainty in the measured signal level will thus drown any variations caused by the presence of varying concentrations of the internal standard species. Of course, this approach requires a certain amount of prior knowledge about the unknowns and their expected composition. To function properly as a surrogate analyte, the internal standard should exhibit similar chemical and physical properties. The greater the similarity, the more likely both are affected to the same extent by drift and non-spectral interference processes. It is also necessary to ensure that both the analyte and the internal standard can be detected without spectral interferences from other components present in the unknowns. Consequently, selection of an internal standard necessitates a considerable investment in terms of preliminary investigations [14–18]. It is certainly worth the effort, as internal standardization is one of the most versatile calibration approaches, by virtue of its ability to correct for sensitivity variations caused by instrument instability and non-spectral interferences. To apply internal standardization, it is necessary to establish the relationship between the sensitivities for the analyte and the internal standard. This is typically achieved by analyzing a series of calibrants containing varying analyte and constant internal standard concentrations. The ratio of analyte (subscript “X”) to internal standard responses, IX=IS;i ; is a function of the concentration ratio, CX=IS;i ; and can be modeled using the relationships: IX=IS;i ¼ aIS þ bIS CX=IS;i þ 1i IX=IS;i ¼

b0IS CX=IS;i

þ 1i

ð3:76Þ ð3:77Þ

79

Douglas C. Baxter and Ilia Rodushkin

depending on whether or not an intercept is included in the linear regression model. The model parameters are appropriate for internal standardization, as identiﬁed by the subscript “IS”, and may be calculated by OLR or WLR as discussed in Sections 3.5.1 and 3.5.2, respectively. Analyte concentrations in unknown samples, spiked with internal standard, can readily be determined on the basis of their response ratios and internal standard levels: IX=IS;u 2 aIS Cu ¼ CIS;u ð3:78Þ bIS IX=IS;u C 0u ¼ CIS;u ð3:79Þ bIS Note that addition of internal standard implies dilution of the unknowns. Obviously this dilution effect must be corrected before reporting the results, using Eq. (3.67), but it will not be considered implicitly in the following. Again, as for the method of standard additions (see Section 3.7.1), it is assumed that dilution does not provide a signiﬁcant contribution to the uncertainty in the concentration estimate.

3.8.1

Estimating uncertainty

If it is further assumed that CIS;u is known exactly, the combined uncertainty in the estimated analyte concentration can be readily determined, because Eqs. (3.78) and (3.79) are of the same form as Eqs. (3.46) and (3.47), respectively. " # 0:5 ðIX=IS;u 2 IX=IS Þ2 CIS;u u2 ðIX=IS;u Þ 1 sc ðCu Þo ¼ þ S2IS;o þ 2 P ð3:80Þ X=IS Þ2 m bIS;o n bIS;o ðCX=IS;i 2 C sc ðC0u Þo

CIS;u ¼ 0 bIS;o

sc ðCu Þw ¼

0:5

ð3:81Þ

CIS;u bIS;w

sc ðC0u Þw ¼

" # 2 IX=IS;u u2 ðIX=IS;u Þ 0 2 þ SIS;o 0 2 P X=IS Þ2 m bIS;o ðCX=IS;i 2 C

u2 ðIX=IS;u Þ ðIX=IS;u 2 IX=IS;w Þ2 1 þ 2 P þ S2IS;w P X=IS;w Þ2 m wi bIS;w wi ðCX=IS;i 2 C

CIS;u b0IS;w

"

2 IX=IS;u

u2 ðIX=IS;u Þ 0 2 þ SIS;w P 02 X=IS;w Þ2 m bIS;w wi ðCX=IS;i 2 C

#

0:5

ð3:82Þ 0:5

ð3:83Þ

Equations (3.80) and (3.81) apply to OLR with and without an intercept, respectively, whereas Eqs. (3.82) and (3.83) are the corresponding expressions

80

Calibration approaches for trace element determination

for WLR. The terms have been deﬁned in Section 3.6.1 following Eqs. (3.52)– (3.55), the same considerations made there applying even here. 3.8.2

Optimizing precision

Strategies for minimizing uncertainties in estimated analyte concentrations are the same as those pertaining to external calibration. Somewhat surprisingly, however, the question as to the optimum internal standard concentration to use does not appear to have received much attention. For this reason, some model calculations were performed, a selection of the results for regression models including the intercept being illustrated in Fig. 3.14. Several situations were considered: (i) a constant level of measurement uncertainty, corresponding to conditions for which OLR would be applicable; (ii) a constant level of relative measurement variance, i.e. u2 ðIÞ / I 2 ; (iii) response variability determined by Poisson (counting) statistics as ideally obtained using a mass spectrometer, i.e. u2 ðIÞ ¼ I; and (iv) a linear combination of (ii) and (iii). Note that WLR is required to treat data for the situations described by scenarios (ii) –(iv). From Fig. 3.14a it transpires that, for situations (i) and (ii), the choice of internal standard concentration is irrelevant. Furthermore, when the requirements for applying OLR are met, the precision attainable is only favorable at very high analyte signal levels. Fortunately, most analytical instruments generate absolute signal variations that are an increasing function of response or concentration, as mentioned earlier. As such, WLR should then be used and this will result in considerably improved precision in analyte concentration estimates. For cases (iii) and (iv), the results of Fig. 3.14b,c demonstrate that using a high concentration of internal standard is optimal at high, but not low, analyte concentrations. It is therefore of importance to be aware of the noise characteristics of the instrument in order to determine the optimum spiking protocol. Of course, if the unknowns are likely to contain small quantities of the internal standard species, high levels must be added, which may be detrimental for the precision achievable at lower analyte concentrations, depending on the uncertainty structure. 3.9

ISOTOPE DILUTION

Isotope dilution mass spectrometry has the potential for application to some 60 elements that have at least two stable isotopes. The availability of long-lived radionuclides of additional elements further extends the suite of analytes that are amenable to this calibration approach, as illustrated in Fig. 3.15 [20]. Although calibration curve-based methods for the calculation of analyte concentrations by isotope dilution have been developed [51–53], these require the use of rational polynomials and are outside the scope of this treatment. We

81

82 Douglas C. Baxter and Ilia Rodushkin

Fig. 3.14. Effect of the internal standard concentration ðCIS Þ used on the relative standard uncertainty ðRSU ¼ sðCu Þ=Cu £ 100%Þ of the estimated analyte concentration. For all examples, the same set of normally distributed random numbers was used, generated at concentration levels ranging from 0 to 106, assuming that IX ¼ CX ; IIS ¼ CIS ; aIS ¼ 110 and bIS ¼ 1; with a constant standard uncertainty, sðIÞ; of 50. Mean values at concentration levels of 0, 104, 105 and 106 ðn ¼ 4Þ were employed to generate calibration curves using internal standardization and OLR or WLR, as appropriate, with a speciﬁc type of noise included. In (a), curve 1 (left axis) was obtained using OLR [m ¼ 1 in Eq. (3.80)] with sðIX Þ ¼ sðIIS Þ ¼ 50; independent of CIS ; curve 2 (right axis) was obtained using WLR (mu ¼ 1 in Eq. (3.82)) with sðIX Þ=IX ¼ sðIIS Þ=IIS ¼ 0:01; i.e. 1% proportional noise at all signals levels, and was independent of CIS : In (b), response precision was assumed to be governed by Poisson statistics, i.e. s2 ðIÞ ¼ I; the curves showing the results for CIS at (1-thick, solid line) 103, (2-dashed line) 104, (3-dotted line) 105 and (4-solid line) 106. In (c), the response precision was described by the relationship s2 ðIÞ ¼ I þ ð0:01 £ IÞ2 ; i.e. a linear combination of counting statistics and proportional noise; CIS as for (b).

Calibration approaches for trace element determination

Fig. 3.15. Periodic table showing the elements for which isotope dilution mass spectrometry can be applied. The number of stable (or long-lived) naturally occurring isotopes is given above the element symbol. For the mono-isotopic elements Al, I and Th, nominal masses for long-lived radionuclides, which are readily available for use as tracers, are given in parentheses. Adapted from Ref. [20].

will therefore conﬁne the ensuing discussion to the most commonly adopted variant of isotope dilution analysis, i.e., that without a calibration curve. Under the provision that the added material is equilibrated with the native analyte, exact compensation can be made for incomplete recoveries or losses during the sample preparation, as well as for non-spectral interferences [54]. Like all the calibration approaches discussed earlier, however, correction for spectral interferences is not directly possible and so selection of the isotopes used must be made carefully to avoid potential contributions from polyatomic ions or isobars having the same nominal mass-to-charge ratios as the native and enriched isotope analyte ions. Alternatively, suitable empirical correction factors must be applied, or the isotope system of interest must be separated from interfering species during sample preparation. Being based on experimental measurements, the former approach leads to the introduction of additional sources of uncertainty that must be accounted for in the uncertainty budget. The latter approach is that commonly adopted prior to TIMS measurements, and is becoming ever more widely applied for ICP-MS-based procedures as well [24]. Isotope dilution requires that the isotopic composition of both the native analyte in the sample and that of the spike [55] be known. When known

83

Douglas C. Baxter and Ilia Rodushkin

amounts of sample and spike are mixed, the measured isotopic composition of the resulting mixture can be used to calculate the concentration of analyte in the original unknown. In many cases, the isotopic composition of the sample can be safely assumed to equal that given in the current listing from IUPAC [56]. In other cases, where considerable natural variations occur, e.g. as a result of radiogenic processes, it may be necessary to ﬁrst establish the isotopic composition of the unknown. One such example is lead, where three of the four naturally occurring isotopes, 206Pb, 207Pb and 208Pb, are the ultimate products of radioactive decay of 238U, 235U and 232Th, respectively. Only 204Pb is nonradiogenic, and the isotopic composition of lead in any sample will reﬂect the origin of the lead incorporated in the material, as well as the original concentrations of the parent isotopes [26]. The isotope ratio, IR, in a spiked unknown is given by: n1;u þ n1;t n1 IRm ¼ ¼ ð3:84Þ n2 m n2;u þ n2;t where n1 and n2 (mol) are the amounts of isotope (or isotopomer in the case of molecular species) present in the mixture (subscript “m”), in the unspiked sample (u) and in the enriched isotopic spike or tracer (t). Note that all isotope ratios mentioned here are consistently deﬁned in terms of moles of isotope 1 divided by moles of isotope 2; other conventions may be encountered in the literature [23]. Rearranging to express the terms as isotope ratios and solving for n1;s yields [22]: n1 n 2 1 IRt 2 IRm IR ðIRt 2 IRm Þ n n t 2 ¼ n2;t u ð3:85Þ n1;u ¼ n2;t 2 m ¼ n2;t n1 n2 IRm 2 IRu IRm =IRu 2 1 21 n2 m n1 u Isotope 1 is preferably the most abundant isotope in the sample, although the choice is also made on the basis of freedom from spectral interferences. Isotope 2 is generally a minor isotope in the sample, but available in highly enriched form. Thus, by deﬁnition, n1;u . n2;u and IRu . IRm . IRt : Equation (3.85) provides the number of moles of isotope 1 present in the unknown relative to the amount of isotope 2 in the spike. To convert the quantity n1;u into more traditional concentration units, due consideration must be given to the relative atomic (or molecular) weights of the native ðMu Þ and spiked ðMt Þ analyte, as well as to the atom fractions or relative abundances of isotopes 1 and 2 in the original unknown ðA1;u Þ and in the tracer ðA2;t Þ; respectively. The volumes (or masses, depending on the units of concentration employed) of unknown ðVu Þ and tracer ðVt Þ must also be taken into account: n1;u ¼

84

Cu Vu A1;u Mu

ð3:86aÞ

Calibration approaches for trace element determination

n2;t ¼

Ct Vt A2;t Mt

ð3:86bÞ

Substituting Eqs. (3.86) into Eq. (3.85), the isotope dilution equation is obtained, enabling the desired analyte concentration in the unknown to be calculated:

Mu Vt A2;t IRu ðIRt 2 IRm Þ ð3:87aÞ Cu ¼ Ct Mt Vu A1;u IRm 2 IRu

IRu ðIRt 2 IRm Þ Cu ¼ Ct F ð3:87bÞ IRm 2 IRu where F is a coefﬁcient summarizing the ratio preceding the bracketed term. Various alternative forms of the isotope dilution equation may be found in the literature [20,23], all giving the same result. The advantage of the present form is that the contributions of the unknown, tracer and mixture isotope ratios are shown explicitly. As isotope ratios are computed from the measured count rates for isotopes 1 and 2 as I1 =I2 ; it is important to use blank and background corrected responses for the calculations. Subtraction of the blank concentration (calculated in the same way as for an unknown) from the result for an unknown could lead to systematic errors unless the blank and all unknowns exhibit exactly the same measured intensity ratio. Many mass spectrometers, particularly older quadrupole models, yield fairly high background count rates across the entire mass-to-charge range that have no relation to true isotope ratios [57]. 3.9.1

Mass discrimination and detector dead time

As yet, no mention has been made of the fact that the raw isotope ratios generated by a mass spectrometer are not accurate reﬂections of the true values. In Eqs. (3.85) and (3.86), it is actually implied that ðn1 =n2 Þx ¼ IRx ¼ ðI1 =I2 Þx ; i.e., that the measured count rate ratio equals the molar ratio of isotopes 1 and 2 in the analyzed test portion. That this is not the case that depends on two clearly deﬁned factors, the most important resulting from variations in the transmission efﬁciencies of ions of differing mass-to-charge ratios, causing an instrumental mass discrimination effect [21,24,58 –60]. Thus, a measured isotope ratio ðIRmeas Þ will differ from the true value ðIRtrue Þ by a factor K : IRtrue ¼ IRmeas K

ð3:88Þ

If no correction for instrumental mass discrimination were made, the concentration calculated using Eq. (3.87) would be biased. For example, if all three isotope ratios were measured, and the uncorrected data were inserted in

85

Douglas C. Baxter and Ilia Rodushkin

Eq. (3.87), the calculated analyte concentration would be in error by a factor of 1=K (assuming that the same mass discrimination factor applied to each ratio). Habfast [59] and Mare´chal et al. [60] have given useful recent discussions of mass discrimination factors, and the inclusion of uncertainty contributions has been described by Garcı´a Alonso [23]. For systems consisting of three or more isotopes (refer to Fig. 3.15), it is possible to correct for mass discrimination effects using an internal normalization technique [61]. Alternatively, again for polynuclidic elements, double or triple spiking can be used to provide mass discrimination correction [8,62 –65]. For descriptions of these methodologies, reference should be made to the relevant literature. At low concentrations, or when measuring isotopes of low abundance, electron multipliers are employed to enable registration of the smallest ion count rates. As a result of the ﬁnite time required by the detection system to process each ion signal pulse, the electron multiplier exhibits a dead time during which no further ions can be detected. At higher concentrations, the probability that an ion reaches the detector while a previous signal pulse is still being processed increases, leading to a concomitant loss in count rate [21,66, 67]. Thus, the detector dead time constitutes a second source of error in measured isotope ratios. For most ion counting systems, it is necessary to determine the dead time experimentally. This has been a subject of considerable interest in recent years, leading to the development of a variety of experimental methods for dead time determination [21,68–70]. Having determined the dead time, t (s), correction of the measured count rates, It ¼

I 1 2 It

ð3:89Þ

and hence of the isotope ratio, can be implemented: IRK;t ¼ K

I1 I2

1 2 tI2 1 2 tI1

¼

IRmeas K 2 tI1 1 2 tI1

ð3:90Þ

Note that correction for mass discrimination is also required, and included in the above expression. The rightmost form of Eq. (3.90) has been introduced to reduce the number of variables, as this will assist in the uncertainty analysis later. 3.9.2

Estimating uncertainty

The coefﬁcient F in Eq. (3.87b) can sometimes be assumed to contribute little uncertainty to the concentration estimate [22]. Nevertheless, some caution in making this assumption is warranted, particularly when the exact isotopic composition of the analyte or the spike is uncertain. The combined uncertainty

86

Calibration approaches for trace element determination

in F can be readily estimated: sc ðFÞ ¼ F

s2 ðA1;u Þ s2 ðMu2 Þ s2 ðVt2 Þ s2 ðA2t Þ s2 ðMt2 Þ s2 ðVu2 Þ þ þ þ þ þ 2 2 2 2 2 A1;u Mu Vt At Mt Vu

0:5

ð3:91Þ

and factored into the calculation of the combined uncertainty for the analyte concentration, if deemed necessary. Garcı´a Alonso [23] provides detailed accounts of the calculation of uncertainties in atomic weights and relative abundances. Note that the measurement of volumes (or masses) of unknowns and tracers can also introduce substantial uncertainties to the factor F: Propagating the uncertainties in Eq. (3.87b) and evaluating the partial derivatives provides the necessary relationship for the combined uncertainty in the analyte concentration estimated by the isotope dilution analysis of the unknown [22,23,71,72]: (

2 s2 ðCt Þ s2 ðFÞ s2 ðIRu Þ IRm sc ðCu Þ ¼ Cu þ c 2 þ 2 2 IRm 2 IRu Ct F IRu ð3:92Þ )

2 2 0:5 s2 ðIRt Þ IRt s2 ðIRm Þ IRm ðIRu 2 IRt Þ þ þ IRt 2 IRm ðIRt 2 IRm ÞðIRm 2 IRu Þ IR2t IR2m Note that, when the purity or stoichiometry of the enriched tracer is in doubt, the terms Ct and IRt must be determined experimentally, and might thus constitute a major source of uncertainty in the analytical result. It also follows that the terms Mt and A2;t will have to be evaluated in Eq. (3.87a), introducing further uncertainties. We should also emphasize that it has been assumed that all the uncertainties are independent, thus covariance terms have been omitted. Adding a term allows the variance introduced by correcting for mass discrimination effects to be included in the combined uncertainty for the isotope ratio [23]: " #0:5 sc ðIRK Þ s2 ðIRmeas Þ s2 ðKÞ ¼ þ ð3:93Þ IRK IR2meas K2 where the subscript “K” denotes the nature of the correction applied to the isotope ratio. The combined relative uncertainty, in a mass discrimination [58] and dead time corrected isotope ratio [22,66], is given by [23,73]: ( 2 sc ðIRK;t Þ s2 ðIRmeas Þ 1 s2 ðKÞ ¼ þ 2 IRK;t 1 2 tI1 =IRmeas IRmeas K2 " þ

s2 ðI1 Þ s2 ðtÞ þ I12 t2

#

tI1 ðIRmeas 2 1Þ ðIRmeas 2 tI1 Þð1 2 tI1 Þ

2 )0:5

ð3:94Þ

87

Douglas C. Baxter and Ilia Rodushkin

Equations (3.93) and (3.94) can replace any, or all, of the affected isotope ratio terms in Eq. (3.92), as appropriate for the system under consideration. 3.9.3

Optimizing precision

Equation (3.92) provides a useful theoretical basis for optimizing the precision of isotope dilution measurements [20,22]. Analyte isotope ratios in the unknown and in the tracer are obviously ﬁxed, whereas that of the mixture can be controlled. The differences between isotope ratios appearing in the denominators of the last three terms in Eq. (3.92) should be maximized in order to minimize uncertainty magniﬁcation, i.e. those factors enclosed in square brackets. As the isotope ratio of the mixture approaches that of the unknown, a condition denoted underspiking, the term ðIRm 2 IRu Þ tends to zero and two of the uncertainty magniﬁcation factors become very large. In the case of overspiking, ðIRt 2 IRm Þ tends to zero, again resulting in considerable uncertainty magniﬁcation. Purely from the standpoint of uncertainty propagation, the optimum precision will be realized when [20]: IRm ¼ ðIRu IRt Þ0:5

ð3:95Þ

Note that this equation only applies assuming that the relative uncertainties are constant, i.e. uðIÞ / I: In practice, additional considerations must also be observed. On the one hand, mass spectrometric measurement yields optimum precision at isotope ratios near unity. On the other hand, if the analyte concentration in the unknown is very low, then it may be advantageous to add a three- to 10-fold excess of tracer to avoid measuring both isotopes close to the detection limit [20]. Potentially, such overspiking could result in the introduction of analytical errors in determined concentrations, unless dead time effects are accurately corrected for [54], so care must be exercised. Mass spectrometric measurements tend to obey Poisson statistics, i.e. uðIÞ < I 0:5 at least at low signal levels, and as such, the isotope ratio yielding the best precision for the spiked unknown tends to be overestimated by Eq. (3.95). This is best illustrated by an example, taken from the excellent treatment of uncertainties in isotope dilution mass spectrometry by Adriaens et al. [22]. Three uncertainty models are considered: (a) a constant level of relative measurement variance, i.e. u2 ðIi ti Þ=ðIi ti Þ2 ¼ constant ¼ ð0:002Þ2 in this example; (b) response variability determined by Poisson (counting) statistics, i.e. u2 ðIi ti Þ ¼ Ii ti ; and (c) a linear combination of (a) and (b). The corresponding uncertainties in the isotope ratio for the mixture are then: sðIRm Þ ¼ IRm

88

"

s2 ðI1 t1 Þ s2 ðI2 t2 Þ þ ðI1 t1 Þ2 ðI2 t2 Þ2

#0:5 ¼ ½2 £ ð0:002Þ2 0:5

ð3:96aÞ

Calibration approaches for trace element determination

0:5 1 1 0:5 1 t þ ¼ 1 þ IRm 1 t2 I1 t1 I2 t2 I 1 t1

0:5 sðIRm Þ 1 t ¼ 1 þ IRm 1 þ 2 £ ð0:002Þ2 t2 IRm I1 t1 sðIRm Þ ¼ IRm

ð3:96bÞ ð3:96cÞ

where I1 and I2 are the count rates (counts s21) measured for total times (s) of t1 and t2 for isotopes 1 and 2, respectively. Insertion of the appropriate form of Eqs. (3.96) in Eq. (3.92) allows the uncertainty in determined analyte concentrations to be assessed as a function of the isotope ratio in the mixture. (Here it is assumed that dead time correction is unnecessary, e.g. by using a Faraday cup detector [24].) Results of the calculations are plotted in Fig. 3.16. In the case of constant proportional noise, curve (a), the optimum isotope ratio in the mixture is that predicted by Eq. (3.95). The relative uncertainty curve is, in this example, symmetrical about the optimum on a logarithmic scale [22]. This is only true, however, when the relative uncertainties in IRu and IRt are equal. If the relative uncertainty in the ratio for the spiked unknown is a function of the isotopic composition, the position of the optimum is clearly shifted toward

Fig. 3.16. Effect of the isotope ratio in the spiked unknown on the uncertainty in the estimated analyte concentration determined by isotope dilution analysis. Parameters used in the simulation were: I1 ¼ 10,000 counts s21; t1 ¼ 100 s; t1 =t2 ¼ 1; sðCt Þ=Ct ¼ 0:001; sðFÞ=F ¼ sðKÞ=K ¼ 0; sðIRu Þ=IRu ¼ sðIRt Þ=IRt ¼ 0:01; sðtÞ=t ¼ 0:25; IRu ¼ 415:7 and IRt ¼ 1:818: Curve (a) was obtained assuming the presence of proportional noise, Eq. (3.96a); curve (b) for Poisson statistics, Eq. (3.96b); and curve (c) for a linear combination of both types of noise, Eq. (3.96c).

89

Douglas C. Baxter and Ilia Rodushkin

the tracer composition, as shown by curves (b) and (c) in Fig. 3.16, corresponding to Eqs. (3.96b) and (3.96c), respectively. This means that a greater quantity of enriched isotopic tracer must be added to the unknown to minimize the uncertainty in the determined analyte concentration. Prior knowledge of the uncertainty structure is therefore of importance in determining optimal spiking regimes for isotope dilution analysis. It is also worth mentioning that, for the conditions used for the simulations in Fig. 3.16, the uncertainty contribution made by dead time correction was negligible. Indeed, uncertainty budgets suggest that correction for mass discrimination is often the precision-limiting factor in isotope ratio measurement [73].

Acknowledgements This work was ﬁnancially supported by EUs structural fund for Objective 1 Norra Norrland and Analytica AB, Lulea˚, Sweden, http://www.analytica.se/. REFERENCES 1 2 3

4

5 6 7

8

9 10

11 12

90

E. Prichard, Quality in the Analytical Chemistry Laboratory. Wiley, Chichester, 1997, Ch. 1. D.A. Skoog, F.J. Holler and T.A. Nieman, Principles of Instrumental Analysis, 5th edn. Harcourt Brace, Philadelphia, PA, 1998, Ch. 23. S. Kapila, D.O. Duebelbeis, S.E. Manahan and T.E. Clevenger, Flame photometric detectors. In: R.M. Harrison and S. Rapsomanikis (Eds.), Environmental Analysis Using Chromatography Interfaced with Atomic Spectroscopy. Ellis Horwood, Chichester, 1989, Ch. 3. J. Vessman, R.I. Stefan, J.F. van Staden, K. Danzer, W. Lindner, D. Thorburn Burns, A. Fajgelj and H. Muller, Pure Appl. Chem., 73 (2001) 1381. Available at http://iupac.chemsoc.org/publications/pac/2001/pdf/7308x1381.pdf. International Union of Pure and Applied Chemistry, Spectrochim. Acta, 33B (1978) 247. ISO, Guide to the Expression of Uncertainty in Measurement. International Standards Organization, Geneva, 1993. NIST, Guidelines for Evaluating and Expressing the Uncertainty of NIST Measurement Results, Technical Note 1297, National Institute of Standards and Technology, Gaithersburg, MD, 1994. Available at http://physics.nist.gov/ Document/tn1297.pdf. EURACHEM/CITAC Guide, Quantifying Uncertainty in Analytical Measurement, 2nd edn. 2000. In: S.L.R. Ellison, M. Rosslein and A. Williams (Eds.) Available at http://www.eurachem.bam.de/guides/quam2.pdf. International Union of Pure and Applied Chemistry, Pure Appl. Chem., 45 (1976) 99. Available at http://iupac.chemsoc.org/reports/V/spectro/partII.pdf. M. Thompson, S.L.R. Ellison, A. Fajgelj, P. Willetts and R. Wood, Pure Appl. Chem., 71 (1999) 337. Available at http://iupac.chemsoc.org/publications/pac/1999/ 71_02_pdf/thompson.pdf. B. Welz, Fresenius Z. Anal. Chem., 325 (1986) 95. E.H. Evans and J.J. Giglio, J. Anal. At. Spectrom., 8 (1993) 1.

Calibration approaches for trace element determination 13 14 15 16 17 18 19 20 21

22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49

I.I. Stewart and J.W. Olesik, J. Anal. At. Spectrom., 13 (1998) 843. J.J. Thompson and R.S. Houk, Appl. Spectrosc., 41 (1987) 801. R.F.J. Dams, J. Gossens and L. Moens, Mikrochim. Acta, 119 (1995) 277. I. Rodushkin, T. Ruth and D. Klockare, J. Anal. At. Spectrom., 13 (1998) 159. D.A. Skoog, F.J. Holler and T.A. Nieman, Principles of Instrumental Analysis, 5th edn. Harcourt Brace, Philadelphia, PA, 1998, Ch. 1. D.C. Harris, Quantitative Chemical Analysis, 6th edn. Freeman, New York, 2002, Ch. 5. R.J. Herberg, Anal. Chem., 35 (1963) 786. J.D. Fassett and P.J. Paulsen, Anal. Chem., 61 (1989) 643A. G.P. Russ III, Isotope ratio measurements using ICP-MS. In: A.R. Date and A.L. Gray (Eds.), Applications of Inductively Coupled Plasma Mass Spectrometry. Blackie, Glasgow, 1993, Ch. 4. A.G. Adriaens, W.R. Kelly and F.C. Adams, Anal. Chem., 65 (1993) 660. J.I. Garcı´a Alonso, Anal. Chim. Acta, 312 (1995) 57. I.T. Platzner (Ed.), Modern Isotope Ratio Mass Spectrometry. Wiley, New York, 1997. K.P. Huber and G. Herzberg, Molecular Spectra and Molecular Structure IV. Constants of Diatomic Molecules. Van Nostrand, New York, 1979. G. Faure, Principles of Isotope Geology, 2nd edn. Wiley, New York, 1986. V.J. Barwick and L.R. Ellison, Analyst, 124 (1999) 981. J. Kragten, Analyst, 119 (1994) 2161. J.C. Miller and J.N. Miller, Statistics for Analytical Chemistry, 3rd edn. Ellis Horwood, Chichester, 1993, Ch. 2. D.A. Skoog, F.J. Holler and T.A. Nieman, Principles of Instrumental Analysis, 5th edn. Harcourt Brace, Philadelphia, PA, 1998, Appendix 1. D.C. Harris, Quantitative Chemical Analysis, 6th edn. Freeman, New York, 2002, Ch. 3. F.E. Satterthwaite, Psychometrika, 6 (1941) 309. D.L. MacTaggart and S.O. Farwell, J. AOAC Int., 75 (1992) 594. N.R. Draper and H. Smith, Applied Regression Analysis, 2nd edn. Wiley, New York, 1981, Ch. 1. J.C. Miller and J.N. Miller, Statistics for Analytical Chemistry, 3rd edn. Ellis Horwood, Chichester, 1993, Ch. 5. Analytical Methods Committee, Analyst, 113 (1988) 1469. Y. Hu, J. Smeyers-Verbeke and D.L. Massart, J. Anal. At. Spectrom., 4 (1989) 605. P. Vankeerberghen, J. Smeyers-Verbeke, A. Thielemans and L. Massart, Analusis, 20 (1992) 103. J.D. Ingle Jr. and S.R. Crouch, Spectrochemical Analysis. Prentice Hall, Englewood Cliffs, NJ, 1988, Ch. 5. L.M. Schwartz, Anal. Chem., 51 (1979) 723. J.S. Garden, D.G. Mitchell and W.N. Mills, Anal. Chem., 52 (1980) 2310. P.D.P. Taylor and P. Schutyser, Spectrochim. Acta, 41B (1986) 1055. R.L. Watters Jr., R.J. Carroll and C.H. Spiegelman, Anal. Chem., 59 (1987) 1639. S.N. Ketkar and T.J. Bzik, Anal. Chem., 72 (2000) 4762. D.L. MacTaggart and S.O. Farwell, J. AOAC Int., 75 (1992) 608. J.H. Williamson, Can. J. Phys., 46 (1968) 1845. M.H. Feinberg, J. Chemometrics, 3 (1988) 103. M.J. Gardner and A.M. Gunn, Fresenius Z. Anal. Chem., 325 (1986) 263. I.L. Larsen, N.A. Hartmann and J.J. Wagner, Anal. Chem., 45 (1973) 1511.

91

Douglas C. Baxter and Ilia Rodushkin 50 51 52 53 54 55 56 57

58 59 60 61 62 63 64 65 66 67

68 69 70 71 72 73

92

K.L. Ratzlaff, Anal. Chem., 51 (1979) 232. J.A. Jonckheere, A.P. De Leenheer and H.L. Steyaert, Anal. Chem., 55 (1983) 153. A.R. Driedger III, D.C. Thornton, M. Lalevic and A.R. Bandy, Anal. Chem., 59 (1987) 1196. H. Klinkenberg, W. Van Borm and F. Souren, Spectrochim. Acta, 51B (1996) 139. T. Catterick, B. Fairman and C. Harrington, J. Anal. At. Spectrom., 13 (1998) 1009. P. De Bie`vre, J.R. De Laeter, H.S. Peiser and W.P. Reed, Mass Spectrom. Rev., 12 (1993) 143. K.J.R. Rosman and P.D.P. Taylor, Pure Appl. Chem., 70 (1998) 217. Available at http://www.iupac.org/reports/1998/7001rosman/index.html. D. Beauchemin, Current status of ICP-MS. In: D. Barcelo´ (Ed.), Discrete Sample Introduction Techniques for Inductively Coupled Plasma Mass Spectrometry. Elsevier, Amsterdam, 2000, Ch. 1. P.D.P. Taylor, P. De Bie`vre, A.J. Walder and A. Entwistle, J. Anal. At. Spectrom., 10 (1995) 395. K. Habfast, Int. J. Mass Spectrom., 176 (1998) 133. C.N. Mare´chal, P. Te´louk and F. Albare`de, Chem. Geol., 156 (1999) 251. L.J. Moore, L.A. Machlan, W.R. Shields and E.L. Garner, Anal. Chem., 46 (1974) 1082. L.A. Dietz, C.F. Pachucki and G.A. Land, Anal. Chem., 34 (1962) 709. M.H. Dobson, Geochim. Cosmochim. Acta, 34 (1970) 1241. R.D. Russell, J. Geophys. Res., 76 (1971) 4949. S.J.G. Galer, Chem. Geol., 157 (1999) 255. J.M. Hayes and D.A. Schoeller, Anal. Chem., 49 (1977) 306. P.J. Turner, D.J. Mills, E. Schro¨der, G. Lapitajs, G. Jung, L.A. Iacone, D.A. Haydar and A. Montaser, Instrumentation for low- and high-resolution ICPMS. In: A, Montaser (Ed.), Inductively Coupled Plasma Mass Spectrometry. Wiley, New York, 1998, Ch. 6. A.J. Fahey, Rev. Sci. Instrum., 69 (1998) 1282. S.M. Nelms, C.R. Quetel, T. Prohaska, J. Vogl and P.D.P. Taylor, J. Anal. At. Spectrom., 16 (2001) 333. H. Rameba¨ck, M. Berglund, D. Vendelbo, R. Wellum and P.D.P. Taylor, J. Anal. At. Spectrom., 16 (2001) 1271. C.S.J. Wolff Briche, C. Harrington, T. Catterick and B. Fairman, Anal. Chim. Acta, 437 (2001) 1. L. Yang, Z. Mester and R.E. Sturgeon, Anal. Chem., 74 (2002) 2968. P.K. Appelblad, I. Rodushkin and D.C. Baxter, Anal. Chem., 73 (2001) 2911.

Chapter 4

Stated references for ensuring traceability of trace element analysis Ph. Quevauviller

4.1

INTRODUCTION

Measurements constitute one of the foundations of modern society. Among the billions of analyses performed every year around the world, trace element determinations are a key issue in many sectors of industrial and societal importance (health, food, environment, industrial production). Sound decisionmaking, hence, calls for measurement systems capable of producing analytical data of demonstrated quality. Analytical problems to be tackled with respect to trace element analyses are numerous, and concern a wide range of different matrices, which are analysed for a wide variety of elements and their species at various levels of concentrations. This issue is becoming increasingly complex regarding the analytical problems encountered and the pressure that laboratories are facing with respect to providing ﬁt-for-purpose data quality. The last decade has seen an increasing awareness of the need for quality assurance (QA) in all the sectors directly relying on analytical data, which has been reﬂected by the development of a number of guidelines and documented standards, e.g. for managerial aspects and technical operations (e.g. sampling, method validation, etc.) and tools [e.g. reference materials (RMs), proﬁciency testing schemes] [1 –4]. In this context, traceability of chemical data is at the heart of the ongoing discussions. This concept is a heritage of metrology as conceived for physical measurements (e.g. mass, length, time, temperature, etc.) more than a century ago. Metrology in chemistry is now actively discussed among experts in metrology and analytical chemistry in order to propose a system that would be applicable to complex chemical measurements [5]. Recently, the application of metrology concepts to environmental analysis has also been examined [6]. The discussions generally point out that the direct application of theoretical metrology concepts to chemical measurements is not possible, because of major differences between chemical and physical measurement processes, e.g., chemical analysis results are often strongly dependent upon the nature of Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

93

Ph. Quevauviller

samples (whereas physical measurements are less or not affected). With respect to trace element analysis, a wide variety of analytical problems are encountered in relation to different elements (and their species) and matrices. Preliminary steps are necessary (e.g. sampling, sample pre-treatment) that may have an effect on the ﬁnal result. When dealing with (routine) trace element analyses, these theoretical discussions seem to be very distant from real-life situations, and the practice is, in most cases, very far from theory. Even though the situation has drastically improved within the last few years, the warning made at the beginning of the 1990s [7] is still relevant: many chemists still do not pay sufﬁcient attention to the reliability of analytical results and confuse trueness and precision. With respect to traceability, the situation is even worse and this concept is prone to many misunderstandings when applied to chemical measurements. This chapter discusses traceability of trace element analysis and examines, in particular, the various stated references to which the chemical data may be traceable. 4.2

MEANING OF TRACEABILITY FOR CHEMICAL MEASUREMENTS

ISO deﬁnes traceability as “the property of the result of a measurement or the value of a standard whereby it can be related to stated references, usually national or international standards, through an unbroken chain of comparisons all having stated uncertainties” [8]. In this deﬁnition, three key elements may be distinguished, which have been extensively discussed with respect to their applicability to chemical measurements: (1) the link to stated references, (2) the unbroken chain of comparisons and (3) the stated uncertainties. Detailed discussions have already investigated how these elements apply to chemical measurements [9 –12]. Let us now examine how they may be understood in the context of trace element analysis. In the deﬁnition, the stated references may be reference methods, RMs or SI units (kg or mole for chemical measurements) [13]. In theory, all chemical measurements should aim at being traceable to SI units. In practice, measurements correspond to approximations via comparisons of amounts, instrumental response generated by a number of particles, etc. Establishing SI traceability nowadays implies demonstrating to what extent these approximations are clearly related to the stated references [9]. As discussed below, most of the trace element analyses performed nowadays are actually traceable either to an RM [pure substances or matrix certiﬁed reference materials (CRMs)] or to a reference method (e.g. standardised method). The unbroken chain of comparison basically means that there is no loss of information during the analytical procedure (e.g. incomplete recovery, contamination). Achieving this requirement is more or less difﬁcult according to the analytical problem considered. It will be more critical for techniques involving successive analytical steps (e.g. extraction, separation, detection in

94

Stated references for ensuring traceability of trace element analysis

the case of the determination of chemical species of elements) and less acute for direct measurement procedures (e.g. sensors) which may, however, be faced with other difﬁculties (e.g. lack of sensitivity or selectivity). An additional difﬁculty, which is being faced in many instances (e.g. in the case of environmental analyses), is sample collection and storage. These two steps form an integral part of the traceability chain which is too often forgotten. The third key element, the stated uncertainties, is also a critical feature that many analysts still overlook. The theory implies that the uncertainty of a measurement is based on the traceability and uncertainty of all the stated references that contribute to this measurement. In other words, uncertainty components should be estimated at each step of an analytical process, i.e., the smaller the chain of comparison the better the uncertainty of the ﬁnal result. Here again, theory is confronted by practice when dealing with complex trace element measurements (e.g. analysis of complex environmental or biological matrices). This chapter will not consider uncertainty matters, which are widely discussed in the literature [1,9,14], but will rather focus on the ﬁrst two elements, which are more likely to be prone to misunderstandings. Before starting these discussions, it is useful to remind the reader that traceability should not be confused with accuracy. The latter covers the terms trueness (closeness of agreement between the “true value” and the measured value) and precision (closeness of agreement between the results obtained by applying the same experimental procedure several times under prescribed conditions). General aspects of QA of analytical measurements (including considerations on accuracy) have been extensively discussed in the literature [1] and will not be repeated here. Let us underline the fact that a method that is traceable to a given stated reference is not necessarily accurate (i.e., the stated reference does not necessarily correspond to the “true value”), whereas an accurate method is always traceable to what is considered to be the best approximation of the true value (deﬁned as “a value, which would be obtained by measurement, if the quantity could be completely deﬁned and if all measurement imperfections could be eliminated” [8]). At another level, precision and uncertainty are also often confused and considered to be similar concepts, which is not correct because uncertainty includes both random and systematic errors (while precision is solely linked to random errors) [10]. Trace element analyses are generally based on a succession of actions, namely (i) sampling, storage and preservation of representative samples, (ii) pre-treatment of a sample portion for quantiﬁcation, (iii) calibration, (iv) ﬁnal determination and (v) calculation and presentation of results. Based on this, we may now consider in further detail the types of stated references that are relevant to ensure the traceability of trace element determinations.

95

Ph. Quevauviller

4.3

SI UNITS

Units of the “Syste`me International” (SI) correspond to internationally recognised fundamental units that are used in metrology. They establish units of length (metre), mass (kilogram), time (second), temperature (Kelvin), etc. The unit that underpins chemical measurements is the unit of amount of substance, the mole. In principle, all chemical measurement data should be traceable to the mole [5]. In practice there is no “12C mole” standard, and kg is needed to deﬁne the mole [8]. Therefore, chemical measurements, in general, (and trace element analysis in particular) are actually traceable to the mass unit, the kg. Trace element analyses are based on the determination of amount of substance per mass of matrix. One should not confuse this traceability to mass units with the traceability to the “true value” of the substance in the matrix. This is discussed in section 4.6.5. 4.4

DOCUMENTED STANDARDS

Standardisation is an important aspect of routine analytical work. Documented standards (norms) related to measurement procedures are designed to establish minimum quality requirements and to improve the comparability of analytical results. They also often represent the ﬁrst step of the introduction of techniques/methods into regulations. In this case, the reference is closely related to the documented protocol, representing one of the main links of the traceability chain. This aspect will be particularly acute when dealing with operationally deﬁned parameters (i.e. parameters determined following a strict analytical protocol), e.g. fractionation of elements in soils or sediments [15], leachable trace elements from paint matrices [16], etc., as the traceability chain may be broken if the protocol is not strictly followed. Standardised procedures (documented standards) have been developed for sampling strategies and analytical techniques (documented protocols describing in detail analytical procedures, from the sub-sampling to the actual determination). In the area of environmental monitoring, the implementation of documented standards bound to regulations has been criticised (e.g. EPA methods) as standardised methods may become outdated while still being enforced by regulations that have not been revised. It may indeed happen that the analytical state of the art in a certain ﬁeld has drastically improved but laboratories are still obliged to use old-fashioned inferior methods for legal reasons. Standardisation bodies have recognised this problem and now allow the progress in analytical technologies to ﬂow into standardisation work. The use of a standardised method does not guarantee that no errors will occur; it only provides an analytical framework that is considered as the reference for a given measurement. Let us consider a speciﬁc example: the determination of extractable forms of elements using a single or sequential extraction procedure. This approach is

96

Stated references for ensuring traceability of trace element analysis

widely used for soil and sediment analysis for studies on plant bioavailability or mobility of trace elements [15,17]. The measurements do not relate to speciﬁc forms of elements, but rather to amounts extracted by a given procedure and operationally deﬁned according to interpretations such as “mobile”, “carbonatebound” forms, etc. The comparability of data is only possible if the extraction (documented) protocols are strictly followed [18], i.e., the traceability of the ﬁnal results will be linked to this documented extraction protocol taken as reference. If a change in operational parameters is made by a laboratory, the traceability link will be broken. This comment applies to all the measurements that correspond to a partial extraction of a substance in a given medium. Detailed (documented) guidelines are difﬁcult to set up for sample collection and storage which, however, remain the primary source of error (and hence one of the weakest links of the traceability chain). As an example, recommendations are available in the scientiﬁc literature with respect to environmental monitoring [3,19], but there are very few examples of documented standards that can formally be used as stated references in this ﬁeld. Recent progress has been achieved in the ﬁeld of soil monitoring through a systematic evaluation of sampling and sample pre-treatment procedures in the framework of an interlaboratory study; a group of laboratories actually used their own methods on a reference site and compared them with a reference method that was optimised and ﬁnally proposed as a harmonised method [20]. Sampling standards generally deﬁne the method of sampling, the number of samples to be collected, their representation, the frequency of sampling (taking into account natural variations), the sampling techniques and tools, etc. Statistical sampling tools exist [21,22] but they are often neglected and are hardly applicable to practical cases. The natures of the sample and the substance to be monitored actually dictate the choice of the sampling, which is hence adapted case-by-case. A similar situation is encountered for sample storage for which recommendations are given with respect to protection of the samples from light and elevated temperatures. This situation is obviously unsatisfactory with respect to the comparability of data as no clear stated references may be presently used. 4.5

REFERENCE METHODS

Analytical methods differ in the link between the signal produced by a given determined substance and the signal obtained from the calibration material. For a vast majority of methods, the link is usually related to an amount of substance of established purity and stoichiometry. In some ﬁelds of trace element analysis, e.g. the determination of chemical forms of elements (also referred to as “speciation” [23]), the techniques are based on a succession of analytical steps such as extraction, derivatisation, separation and detection. This multiplies the risk that the traceability chain is broken owing to lack of proper tools (e.g. RMs containing actual analysed species, secondary

97

Ph. Quevauviller

standards, etc.) to accurately determine the result, e.g. extraction recoveries, derivatisation yields, etc. [12]. For other methods, e.g. XRF, the link is through CRMs (see Section 4.6.1). So-called primary methods are methods with the highest metrological qualities for which the uncertainty can be established in terms of SI units and the result is accepted without reference to an external calibrating material. These methods have few random errors and are supposed to be exempt from systematic errors; they are also referred to as “deﬁnitive”, “absolute” or “stoichiometric” methods (e.g. gravimetry, titrimetry, coulometry of simple solutions, etc.) [9]. Using primary methods guarantees, in principle, that measurements will be traceable to SI units, i.e., traceability links will be established to the “true value” of the amount of substance. One would think that “reference methods” should hence systematically be “deﬁnitive methods”. However, these methods mainly exist for trace element determinations. For chemical forms of elements (or organic compounds), there are no real deﬁnitive (or primary) methods for the reasons expressed above (analytical steps with impossibility to ﬁrmly demonstrate full recovery). As stressed above, primary methods theoretically enable the traceability of chemical measurements to the SI unit (i.e., to the mole) to be achieved. This has been demonstrated for relatively “simple” measurements, such as trace elements in sea water, using isotope dilution mass spectrometry (ID-MS) [24]. However, what can be obtained for inorganic parameters in water samples is far from achievable for the analysis of complex organic substances and matrices requiring a series of analytical steps (e.g. extraction, clean-up, etc.). In this case, the traceability chain will be broken at several stages and the stated references will only rely on approximations (recovery estimates). The better these approximations, the closer the traceability of the measurement to the true value. In many cases (e.g. for trace organometallic determinations), true “deﬁnitive” methods do not exist for environmental measurements as there are no means at present to give proof that extraction or chemical reactions (e.g. derivatisation) have yielded a 100% recovery. As an example, ID-ICP-MS has been used for determining tributyltin (TBT) in sediment and mussel matrices after HPLC separation [25]: one could think that the measurements were then of demonstrated trueness because, in principle, no loss could occur after separation. The situation, however, was that the ﬁnal results were traceable to the “true” value of TBT present in the extract but not necessarily to the true value in the sample; the link to extraction recovery simply hampers this traceability being achieved. It has been argued that the development of “reference measurement” procedures that would be adequately applicable to real sample matrices (rather than matrix-dependent methods) would be a much better trend than trying to develop thousands of matrix-matched CRMs (see Section 4.6.1) [24]. However, these “reference methods” also need to be validated, which cannot negate the necessity to develop suitable CRMs.

98

Stated references for ensuring traceability of trace element analysis

Methods based on internal or external calibration rely on the availability of calibrants of high purity and veriﬁed stoichiometry but this is only the last link of the traceability chain (i.e., calibration of the detector signal). In principle, all steps of an analytical technique should be recorded in such a way that the result of the ﬁnal determination is linked through an unbroken chain of comparisons to appropriate standards. In other words, ﬁrmly establishing traceability in analytical measurements means that several “primary” chemical RMs in the form of (ultra-)pure substances are interlinked by well known, quantitative, high-precision high-accuracy chemical reactions [24]. In practice, this is not achievable for a vast category of chemical measurements. In the case of the analysis of chemical forms of elements and starting from extraction, there is no way at present to ﬁrmly ensure that a substance has been fully recovered from a complex matrix; methods that are generally used (e.g. successive extractions or spiking procedures) enable the estimate of method reproducibility but do not necessarily demonstrate full recovery. For methods including a derivatisation step, there are few or no appropriate calibrants available to date to check the yield of derivatisation reactions, which represents an additional gap in the traceability chain. As a conclusion, these methods are dependent on a number of more or less well-controlled parameters that may vary from one sample to another. As stressed below, the analytical steps that rely on a recovery estimate can only be validated in comparison to independent methods, giving a good indication on data comparability but not necessarily on accuracy. Hence, few of these methods may be considered as reference methods unless they are documented down with a great level of detail, describing all the analytical operations and the limits of applicability of the methods. This is the case of “ofﬁcial methods” that are required regulations. These are faced with the problem discussed above regarding documented standards (possible risks of becoming outdated).

4.6

REFERENCE MATERIALS

4.6.1

The various categories of materials and related requirements

The role of RMs is, in principle, well known. Certiﬁed Reference Materials may be calibration materials (pure substances or solutions or materials of known composition for techniques requiring matrix-matched calibrants, e.g. XRF) or matrix materials representing as far as possible “real matrices” for the veriﬁcation of measurement processes. They must meet certain requirements before being accepted as references by the analytical community: †

Homogeneity has to be veriﬁed to ensure that all sub-samples of the RM taken for measurement will produce the same analytical result within the stated measurement uncertainty. This requires a veriﬁcation of the within-

99

Ph. Quevauviller

†

†

†

unit and between-unit for each batch of RM produced. The product must also specify the minimum sample intake for which homogeneity has been measured and may be guaranteed. Finally, the ease of re-homogenising the material after packaging must be taken into consideration. Stability is another important requirement, i.e., producers must test the stability of the RM and its sensitivity to light, humidity, microbial activity, temperature, time, etc., and state the usable lifetime of the RM. Long-term testing is required to validate the stability of a material under a variety of storage and transport conditions. The similarity of the RM to the real sample should, in principle, be as close as possible in terms of matrix composition. This is not always achievable in practice, and compromises are sometimes necessary (e.g. production of synthetic samples mimicking the composition of natural samples). Accuracy, uncertainty and traceability of certiﬁed values are other crucial parameters which have to be duly considered at the stage of certiﬁcation. They are discussed below.

Laboratory reference materials (LRMs) [also known as quality control materials (QCMs)] have the same basic requirements of representativeness, homogeneity and stability as CRMs. The distinction of these “LRMs” or “QCMs” with CRMs is only related to the fact that these materials are not certiﬁed and are generally produced on a much smaller scale, e.g., for interlaboratory studies or internal quality control (control charts), i.e., to monitor the performance of analytical methods with time (reproducibility) through the establishment of control charts [26]. In this view, control charts and related RMs may be considered as long-term stated references for analytical measurements. It has been stressed that the “reference” represented by an RM may not always be reliable because, in many cases, the RM does not have the “same” matrix as the unknown sample [24]. This is discussed in section 4.6.5. 4.6.2

Production

Several recent books have been written describing the requirements and procedures for the production of RMs in general [4] and for environmental science in particular [2,27], of which only the main elements are presented in this chapter. The general steps for the production of an RM are summarised in Table 4.1, which illustrates that the preparation of an RM requires substantial planning and appropriate production facilities. There are obvious differences between producing a small batch of RM for internal quality control of a laboratory and a large amount of material for certiﬁcation purposes (requiring half-industrial capacities), and the producer needs to be equipped accordingly. Various equipment for sampling, treatment (e.g. grinder, milling devices, etc.), packaging, stabilising procedures, etc. are described in the literature [2,4].

100

Stated references for ensuring traceability of trace element analysis TABLE 4.1 General steps in the production of an RM (adapted from Stoeppler et al. [4]) 1. Deﬁnition of the RM, including the matrix, the properties to be certiﬁed and the desired levels 2. Design of the sampling procedure 3. Design of sample preparation procedure 4. Selection of an appropriate method for homogeneity and stability testing 5. Design of the characterisation of the RM 6. Acquisition of the samples 7. Preparation of the samples 8. Homogeneity testing 9. Stability testing 10. Characterisation of the RM 11. Combination of the results of homogeneity and stability tests and of the characterisation for calculation of the expanded uncertainty 12. Establishment of certiﬁcate and, if appropriate, preparation of a certiﬁcation report

Once a sample is acquired, the producer may start to prepare the material in the state it will be used. It is particularly important at this stage to stabilise the material (if required) to prevent possible changes in composition of critical components and to homogenise the material so that all future sub-samples will be as identical as possible. If additional treatment is needed (e.g. reduction of grain size), it should occur at this stage. At this point, a preliminary assessment of the homogenisation process should be performed. As one of the main goals of RM production is to provide a stable material, stability tests should begin at an early stage of the production process. Ideally, these tests should be conducted over the expected lifetime of the RM prior to its distribution. The packaging of the homogeneous and stable material will be conducted according to the analyst’s expectations, i.e., some materials will be packed in large units (e.g. bottles containing 50 g of material allowing several analytical repetitions) whereas in some cases materials may be packaged as single-use samples (e.g. in the case of potentially unstable properties). Additional uniformity assessment will be performed after the packaging (e.g. grain size, colour), which will be followed by the analysis of the material for the analyte(s) of interest for certiﬁcation purposes. The certiﬁcate of analysis will be issued at the end of the overall process. Stability testing should, in principle, be continued over the useable lifetime of the RM, depending on its inherent stability and also on its rate of use.

101

Ph. Quevauviller

4.6.3

Methods used for material characterisation or certiﬁcation

There are several accepted methods for characterising and producing reference values of RMs and/or certiﬁed values of CRMs. The more widely accepted methods include: † †

†

Certiﬁcation using one deﬁnitive method, used when a primary method is available and m for the purpose of the certiﬁcation analyses. An example of such a method for trace element analysis is ID-MS. Certiﬁcation through interlaboratory testing, in which the reference/certiﬁed values are obtained by pooling results from several laboratories (having demonstrated their quality) and using laboratory means for the calculation of the values. Certiﬁcation using at least two independent methods.

Combination of the above approaches are often used to certify RMs, e.g. using two or more independent methods in the frame of an interlaboratory testing programme, after which the data are combined to obtain the reference/certiﬁed value. Note that the calculation of the uncertainty of reference/certiﬁed now takes into account ISO recommendations for assessing expanded uncertainties that include contributions from homogeneity and stability tests, characterisation results and results of certiﬁcation work [4]. Let us note that ofﬁcial organisations attempt, wherever possible, to produce RMs estimating the true values as closely as possible, following the above-mentioned approaches. In the case of matrix CRMs, this is mainly achieved by employing a variety of methods with different measurement principles in the material certiﬁcation study; if these methods are in good agreement, one may assume (but not ﬁrmly demonstrate) that no systematic error has been left undetected, and the reference (certiﬁed) values are the closest estimate of the true value. This approach possibly includes deﬁnitive methods (see above), which seldom exist for analyses involving an extraction or derivatisation step. In many instances, consensus values are accepted as true values reﬂecting the state of the art (hence ensuring data comparability). Discussions are ongoing on the fact that many matrix CRMs do not guarantee a full veriﬁcation of accuracy owing to possible remaining bias (e.g., all extraction methodologies, although being in good agreement, could be biased to a certain degree with no means to demonstrate it at the present stage). This is a point of discussion in section 4.6.5. 4.6.4

Use of reference materials

The use of RMs is also widely described in the literature [4,28], e.g. for validation purposes, interlaboratory testing, control charting, etc., and the reader is encouraged to consult these works directly if additional information is needed. As mentioned above, some (certiﬁed) RMs are intended for calibration

102

Stated references for ensuring traceability of trace element analysis

purposes; in these cases, the uncertainty of the certiﬁed value is of prime importance as it will affect the ﬁnal uncertainty of the measured value in the unknown sample. In the case of certiﬁed pure substances or calibrating solutions, the uncertainties of certiﬁed values are usually negligible in comparison with the method uncertainty. This is not the case with matrix CRMs that are, in principle, reserved for the validation of methods; these materials are used for calibration purposes for non-destructive methods (e.g. XRF) and the larger uncertainty of the certiﬁed values may lead to a too large uncertainty of the ﬁnal results (thus leading to semi-quantitative measurements). 4.6.5

Traceability of reference materials

In the ﬁeld of trace element analysis, the wide variety of matrices and substances encountered calls for a large availability of matrix RMs representative from various sample types (e.g. sediments, soils, plants, waters, biological tissues, industrial products, etc.). Each of these sample types has a wide range of samples, e.g., for a soil—clay, sandy, humic-rich and calcareous soils). Reference materials represent “physical” stated references to which measurements can be linked. As mentioned above, this traceability is often criticised because the requirement of matrix similarity between unknown samples and matrix CRMs is never achievable in practice and often compromises have to be found. It should be stressed that a correct result obtained with a matrix CRM does not give full assurance that “correct results” will be achieved when analysing unknown samples, owing to differences in matrix composition [12]. The question of traceability of matrix CRMs (representing complex chemical systems) to SI units, and hence their values as “reference”, is an ongoing debate. Traceability implies “an accurate realisation of the unit in which the property values are expressed”. Similar to the achievement of traceability to SI units, the “accurate realisation” is often hardly demonstrated in practice. Indeed, as underlined in Section 4.5, it is difﬁcult to demonstrate, e.g., that a 100% extraction recovery has been obtained for a given substance in a complex environmental matrix. The assumption of a 100% recovery will actually be more valid if the certiﬁed values have been obtained in the frame of interlaboratory studies using a variety of (different) techniques. Even though, in the absence of “primary” (or “deﬁnitive”) methods, the collaboratively obtained value may only be considered as a “consensus value”, reﬂecting the state of the art of a given method. This consensus value represents an excellent reference to achieve traceability in a given area, but does not necessarily correspond to the “true value” (which is actually not quantiﬁable in most complex environmental measurements). In addition, there are numerous ﬁelds of environmental monitoring for which RMs are lacking (or are available but with a matrix too far removed from

103

Ph. Quevauviller

the analysed samples) or cannot be prepared owing to their instability. This hampers traceability being achieved. In this case, other approaches have to be followed (e.g. interlaboratory studies, see below). In the case of good correspondence between the matrix of samples and the matrix of CRMs, this reference is certainly the most appropriate one to check the accuracy of analytical methods and compare the performance of a method with other methods (or other laboratories). Similar comments with respect to representativeness may be made concerning matrix LRMs (not CRMs, see Section 4.6.1) used for internal quality control purposes (establishment of control charts). In order to clarify the traceability links represented by RMs, a classiﬁcation has been established, categorising the various types of materials [29], as shown in Table 4.2. In this classiﬁcation, primary RMs are traceable to SI units through primary methods and physical standards (i.e., mass standards); CRMs and LRMs certiﬁed or reference values are obtained by reference or validated methods, but they are not mentioned to be necessarily traceable to SI units. The TABLE 4.2 Classiﬁcation of chemical RMs (adapted from Pan [29]) Level

Appellation

Criteria

I

Primary RM

II

Certiﬁed RM

III

Working RM (or LRM or QCM)

Materials with the highest metrological qualities, whose values are determined (certiﬁed) by a primary method Developed by a national metrological institute Recognised by national decision Traceable to SI units and veriﬁed by international intercomparisons Fulﬁl the ISO Guide 30 deﬁnition Generally developed by a national reference laboratory or a specialised organization Certiﬁed by reference methods, by comparisons of different methods or a combination of the two approaches Recognised by national or specialised organisations Accompanied by a certiﬁcate indicating the uncertainty of the certiﬁed values and describing the traceabilty Fulﬁl the ISO Guide 30 deﬁnition Produced by an accredited organisation Establishment of reference values by one or more validated methods Accompanied by a description of the achieved traceability and giving an estimate of the uncertainty

104

Stated references for ensuring traceability of trace element analysis

table suggests a traceability link between LRMs, CRMs and primary RMs, which may be achieved in some cases but not as a general rule, in particular for matrix RMs as discussed above. In other words, even if primary RMs are used for calibrating an analytical method used for obtaining certiﬁed (or reference) values, the uncertainties that may remain, e.g. on recoveries, do not allow one to ﬁrmly establish traceability to the primary RMs (and hence to SI units). This is discussed in one of the case studies below. 4.7

SPECIMEN BANKING

Specimen banking is another type of stated references that may be used in relation to trace element analysis. A speaking example of this application is environmental monitoring. The approach consists of collecting environmental samples, processing them and storing them on a long-term basis under conditions which prevent any signiﬁcant changes in their chemical composition [30]. The aim is to create a systematic repository of samples, providing information about current levels of pollution and tools to evaluate contamination trends. This approach certainly represents the best referential system for long-term environmental monitoring, enabling use of specimens as stated references for possibly repetitive analysis focusing on speciﬁc contamination studies (e.g. when more sophisticated (accurate) analytical methods will become available). Besides specimens that are “true” stated references of the environment status at the time, RMs can also be produced from surplus specimen material (i.e., stabilising them, e.g. by freeze-drying) in order to monitor the reproducibility of analytical techniques, hence ensuring internal quality control using the most representative samples [28]. “Fresh” RMs may also be prepared from samples similar to collected specimens and processed the same way (i.e., in an uninterrupted cryo-chain to preserve their integrity), homogenised and stored as fresh powder materials for the purpose of developing new analytical procedures, optimising existing methods, internal quality control and stability experiments of environmental specimens [31]. 4.8

PROFICIENCY TESTING

Participating in interlaboratory studies (or proﬁciency testing, which is the equivalent term used in regulations) is a way for laboratories to establish stated references for evaluating the performance of their methods. These exercises imply that one or more materials are distributed to several laboratories for the determination of given substances. The comparison of different methods enables the detection of possible sources of errors linked to a speciﬁc procedure or the way a method is applied by a given laboratory. Exercises focusing on a single method enable the establishment of performance criteria (e.g. precision).

105

Ph. Quevauviller

The stated references, here again, may be RMs that should meet homogeneity and stability requirements (see Section 4.6). However, in contrast to RMs used for internal quality control, proﬁciency testing may involve samples with a limited shelf life that are distributed to laboratories for analysis of particular parameters that could not be evaluated using stabilised RMs. Examples include “fresh” materials (e.g. biological samples) treated with a short-term preservation period. Similarly to what has been discussed for RMs, the measurement values obtained in relation to interlaboratory studies (using different techniques) are taken as the “best representation of the state of the art”, i.e., offering an excellent means for laboratories to achieve comparability (i.e., traceability) of their results to a recognised reference, which is in this case a consensus value (generally the mean of laboratory means). This reference does not enable traceability to the true value of the substance in the medium to be achieved, but it represents a very useful method for achieving comparability of environmental measurements. 4.9

4.9.1

REAL-CASE ACHIEVEMENT OF TRACEABILITY OF TRACE ELEMENT ANALYSIS Total trace element determinations

The ﬁrst example corresponds to the most “simple” system that may enable the traceability of trace element analysis to the SI unit to be achieved. Traceability of trace element determinations in various matrices to the mole is, in principle, achievable using primary methods, such as ID-MS. This does not infer that accurate results cannot be obtained with other methods (e.g. spectrometric methods relying on external calibration), but the traceability chain will be more prone to systematic errors in the case of calibration-dependent methods in comparison to primary methods. The example below, therefore, uses ID-MS as primary method and examines how traceability may be achieved in the framework of a “real-case” analysis. Figure 4.1 distinguishes four steps, separated into an operational part (sampling to detection) and a reference part representing the traceability to stated references. (A) Sampling is recognised to represent the major contribution to analytical errors. This has been clearly demonstrated in some areas, e.g. sampler contamination hampering use of trace element data in oceanic waters produced over the years 1965–1986 [32]. The high variability of results also depend on the way samples are collected, as illustrated by systematic comparisons of sampling methods used for trace element analysis in soils [20]. Considerable improvements in sampling methodologies, and in some instances standardisation, have permitted minimisation of possible contamination and losses of trace elements. The “stated reference” at

106

Stated references for ensuring traceability of trace element analysis

Fig. 4.1. Example of traceability chain for total trace element determinations.

this stage, hence, relies on the experience gained in the past and documented guidelines (standardised or not) that have been adopted as a consensus. It is not possible to demonstrate that all contamination sources have been avoided (explaining the dotted line in the reference links), but this is the best consensus that may be achieved at the present stage for the analysis of most matrices. A way to control the likelihood of contamination is to process a blank sample using the same procedures and materials as used for the unknown sample. (B) Similar to sampling, sample pre-treatment and storage represent a high risk of contamination and procedures are generally tested to minimise these sources of errors, in particular the sample stabilisation and storage. Here again, one has to rely on the common experience to assume that the traceability has not been drastically affected at this stage. Errors can be due to the addition of acids of insufﬁcient purity (e.g. when used to stabilise water samples at the collection stage), errors of manipulation, etc. As observed in step A, the reference link is also weak in this case as it is hardly possible to ﬁrmly identify all errors that may have occurred on the ﬁeld.

107

Ph. Quevauviller

(C) This step corresponds to the laboratory work, in this case determination of the trace elements by ID-MS. The method is considered to be a primary method as it is based on the addition of known amount (determined by weighing) of the analyte in an isotopic composition different from that of the analyte present in the sample. Spiking is achieved prior to chemical treatment of the sample and has to be performed in such a way that an equilibrium is reached between the spike and the isotopes naturally present in the sample. Considering the similarity between the isotopes, the chemical treatment does not affect the isotopic ratio even if the analyte recovery is not complete. Possible contamination or loss has no effects on the traceability of the result, as the analyte and spike will undergo the same pattern and the ratio will not be affected. (D) The amount of analyte is related to the amount of the spiked isotope according to a known formula [33]. The calculation is only based on isotopic ratios that may accurately be measured by mass spectrometry. The measurement is hence considered to be traceable to the mole. In the context of trace element analysis in various matrices, using ID-MS as a determination technique, the weakest parts of the chain are hence the sample collection and pre-treatment steps. An improvement in the traceability chain could be achieved if the isotope spiking was to be conducted immediately after collection, i.e., at the pre-treatment stage (step B). Isotope dilution mass spectrometry measurements, in theory, guarantee the traceability of the pretreated sample to the SI unit. However, an example of application of this technique in interlaboratory studies on trace elements in sea water has shown that even if this method is considered as a primary method, it is not without errors owing to the operating difﬁculties; an example has been published, showing that 5 laboratories out of 16 using ID-MS reached the required performance for the certiﬁcation of a sea water CRM [34]. As a conclusion, one may realise that even a relatively “simple” system with respect to the analytical measurements is not exempt from sources of errors, either due to possible contamination occurring, e.g. at the sampling stage (with few means to ﬁrmly demonstrate the lack of contamination, in particular, for the sampling itself) or manipulation errors at the determination stage (due to the operating complexity of ID-MS).

4.9.2

Operationally deﬁned trace element determinations

The second example deals with the determination of “extractable” contents of trace elements, i.e., analyses based on a given operational protocol (e.g. single or sequential extraction scheme, or leaching procedure). In this case, all steps prior to the analytical work are basically the same as for total trace element determinations, but the pre-treatment actually corresponds to a strictly

108

Stated references for ensuring traceability of trace element analysis

deﬁned protocol (that may be an ofﬁcial standardised method), representing a key “stated reference”. Traceability of determinations relies, in this case, on the proper application of the operational protocol and, of course, on the analytical measurement itself (ﬁnal detection of trace elements in the extract or the leachate). Figure 4.2 gives an account of the different steps, with indication of the references linked to traceability. (A,B) (C)

These steps are basically similar to their equivalent steps in Section 4.9.1. As stressed above, the laboratory work will start by an extraction (or leaching) procedure, strictly following a written procedure. The traceability will hence rely heavily on compliance to the written procedure (errors being made if different reagents are used or if the extraction/ leaching scheme is not applied as written). Trace element contents in the extract or leachate may be determined by ID-MS (as

Fig. 4.2. Example of traceability chain for extractable trace element determinations (based on the use of a single or sequential extraction scheme).

109

Ph. Quevauviller

(D)

discussed in step C of Section 4.9.1), even if this is not a current practice, i.e., ﬁnal trace element determinations are generally conducted by spectrometric methods. If spiking with a known amount of the analyte having an isotopic composition different from that of the analyte is achieved prior to the extraction procedure, possible contamination or losses will have no effect on the result traceability of the trace element contents in the extract. If we consider the possible ID-MS determination of trace elements in the extract, the calculation will be based on isotopic ratios (similar to step D of Section 4.9.1) and the measurement of extractable content will hence be considered to be traceable to the mole. In other words, we will be able to determine the “true value” of the trace elements in the extract, which does not necessarily mean that traceability to a welldeﬁned chemical form is achieved (i.e., traceability to the mole is hardly achievable).

This particular case represents a transition between the total trace element analysis and the next example, corresponding to well-deﬁned chemical forms of elements. This type of measurement is only comparable from one laboratory to another if a strictly followed standardised procedure is applied. Examples exist of procedures that have been adopted as a consensus method, e.g. single and sequential extraction procedures for trace element analysis in soils and sediments [15], or internationally adopted standardised procedures, e.g. the aqua regia extraction (ISO 11466 Standard) for the determination of “pseudo-total” contents of elements, and many others such as the EN 71-3: 1994 Standard for the evaluation of toxic trace element mobility from paints in the context of toy testing [16]. Very few of these schemes are, however, backed up by relevant CRMs (i.e., certiﬁed for their extractable trace element contents, following deﬁned protocols). The references [15,16] give examples of such CRMs, namely for soil and sediment analyses, and paint analysis.

4.9.3

Determinations of chemical forms of elements

In the case of determination of chemical “species”, the substance of concern is more complex (e.g. elements in different oxidation states, organometallic compounds) and requires a more sophisticated measurement approach (Fig. 4.3). The element species or compound is prone to possible alteration (change of oxidation state or degradation of organometallic compound) if insufﬁcient care is taken at the sample collection and storage steps. Analytical procedures have been extensively reviewed for speciation analyses [23,35,36] but, as discussed below, the measurement traceability chain presents many weak links in comparison to the ﬁrst case study.

110

Stated references for ensuring traceability of trace element analysis

Fig. 4.3. Example of traceability chain for the determination of chemical forms of elements.

(A,B)

In contrast to total trace element analysis, there are no formal recommendations regarding sample collection (step A) and storage (step B) for the analysis of chemical species in various matrices. A laboratory will only rely on published procedures, which are often described along general lines, lacking details on QA. Procedures will hence be adapted case-by-case, following “home-made” approaches, which are often hardly comparable sensu stricto from one laboratory to another and do not allow ﬁrm establishment of traceability owing to the lack of well-deﬁned stated references. In other words, the data

111

Ph. Quevauviller

(C)

(D)

(E)

(F)

produced on the basis on “in-house-made recipes” may or may not be of good quality. There is simply no means to anchor them to a ﬁrm reference system. Extraction methodologies also vary considerably from one laboratory to another. The extraction recoveries are, in principle, calculated, but here also there is no real consensus as to the approach to be followed [37]. One may say that there is no real need of setting up documented extraction protocols that could rapidly become outdated with the constantly improving analytical methodologies and it is only necessary to demonstrate that the methods used are validated. This validation is indeed possible on the basis of existing matrix CRMs [2]. The recovery check is, however, prone to the uncertainties discussed in Section 4.6 with respect to the “similarity” of matrix composition between the CRM and the unknown. Therefore, the traceability link exists but it is still questionable. Derivatisation reactions (e.g. hydride generation, Grignard reactions) are frequently used for the determination of chemical species in environmental matrices [38]. The traceability chain implies, in principle, that the derivatisation yields are veriﬁed, which is only possible on the basis of available “secondary” standards, i.e., pure derivatised calibrants (e.g. ethylated, pentylated, etc.). This validation has been followed in the case of certiﬁcation of RMs for TBT [39] but it is far from being a routine practice. As in step C, the completeness of the derivatisation yield will be evaluated on the basis of analysis of matrix CRMs, with the already expressed limitations. Separation will be necessary to isolate the different organotin compounds from possible interfering compounds from the matrix. Selectivity is the key feature here. The risk of losing traceability is related to a possible degradation of the compounds (heat-induced degradation on the column) or an insufﬁcient selectivity. Internal standards with a composition close to the determined compound are useful stated references to detect possible losses or insufﬁcient species separation. This part of the traceability chain is considered to be reasonably under control. High-purity primary standards for the calibration of chemical forms of elements are generally commercially available (except in some analytical ﬁelds, e.g. chemical species of As) and this ﬁnal link of the traceability chain (detection) is considered to be satisfactory.

This case study shows that a “real case” monitoring exercise is subject to many questions with respect to measurement traceability of chemical forms of elements. The weakest links are certainly the sample collection and storage, for which no strong reference system exists. The situation is better with regard to the analytical measurements, even if the links are not considered to

112

Stated references for ensuring traceability of trace element analysis

be that strong (e.g. with respect to evaluation of analyte recoveries). At present, one must admit that we are far from being able to ﬁrmly demonstrate measurement traceability to the “true value” of chemical forms of elements in many instances. 4.10

CONCLUSIONS

Trace element analyses can only be valid if the data are obtained under a reliable QA regime. Comparability of data is mandatory for evaluating spatial and temporal contamination trends, studying bio- or geo-chemical pathways, assessing product quality, evaluating risks, etc., and this is only achievable if harmonised approaches are considered, from sampling to ﬁnal detection, for the analysis of given substances in given media. Without the demonstration of data traceability to any kind of well-documented stated references, a considerable amount of data published in the scientiﬁc literature is actually totally useless and this represents a huge waste of resources. As a ﬁnal word, one should not confuse the search for traceability to welldeﬁned and accepted stated references to the achievement of accuracy. The ﬁrst concept is a moving feature, i.e., stated references may evolve with progress of knowledge and technical capabilities, while still maintaining comparability of data, but it does not necessarily mean that the resulting data are accurate (i.e., close to “true value”). This is partly compatible with the metrological principle of traceability which implies that “if the traceability of measurements is claimed to be other than the mole unit itself, but rather through a procedure, material or standard, then they must be credibly described and their relation to the mole clearly established” [24]. Indeed, the traceability of trace element analyses can be demonstrated to pure calibrating substances, CRMs or documented standards, the latter two often corresponding to “consensus” values and not “true” values, except in some speciﬁc cases (e.g., certiﬁed trace element contents in water matrices). Demonstrating traceability of an amount of substance to its true value in a given matrix is, therefore, hardly achievable in practice. We are in a world of compromises and the best compromise to date for many analytical issues is to achieve the best possible comparability of data both spatially (e.g. for trend studies) and geographically (between-laboratory comparability), which relies on physical tools (RMs and, in some cases, specimens). With respect to trend studies, this comparability quest should respond to progress in analytical sciences so that the stated references may be regularly improved while still maintaining a traceability chain with “old data”. Indeed, if analytical progress actually enables us to reﬁne the determination of certiﬁed (or reference) values, approaching their closeness to the true value, links with measurements conducted decades ago (but veriﬁed with RMs of lesser conﬁdence) will still be possible and data will not be lost. In other words, small detected biases (detected with more advanced techniques) could be

113

Ph. Quevauviller

corrected in the future if a system of RM banking is implemented. Therefore, one may hope that progress in analytical chemistry will be such within the next decades that the accuracy of measurements will be ﬁrmly demonstrated, i.e., traceability to the true amounts of contaminants in the environment will be established.

REFERENCES 1 2

3 4

5

6 7 8 9

10 11 12 13 14 15

16 17

18 19

114

H. Gu¨nzler (Ed.), Accreditation and Quality Assurance in Analytical Chemistry. Springer-Verlag, Berlin, 1996. Ph. Quevauviller and E.A. Maier, Certiﬁed Reference Materials and Interlaboratory Studies for Environmental Analysis—The BCR Approach. Elsevier, Amsterdam, 1999. D. Barcelo´ (Ed.), Sample Handling and Trace Analysis of Pollutants, 2nd edn. Elsevier, Amsterdam, 2000. M. Stoeppler, W.R. Wolf and P.J. Jenks (Eds.), Reference Materials for Chemical Analyses—Certiﬁcation, Availability and Proper Usage. Wiley-VCH, Weinheim, Germany, 2001, ISBN: 3-527-30162-3. B. King, M. Walsh, K. Carneiro, R. Kaarls, V. Komppa, C. Nieto de Castro and J. Lexow, Metrology in Chemistry—Current Activities and Future Requirements in Europe, EUR Report, EUR 19074 EN, European Commission, Brussels, 1999, ISBN: 92-828-7465-6. Ph. Quevauviller and O.F.X. Donard, Trends Anal. Chem., 20 (2001) 600. B. Griepink, Fresenius J. Anal. Chem., 338 (1990) 360. ISO, International Vocabulary of Basic and General Terms in Metrology, 2nd edn. International Standardisation Organisation, Geneva, Switzerland, 1993. M. Valca´rcel, A. Rı´os, E. Maier, M. Grasserbauer, C. Nieto de Castro, M.C. Walsh, F.X. Rius, R. Niemela¨, A. Voulgaropoulos, J. Vialle, R. Kaarls, F. Adams and H. Albus, Metrology in Chemistry and Biology—A Practical Approach, EUR Report, EUR 18405 EN, European Commission, Brussels, 1998, ISBN: 92-828-4049-2. M. Valca´rcel and A. Rı´os, Anal. Chem., 65 (1999) 78A. M.C. Walsh, Trends Anal. Chem., 18 (1999) 616. Ph. Quevauviller, J. Environ. Monit., 2 (2000) 292. B. King, Analyst, 112 (1997) 197. A. Maroto, R. Boque´ and F.X. Rius, Trends Anal. Chem., 18 (1999) 577. Ph. Quevauviller (Ed.), Methodologies for Soil and Sediment Fractionation Studies. The Royal Society of Chemistry, Cambridge, United Kingdom, 2002, ISBN: 0-85404453-1. P. Roper, R. Walker and Ph. Quevauviller, Fresenius J. Anal. Chem., 366 (2000) 289. A.M. Ure and C.M. Davidson (Eds.), Chemical Speciation in the Environment. Blackie Academic and Professional, London, United Kingdom, 1995, ISBN: 0-75140021-1. Ph. Quevauviller, Trends Anal. Chem., 17 (1998) 289. Ph. Quevauviller (Ed.), Quality Assurance in Environmental Monitoring— Sampling and Sample Pretreatment. VCH, Weinheim, Germany, 1995, ISBN: 3-527-28682-9.

Stated references for ensuring traceability of trace element analysis 20

21 22 23

24 25 26 27 28

29 30 31 32 33 34 35 36 37 38 39

G. Wagner, Ph. Quevauviller, A. Desaules, H. Muntau and S. Theocharopoulos (Eds.), Comparative Evaluation of European Methods for Sampling and Sample Preparation of Soils, Special issue of Sci. Total Environ., 2001, p. 264. F.M. Garﬁeld, Quality Assurance Principles for Analytical Laboratories. AOAC International, Arlington, USA, 1991. P.M. Gy, Mikrochim. Acta, II (1991) 457. L. Ebdon, L. Pitts, R. Cornelis, H. Crews, O.F.X. Donard and Ph. Quevauviller (Eds.), Trace Element Speciation for Environment, Food & Health. The Royal Society of Chemistry, Cambridge, United Kingdom, 2001, ISBN: 0-85404-459-0. P. de Bie`vre, in: H. Gu¨nzler (Ed.), Accreditation and Quality Assurance in Analytical Chemistry. Springer, Berlin, Germany, 1996. S.J. Hill, L.J. Pitts and A.S. Fisher, Trends Anal. Chem., 19 (2000) 120. T.H. Hartley, Computerized Quality Control: Programs for the Analytical Laboratory, 2nd edn. Ellis Horwood, Chichester, 1990. Ph. Quevauviller, Mate´riaux de Re´fe´rence pour L’environnement. Tec&Doc Editions, Paris, France, 2002, ISBN: 2-7430-0579-3. V. Barwick, S. Burke, R. Lawn, P. Roper and R. Walker, Applications of Reference Materials in Analytical Chemistry. The Royal Society of Chemistry, Cambridge, United Kingdom, 2001, ISBN: 0-85404-448-5. X.R. Pan, Metrologia, 34 (1997) 35. H. Emons, J.D. Schladot and M.J. Schwuger, Chemosphere, 34 (1997) 1875. T.-M. Sonntag and M. Rossbach, Analyst, 122 (1997) 27. G. Topping, Sci. Total Environ., 49 (1986) 9. W. Richter, Accredit. Qual. Assur., 2 (1997) 354. B. King, Analyst, 112 (1997) 197. Ph. Quevauviller, Method Performance Studies for Speciation Analysis. The Royal Society of Chemistry, Cambridge, United Kingdom, 1998, ISBN: 0-85404-467-1. K.L. Sutton and J.A. Caruso (Eds.), Elemental Speciation—New Approaches for Trace Element Analysis. Elsevier, Amsterdam, 2000. Ph. Quevauviller and R. Morabito, Trends Anal. Chem., 19 (2000) 86. R. Morabito, P. Massanino and Ph. Quevauviller, Trends Anal. Chem., 19 (2000) 113. Ph. Quevauviller, M. Astruc, R. Morabito, F. Ariese and L. Ebdon, Trends Anal. Chem., 19 (2000) 180.

115

This page intentionally left blank

Chapter 5

Detection methods for the quantitation of trace elements Les Ebdon, Andrew S. Fisher, Maria Betti and Maurice Leroy

5.1

INTRODUCTION

This chapter serves as a general introduction to the methods of trace element determination discussed throughout this book. Brief overviews of these instrumental techniques will be given, along with discussions of their analytical capabilities, requirements of the sample, sample throughput, ﬁgures of merit and descriptions of the numerous methods of sample introduction. In addition, a brief overview of some of the sample preparation methods and sample manipulation procedures will also be given. Included in the chapter are selected examples of applications, although many of these will be treated in more detail throughout later chapters in this volume. It is worth noting that many of the sample introduction methods for the more frequently used atomic spectrometric techniques are common to all. Therefore, the description of the theory behind them will only be given once and, thereafter, applications of each will be given for the other methods of detection. 5.2

CLASSICAL METHODS

Classical methods of analysis will be dealt with only very brieﬂy here since it is largely outside the scope of this chapter. It should be noted, however, that titrations are still an important part of an analyst’s armory because an “EDTA” titration readily provides traceability to a primary standard. Titrations can be relatively time consuming and do not usually offer very great sensitivity, so are of limited use for many sample types. For those applications where the analyte is present at an appreciable concentration (contaminants at the 0.01–5% m/m range in metallic samples, or even the major constituent), a titration can offer very accurate and precise results. As an example, inspection of the certiﬁcate for the reference material BCS 177/2 lead-base white metal (available from the Bureau of Analysed Samples, Middlesbrough, UK) Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

117

L. Ebdon et al.

indicates that lead (present at an average of 84.5%), antimony (10.1%) and tin (5.07%) were all determined by different titrimetric methods. In addition, three analysts determined arsenic (0.05%) by titration. A sound knowledge of the chemistry of both the sample and of the analytical method is required to prevent interferences. A useful textbook covering many of the classical “wet-chemical” methods is “Vogel’s Textbook of Quantitative Chemical Analysis” [1] which contains an assortment of titrimetric, gravimetric, potentiometric, electrogravimetric, spectrophotometric and amperometric methods. As well as giving the basic theory behind each of the techniques, it also gives experimental details for some selected applications.

5.3 5.3.1

FLAME SPECTROMETRY Introduction

Flame spectrometry, either atomic absorption spectrometry (AAS) or atomic emission spectrometry (AES), are amongst the most simple and inexpensive of the instrumental methods of trace element analysis. The cost of a basic AAS instrument can be less than US $10,000, although for the more powerful computer controlled instruments containing autosamplers, the cost can easily be double this. A ﬂame photometer (a very basic AES instrument) that can be used to determine analytes such as lithium, sodium and potassium costs even less. 5.3.2

Theory

A detailed description of the basic theory of AAS and AES is not required here; a detailed and theoretical description of the processes within the ﬂame (or plasma for emission) may be found elsewhere [2]. The relaxation of electrons in an analyte atom from different excited energy levels populated by (thermal) ﬂame processes back to the ground state will yield photons of light of different energy, i.e., the wavelength of the light emitted will be different for each transition. A characteristic spectrum for each element will therefore arise. The transition probability governs the sensitivity of a wavelength (an analytical line). If the probability of a transition is low, then the number of analyte atoms (or ions) in which the electrons are excited to that energy level will also be low. This means that the number of atoms/ions emitting light at the corresponding wavelength will be relatively few and hence the overall sensitivity will also be low. Therefore, to obtain a detectable signal, the concentration of that analyte in the sample will have to be higher. Each line of an analyte, therefore, has a different sensitivity. This can be useful analytically, because each line will have its own linear range. If the expected range of

118

Detection methods for the quantitation of trace elements

concentration of an analyte in a sample is known, then the most appropriate analytical line may be chosen, thereby negating the need for large sample dilutions and hence keeping sample manipulation to a minimum and decreasing the likelihood of dilution errors or contamination. In atomic absorption, the amount of light absorbed from an incident beam of light is proportional to the number of analyte atoms in the optical path, and hence to its concentration in the sample. As for all absorption-based techniques, the path length also has an effect on the sensitivity. Typically, a 10 cm path length is used for an air–acetylene ﬂame, but there is no reason why a smaller path length (5 cm, used for nitrous oxide –acetylene), could not be used, which would lead to half the absorbance of that using the larger burner. This also has the effect of extending the linear range by a factor of two. As in atomic emission, numerous wavelengths are available for each analyte and these will each confer a different sensitivity. A comprehensive theory behind atomic absorption may be found elsewhere [3]. It is worth noting that true spectral interferences for atomic absorption are very rare. This is because of the “lock and key” effect of the incident radiation and the analyte atoms. Theoretically, there are no other atoms present in the atom cell that should absorb the radiation and hence false high signals should not be obtained. Unfortunately, the presence of particulate matter or of some molecules may lead to absorption of the light. Under circumstances such as these, erroneously high signals may then be obtained. 5.3.3

Instrumentation

As discussed above, a light source is required to excite the analyte atoms when using the AAS technique. There are two common types of light source, of which the line source hollow cathode lamp (HCL) is the more frequently used. It has a cup-shaped cathode coated internally with (or fabricated from) the analyte of interest. Often, only one element is used per lamp, but multi-element lamps are also commercially available that may contain two, three or perhaps even ﬁve or six elements. Although multi-element lamps are more expensive than single element ones, they have the advantage of being less expensive than investing in ﬁve or six individual lamps. Their disadvantage is that often compromise operating conditions must be used, which may have an unfavorable effect on the signal-to-noise ratio and linear range for some of the analytes. The other light source commonly used is the electrodeless discharge lamp (EDL). These are more expensive to purchase but offer an increased light intensity and for some analytes, e.g., arsenic, and provide for higher sensitivity and enhanced detection limit. Sample is usually, but not always (see Section 5.3.6), introduced to the instrument as a liquid via a nebulizer/spray chamber assembly. As the gas used for combustion passes the end of a capillary, a pressure drop is obtained. If the other end of the capillary is immersed in a liquid sample, it will be drawn

119

L. Ebdon et al.

through the capillary by the Venturi effect. As it enters the gas stream rushing across the end of the capillary, the stream of liquid is shattered into a nebular (an aerosol or mist of droplets). This nebular then enters a spray chamber where the larger droplets are separated from the smaller ones by a combination of an impact bead (that helps shatter the droplets into smaller droplets) and a series of bafﬂes. The smallest droplets are then carried in the gas ﬂow towards the atom cell whilst the larger ones pass to waste under the inﬂuence of gravity. For ﬂame spectrometry, typically 10– 15% of the sample reaches the atom cell whilst 85 –90% is wasted. Once the sample aerosol enters the atom cell, the ﬂame desolvates the aerosol and then dissociates the salts present into their constituent atoms. The atoms of the analyte will then absorb the light emitted from the light source and the amount of light absorbed can be related to the concentration of the analyte in the sample. There are several ﬂame types that can be used, the most common being a mixture of air and acetylene. These can be mixed in several different proportions, including fuel rich (here a yellow ﬂame that has reducing properties is produced), fuel lean (a blue ﬂame that is chemically oxidizing) or stoichiometric (an intermediate ﬂame that is blue but also has yellow “feathers” at its base). The type of ﬂame chemistry used will depend on the analyte, and should be optimized for every element determined. The temperatures of these ﬂames range between 1700 and 2200 K, which is sufﬁcient to dissociate the majority of compounds. A hotter ﬂame, e.g., nitrous oxide –acetylene (2500 – 2700 K) may be necessary for the more refractory compounds. Again, different proportions of nitrous oxide and acetylene may be required for optimal determination of different analytes. Other ﬂames used, albeit less frequently, include a hydrogen diffusion ﬂame and a methane ﬂame. The former has the advantage of being very optically clean at lower wavelengths, which will improve the signal-to-noise characteristics for wavelengths such as 193.7 nm (As), 196 nm (Se), 213.9 nm (Zn) and 217 nm (Pb). The methane ﬂame is useful when the sample matrix may contain a very high concentration of a component that forms an explosive acetylide compound (e.g., Ag or Cu). The analyte wavelength used for the measurement process is usually isolated with the use of a low-resolution monochromator, since in AAS the resolution of the instrument is essentially derived from the narrow wavelength output of the line source. Once the wavelength of light of interest has been isolated, it may be detected using a photomultiplier tube or solid-state electronic device (such as a charge coupled or charge injection device or diode array). These convert photons to an electrical signal, the magnitude of which may be related to the concentration of the analyte within the sample. Since atomic absorption is a ratio technique, i.e., a comparison is made of the initial light intensity with the intensity after absorption by the analyte has occurred, there are no units. Tuned electronic circuits are used to ensure that light produced by emission

120

Detection methods for the quantitation of trace elements

processes arising from the analyte within the ﬂame do not interfere with the measurement of atomic absorption. Atomic emission spectrometry utilizes all of the above procedures with the exception of the HCL light source. A monochromator is usually used to isolate the wavelengths of interest but very low-resolution optical ﬁlters may be used in the less expensive ﬂame photometers. Analyte atoms thermally excited by the ﬂame emit multiple wavelengths of light, one of which is isolated and detected. Since the advent of inductively coupled plasma instrumentation for atomic emission, the ﬂame emission technique has been in decline and is now rarely used. A more detailed description of the instrumentation used and of the processes occurring within it may be obtained in several other publications [2,3]. 5.3.4

Interferences and background correction techniques

The majority of interferences that are encountered are either physical or chemical in nature, although a few spectral interferences arising from molecular species also exist. Transport efﬁciency of the sample through the sample introduction system can lead to interferences if standards are not closely matrix matched with the samples. For example, differences in viscosity between samples and standards result in different nebulization efﬁciencies. If less of the sample reached the ﬂame, an underestimate of the true concentration of the analytes would be made. If the concentration of the analyte is sufﬁciently high, it may be possible to dilute the sample such that the dissolved solid content becomes negligible. If the analyte cannot be diluted, then it may be necessary to perform a standard additions analysis. Many analysts are not overly keen to use the standard additions technique, because it means that the same sample must be analyzed up to four times with different added concentrations, thereby lengthening the analytical process fourfold. In addition, the volume of sample used will be increased fourfold, which may be problematic if only a limited supply is available. There are several types of interference that may occur in ﬂame spectroscopy. Chemical interferences may cause either depressions or enhancements in the signal, depending on the particular interferent. If a species is present in the sample that will combine with the analyte to form a less volatile compound that is difﬁcult to dissociate in the ﬂame, then a depression in signal may occur, an example being the presence of phosphate during the determination of calcium. There are various methods for overcoming this, including adjusting the nebulizer so that smaller droplets are produced; making observations higher in the ﬂame so that the less volatile compound has a longer time to become dissociated; using a releasing agent (e.g., lanthanum), that preferentially combines with the phosphate, and using a chelating agent (e.g., EDTA) to complex with the analyte so that it cannot combine with the phosphate. The addition of other chemicals to the samples may lead to

121

L. Ebdon et al.

contamination and error, so possibly the most simple and reliable method of overcoming this type of interference is the use of a hotter ﬂame, i.e., nitrous oxide–acetylene. Signal depressions may also occur if the analyte becomes occluded in a refractory compound, such as is the case of the oxides of rare earth elements, uranium or zirconium. Again, the use of a hotter ﬂame usually overcomes this problem. Signal enhancements are much rarer, but an example is the formation of an EDTA complex for calcium. The complex is more volatile than many other calcium compounds. Matrix matching may overcome this, but the use of a hotter ﬂame will ensure that all of the analyte in both standards and samples is atomized. Similarly, if an analyte is occluded into a volatile compound (e.g., ammonium chloride), the atomization of the analyte may be enhanced. Again, matrix matching usually provides a solution to the problem. Ionization interferences occur mainly for the alkali metals that have an exceptionally low ﬁrst ionization potential (IP). Since ﬂame spectroscopy usually determines atoms (either by atomic absorption or by atomic emission), the formation of ions may lead to problems because these will not absorb or emit at the same wavelength. If, for example, sodium is to be determined by either atomic absorption or emission, it is usually necessary to add a high concentration of another easily ionized element such as potassium or cesium (assuming that these are not amongst the analytes), to the samples, standards and blanks. These will become ionized in the ﬂame, producing a large excess of electrons that then force the ionization equilibrium to favor the neutral analyte species. Samples are more likely to contain other easily ionizable elements than pure aqueous standards, and therefore the extent of ionization will be less than for the standards. Unless a large excess of ionization buffer is added to all samples, standards and blanks, an overestimate of the analyte concentration could result. Spectral interference, caused by direct line overlap, is negligible for ﬂame spectroscopy. However, since molecules exhibit a much wider wavelength band of absorption/emission, these can occasionally prove to be problematic. Examples include phosphate and sulfate interferences on the arsenic and selenium lines at 193.7 and 196 nm, respectively. Similarly, small particulate matter within the ﬂame may attenuate the light beam, leading to an erroneously high signal unless it is corrected for by a method of background correction. There are several types of background correction systems used for AAS. These include the deuterium lamp (continuum source), Zeeman effect and Smith-Hieftje systems, and each has been available in commercial instrumentation. Each background correction system has its own relative advantages and disadvantages. A detailed description of their operation is not necessary here, but may be found in the literature [3]. However, a brief description of the advantages and disadvantages of each is appropriate. The deuterium lamp has relatively ineffective output above 350 nm and so the most sensitive resonance line for chromium (at 357.9 nm) may occasionally be problematic if particulate

122

Detection methods for the quantitation of trace elements

material is present in the ﬂame. The deuterium lamp is, however, fairly inexpensive to purchase and to operate and provides an adequate correction for those analytes that have a primary wavelength in the UV region. Since the large majority of background absorption phenomena occur below 350 nm, it is adequate for most applications. Another disadvantage of this continuum source system is that the beam from this light source must follow an identical optical path to the beam from the HCL. The Smith-Hieftje correction system operates on the principle of self-reversal, i.e., if the source HCL is pulsed to much higher operating current, the narrow emission line proﬁles are broadened and suffer some self-reversal. The analyte atoms absorb only a fraction of this broadened line radiation whereas the background absorption is unaffected. By operating the HCL at normal current (2 –25 mA) and at much higher current (. 100 mA) in a rapidly oscillating manner, estimates of the total absorbance (atomic and background) and the background absorbance may be made by subtracting the two absorbance signals. The technique works fairly well for many types of interference, but has several drawbacks. These include the shortened lifetime of the HCL, the “assumption” that the atomic absorption during the high current pulse is negligible, which leads to reduced analytical sensitivity, and increased curvature of the calibration curve. The Zeeman background correction system is used almost exclusively for electrothermal AAS (ET-AAS), but will be included here amongst the other background correction techniques for completeness. It is a technique that uses a powerful magnet (approximately 1 T) to separate the normal single atomic line proﬁle into several different components, as described in more detail elsewhere [3,4]. The signiﬁcant advantage of this type of correction system is that it is capable of correcting much larger background signals than any of the other methods. Unfortunately, it also suffers from decreased sensitivity and increased curvature (and ultimately complete roll-over) of the calibration function. 5.3.5

Conventional nebulization

The process by which conventional nebulization occurs and some of the potential problems that may arise (i.e., different viscosity of samples and standards leading to different nebulization efﬁciency and hence, different sensitivity) have been described previously and several other factors need to be discussed. The sample uptake rate for conventional nebulization in a typical ﬂame AAS/AES instrument is between 4 and 8 ml min21. This may usually be changed by careful adjustment of the nebulizer. Fortunately, the time required for the analyte to pass through the nebulizer/spray chamber system, into the burner head and then into the ﬂame, is only about a second. Therefore, measurements may be made only 2–5 s (depending on the integration time used) after sample introduction commences. The amount of sample consumed will depend on the number of replicate readings taken, but usually, a volume of 3–4 ml is adequate to determine an analyte. However, for most instruments,

123

L. Ebdon et al.

ﬂame AAS is a single element technique, i.e., the operating conditions will have to be changed for another analyte, 3–4 ml of sample is required for every analyte. This may prove problematic if 10 –20 analytes need to be determined and only 25 ml of sample is available. It should be noted, however, that some modern instruments have a rapidly scanning monochromator (2000 nm min21) and specialized valves that enable a very rapid change of the ﬂame chemistry. These attributes, combined with a series of fast rotating mirrors to change between different HCLs, enable very rapid sequential determinations to be made. Such an instrument offers signiﬁcant improvements in analysis time, although sample consumption may be marginally higher. Conventional nebulization into ﬂame AAS is, as discussed above, very rapid. After the measurement of one sample or standard, a washout period is necessary in which water, dilute acid or a chemical matched to the matrix of the samples is used to ensure that there is no signal carry over between samples. Depending on the matrix and the analyte, this requires anywhere from just a few seconds to in excess of a minute. The washout may have to be especially long if the sample contains a very high concentration of dissolved salts. Once these samples enter the burner head, they will desolvate and, unless a long wash period is used, there is a chance that the salts will start to block the burner head. This would result in several effects, including a reduction in the sensitivity (because if gaps start appearing in the ﬂame, the path length is effectively decreased) and excessive signal drift. Occasionally, the burner may be cleaned by gentle scraping with a non-combustible item, e.g., a stainless steel spatula but often, ﬂame extinction followed by dismantling of the burner assembly is necessary to clean it. This would obviously lead to an increase in the analysis time. A single element may be determined in only a few seconds per sample, and therefore a batch of 20 samples could be analyzed for one analyte in less than 10 min. Using the rapid sequential instruments, sample throughput for several analytes can be improved signiﬁcantly. Many modern instruments come equipped with an autosampler, which facilitates the unattended operation of the instrument, thereby maximizing sample throughput with minimal human intervention. Many modern instruments have software that enables the complete analysis to be pre-programmed, incorporating assorted quality control measures, e.g., check standards, collection of data from multiple replicates and the calculation of mean values, standard deviations and precision, etc. Some instruments have a moveable turret in which three or four HCLs may be inserted. The software then controls the monochromator, changing to the wavelength necessary for each analyte. The ﬁgures of merit of ﬂame techniques tend to be the least impressive of the standard instrumental techniques, but are still adequate for many applications. The limit of detection will depend on several factors, including the analyte itself. Some analytes, e.g., magnesium or cadmium, are extremely sensitive, whereas others, e.g., lead, are not. Other analytes that are extremely refractory, such as tantalum and tungsten, offer relatively poor sensitivity.

124

Detection methods for the quantitation of trace elements

The limit of detection will also depend on which analytical line of the analyte is being measured. As described previously, numerous analytical lines may be used and each has a different sensitivity, LOD, calibration range, etc. The LOD obtained using each of the lines will also depend on the instrumental parameters used, as each element has an optimal ﬂame chemistry, HCL current, spectral bandpass, viewing height, etc., and unless the optimal conditions are used, the optimal ﬁgures of merit will not be obtained. A list of limits of detection for numerous elements is given in Table 5.1, for which it has been assumed that the most sensitive analytical line is used for each analyte under optimal conditions. It should be noted that the ﬁgures given in Table 5.1 relate to liquid samples. If a solid has been dissolved or digested, then a dilution has occurred and the LOD related to the solid would have to be re-calculated. For atomic absorption measurements, the linear range usually spans 1.5 –2 orders of magnitude before there is a departure from linearity. Therefore, assuming that several standards are prepared that cover this range and that steps have been taken to overcome potential interferences, reliable data should be obtained. Any sample that has an absorbance greater than the most concentrated standard should be diluted so that it comes into the working range. If many or all samples contain a concentration of an analyte that is above the most concentrated standard, it would be less time consuming to use an alternative wavelength and prepare a more appropriate calibration range than to dilute perhaps 50 samples. An alternative method is to rotate the burner head slightly. This has the effect of decreasing the path length, i.e., fewer atoms are in the light beam at any one instant, and therefore the linearity may be extended (at the expense of sensitivity). In such a case, it will still be necessary to prepare another standard that contains an analyte concentration closer to that expected in the sample to ensure linearity. The precision expected from a ﬂame instrument (as with any instrumental method) will obviously depend on the concentration being measured. If the concentration is close to the LOD, then precision will be poor when compared with a concentration further up the linear range. For the latter example, a precision of 0.1 –2% relative standard deviation (RSD) is typical. As discussed previously, the majority of samples introduced via conventional nebulization must be liquid based and that the transport efﬁciency is usually between 10 and 15% for aqueous based samples. This ﬁgure will depend, however, on the nature of the sample. The presence of appreciable amounts of dissolved solid is likely to decrease this value. If the sample is present in an organic solvent, then the nebulization characteristics will differ markedly. Water has a fairly high surface tension and viscosity and a low vapor pressure. Organic solvents tend to have a lower surface tension and viscosity and a higher vapor pressure. This means that they more efﬁciently form an aerosol, resulting in an increased transport efﬁciency to the ﬂame and enhanced sensitivity. It is therefore extremely important to prepare standards in the same solvent as that used to dissolve the samples. The presence of organic solvents in

125

L. Ebdon et al. TABLE 5.1 Limits of detection using ﬂame AAS under optimum conditions with the most sensitive line Analyte

LOD (mg l21)

Analyte

Ag Al As Ba Be Bi Ca Cd Co Cr Cs Cu Dy Er Eu Fe Ga Gd Ge Hf Hg Ho In Ir K La Li Lu Mg Mn

2 30 300 20 1 50 1 2 5 6 4 3 30 50 1.5 6 100 2000 200 2000 200 40 40 500 3 2000 2 300 0.3 2

Mo Na Nb Nd Ni P Pb Pd Pr Pt Rb Ru Sb Sc Se Si Sn Sr Ta Tb Te Th Ti Tl Tm U V W Y Zn

LOD (mg l21) 20 0.2 2000 1000 10 40,000 10 10 10,000 100 10 100 40 50 500 300 100 2 2000 700 30 100 20 20 40,000 100 1000 200 1

the ﬂame will radically change the ﬂame chemistry and it is therefore important to optimize the ﬂame conditions using the solvent of interest if optimal sensitivity is to be obtained. Organometallic standards, which are often based on the cyclohexylbutyrates, are available commercially and are soluble in many organic solvents. If the solvent is methanol, then many inorganic standards that are stabilized in nitric or hydrochloric acid are soluble.

126

Detection methods for the quantitation of trace elements

Although the large majority of samples introduced into ﬂame atomic spectrometric instrumentation is liquid based, the introduction of solid materials is possible. This is usually achieved through the preparation of slurries. A slurry is a suspension of a very ﬁnely ground sample in a liquid medium which is usually a dispersant to prevent the particles from ﬂocculating. The subject will be dealt with in far more detail in a later section (Section 5.3.6.6). Brieﬂy, the sample is ground so that the particle size is equivalent to the droplet size in the aerosol formed by the nebulizer. The transport efﬁciency of the slurry particles should therefore be equivalent to aqueous standards and therefore, simple aqueous standards (or standards matrix matched with the dispersant used for the slurry) can be used for calibration. A plethora of other pre-concentration and matrix separation techniques, including solid phase extractions, liquid –liquid extraction, co-precipitation, ﬂotation and evaporation are available and these will be discussed in more detail in other chapters throughout the book. 5.3.6

Alternative methods of sample introduction

There are a number of alternative methods that may be used to introduce samples for ﬂame spectrometry. Some of these offer increased sensitivity and others help overcome potential interferences, thereby yielding more reliable results or better long-term stability. 5.3.6.1 Chemical vapor generation Chemical vapor generation as a method of sample introduction is discussed in detail by Cai in this volume. The topic has also been reviewed by Tsalev [5] and by Howard [7]. Although primarily applicable to elements such as arsenic, selenium, antimony, tellurium and germanium, which are capable of forming gaseous hydrides at room temperature by reaction with sodium tetrahydroborate, and elemental mercury, it has been reported that several other analytes, including Ag, Au, Cd, Co, Cu, Ni, Sn and Zn, have also been determined by vapor generation [6]. There are also alternative reagents that may be used to form volatile vapors, including various salts of tetraethylborates. Mercury may be reduced to its elemental state by stannous chloride. There are several advantages of introducing analytes as a gas rather than as a liquid. The ﬁrst is that the analyte is separated from the bulk of the matrix. Together with higher sample uptake rates this means that spectroscopic interferences are minimal. Also, gases are more easily transported than liquids, and hence the transport efﬁciency of the vapors to the atom cell is closer to 100 than 10– 15% obtained for liquids. This will obviously lead to a sensitivity improvement by a factor of 30–50. Frequently, the atom cell is a quartz T-piece placed on top of the burner head that is heated by the ﬂame and the light beam from the HCL passes

127

L. Ebdon et al.

through it. As the gaseous analytes enter the heated T-piece, the compounds dissociate, forming analyte atoms. The advantage of using the T-piece rather than allowing the analyte vapors to simply enter the ﬂame is that it acts as a sort of trap, increasing the analyte residence time in the optical path. Further, the T-piece may also provide a longer absorption path length (15 cm). There are disadvantages associated with vapor generation techniques. Only relatively few analytes form gaseous compounds at room temperature and not all of the oxidation states react with the same efﬁciency. Arsenic in its þ3 state forms a hydride far more efﬁciently and with a different sensitivity than does As(V). Similarly Se(IV) forms a hydride with relative ease whereas Se(VI) does not form a hydride at all. In addition, when the analyte is an integral part of an organic molecule, e.g., selenium in the form of selenomethionine or arsenic as arsenobetaine (AsB), a hydride is not formed. As such, an underestimate of the total concentration of the element of interest will be obtained unless steps are taken to transform all species of the analyte into a state that will form a hydride. There have been numerous chemical and physical methods used to accomplish this, including the use of L -cysteine to reduce As(V) and monomethylarsonic acid (MMAA) and dimethylarsinic acid (DMAA) to As(III) [8], alkaline persulfate to oxidize arsenobetaine [9], iodide/ iodate reactions [10], etc. Included in the physical methods used are photolysis [9] and the use of microwave energy to accelerate the action of acids [11]. Another potential problem with the technique is the presence in the samples of transition metals such as zinc, copper and iron and of precious group metals such as gold, palladium and platinum. These elements interfere with the hydride formation process and often result in an underestimate of the analyte’s concentration. These potential interferences may be overcome by the addition of a chelating agent, such as 1,10-phenanthroline [12], 8 hydroxyquinoline [13] or picolinic acid [14]. Limits of detection for the vapor generating analytes can be improved by a factor of over 100 compared with their conventional nebulization, with LODs for many of the analytes being at the low ng ml21 level. Precision should again be at the 0.5 –3% RSD level. Sample consumption will depend on the mode of vapor generation. In the continuous mode, a typical analysis is likely to use 10– 12 ml of sample for a measurement time of approximately 30 s. This type of operation is therefore slightly more wasteful of sample than conventional nebulization, but sensitivity is improved still further. The other mode of hydride generation is the “batch” mode. Here, a discrete volume of sample is used and the signal will appear as a transient, i.e., a peak. This method uses far less sample, although several injections will have to be performed so that an estimate of precision can be made. The other drawback is that it may be necessary to have a chart recorder or integrator output to the spectrometer so that measurement of the peak height or area may be performed more accurately. Sample throughput for hydride generation introduction to ﬂame

128

Detection methods for the quantitation of trace elements

spectrometry is less than for conventional nebulization, but it should be possible to analyze 20–30 replicates in an hour. 5.3.6.2 Sampling cups and ﬂow injection If a very limited sample volume is available, e.g., , 2 ml, it will probably not be possible to determine any more than one analyte if conventional nebulization is used. Also, if a sample contains a very high concentration of dissolved solids, there is a chance of both nebulizer and/or burner head blockage. Sampling cups and ﬂow injection (FI) methods are both means of introducing discrete volumes of sample, thereby decreasing the volume of sample introduced and hence the amount of dissolved solid entering the instrumentation. As with the batch mode of vapor generation, a transient signal is obtained. For maximum signal to be obtained, an injection volume of approximately 0.5 ml is required, but smaller volumes may be introduced with a concomitant drop in response. This occurs because approximately 0.5 ml is the minimum volume required to obtain a signal equivalent to that generated with conventional nebulization. The overall result is that 0.5 ml injection volumes are likely to lead to improved precision when compared with smaller volumes. The sampling cup is a device that has a small hole in the bottom of a cup of volume of approximately 1 ml. The nebulizer uptake tube is inserted into the hole, so when sample is dispensed into the cup via a high accuracy and high precision micropipette, it is immediately aspirated into the ﬂame. In between sample replicates, several volumes of water or dilute acid may be injected to ensure that no carry over effect occurs. The use of a micropipette to introduce the sample is a potential source of imprecision, since a worn seal will prevent reproducible volumes from being taken up and dispensed. The method of sample introduction using the sample cup is also called pulse or gulp nebulization. Numerous FI methods have been reported and an overview of the relevant literature from 1972 to 1995 has been presented by Fang et al. [15], with a current treatment available in this volume. The simplest of FI methods requires just a sample injection valve to be coupled to the nebulizer uptake tube, permitting discrete volumes of 0.01 ml upwards to be introduced via a sample loop, although sample introduction via direct injection with a syringe is also possible, but this leads to poorer precision. Flow injection frequently makes use of mini- or micro-columns of an ion exchange or chelating resin to retain the analytes of interest and eliminate or minimize concomitant element interference effects. Pre-concentration may also be readily achieved using FI techniques and will improve the LOD for ﬂame detection (and any other technique) considerably. The pre-concentration factor achievable by FI techniques will depend on the analyte, the sample volume available, time constraints and, in some cases, by the purity of the chemicals used for buffers, etc. Time constraints must also be considered. If the sample is pumped through the column at 3–4 ml min21, it will still take 25 –30 min to introduce 100 ml.

129

L. Ebdon et al.

The volume and concentration of eluent required will have to be optimized, but typically 0.25– 1.0 ml is used. Therefore, large pre-concentration factors are possible theoretically, but it will be at the expense of time. Inevitably, a busy laboratory with 100 samples to analyze cannot afford to take 30 min per sample replicate; especially when typically three replicates per sample are required so that an estimation of precision can be made. In practice, pre-concentration factors of 10 –40 are more common. The precision of FI techniques that use a column of resin to afford matrix removal/pre-concentration will depend on the reproducibility with which the analyte is retained and then eluted from the column. Successful methods have a precision of typically , 5% RSD. Flow injection techniques that simply use a valve to introduce small volumes of sample into the spectrometer should have a precision of 1–2% but, again, this will depend on the concentration of the analyte within the sample. Sample throughput for sampling cups would typically be 60 samples (assuming three or four replicates) per hour. Flow injection methods tend to be slower, but simple FI methods may analyze 20 –30 samples per hour. Methods involving matrix separation/pre-concentration are the slowest and will depend on the pre-concentration factor, but 5–10 samples per hour is typical. 5.3.6.3 Slotted tube atom trap (STAT) The STAT acts in a very similar way to the quartz T-piece used for vapor generation. The tube is placed on the burner head, ensuring that a slot carved into the side of it is directly over the ﬂame slot in the burner head; a smaller slot on the top and/or ends of the tube allow exit of ﬂame gases from the tube. The ﬂame ensures the tube then acts as a heated atom trap. The analyte molecules introduced via conventional nebulization, or often by FI, then enter the tube through the slot, become thermally dissociated into atoms and atomic absorption occurs. An increase in sensitivity by a factor of three- to ﬁvefold is obtained. Precision has also been found to improve through the use of a STAT. A review of the atom trapping procedures in ﬂame spectrometry has been presented by Matusiewicz [16]. Closely related to the STAT is the application of a water cooled atom trap (WCAT) that consists of a water cooled single or dual silica tube suspended in the ﬂame which serves as a condensation site for atoms/molecules introduced into the ﬂame. Following a suitable collection period, the water cooling is terminated by use of a pulse of gas through the tubing, which then rapidly heats to ﬂame temperature and results in volatilization of the collected analyte. Typical 2-minute collection periods can improve the detection limit by an order of magnitude, but the technique is clearly most favorable for volatile elements such as Ag, Cd, Cu, Zn, Pb and Tl. Recently, the STAT and the WCAT have been combined in a synergistic arrangement and used for sample analysis [17]. In general, the sample throughput and sample consumption will be governed by whatever sample introduction method is used.

130

Detection methods for the quantitation of trace elements

5.3.6.4 Chromatography Simply determining the “total” concentration of an analyte does not give any information on the overall toxicity of the sample. Therefore, speciation analysis, the determination of different forms of the analyte, is becoming increasingly frequent. One of the most common methods of achieving speciation analysis is to couple a separation technique, e.g., high performance liquid chromatography (HPLC) or gas chromatography (GC), with an element speciﬁc detector. Flame spectrometry is one of the least sensitive methods of atomic spectrometry and therefore the number of speciation analyzes that may be performed with it is somewhat limited. However, despite this obvious drawback, a large number of applications have been presented. Several reviews of chromatography coupled with ﬂame spectrometry have been published. These include those by Ebdon et al., who covered the earlier literature for liquid [18], and GC [19], and a more recent one by Szpunar Lobinska et al. [20]. Several speciation approaches are discussed in detail in other chapters of this book. Gas chromatography depends on the analyte being volatile. If the analytes are not naturally volatile, it may be necessary to resort to use of derivatization reactions, such as use of a Grignard reagent [21]. For HPLC couplings, the end of the column may simply be attached to the nebulizer uptake tube. The ﬂow rate through the chromatography column is typically 1– 2 ml min21, which is less than the natural uptake rate for the nebulizer. It may therefore be necessary to insert a small air bleed to compensate for this mismatch [22]. For GC couplings, it is usually necessary to utilize a heated transfer line from the GC oven to the atom cell. The end of the heated transfer line is usually placed in one of the slots of a STAT so that extra sensitivity is obtained. Sample throughput will depend largely on the chromatographic stage. Both liquid and gas chromatograms frequently take in excess of 10 min per sample, and so sample throughput is very limited. Instrumental precision will depend on the method of sample introduction. If a sample loop is used for HPLC, then precision should be less than 5% RSD. If a syringe is used to inject 1–10 ml into a gas chromatograph, then precision can be . 10%. This may be improved substantially if an internal standard is used. For speciation techniques, this is normally a compound that has similar properties to the analyte compounds, but is not found naturally in the sample. Ideally, the internal standard should elute in the middle of the chromatogram, and not co-elute with any of the species of interest. Another potential source of error, inaccuracy and imprecision, is the extraction technique used to remove the analyte species in an unchanged state from solid samples. This topic will be discussed in later chapters, but it is worth noting here that an inadequate extraction method (one that changes the speciation or that does not yield reproducible recovery, etc.) will render the entire analysis irrelevant. One point that is worth noting for all speciation analyzes is that the concentrations quoted should specify whether the values are related to the

131

L. Ebdon et al.

concentration of the species or of the analyte element. For instance, if during the analysis of ﬁsh extracts a concentration of 5 mg kg21 is quoted, the report should specify whether this is 5 mg of arsenic kg21 or 5 mg of arsenobetaine kg21, etc. Failure to do this is likely to lead to confusion and error. Similarly, LODs should also be quoted with the same qualiﬁcations. 5.3.6.5 Multiple couplings Occasionally, when extra sensitivity is required for speciation analysis and when instrumental costs preclude the purchase of a more sensitive detector, it is necessary to couple together several techniques. A technique that has become relatively common (in the research literature) is the coupling of HPLC with HG-AAS [23,24]. After the species have been separated using HPLC, either a chemical or physical process is used on-line to convert each of the species to a state that will form a hydride and may require a chemical oxidation using alkaline persulfate or photolysis. In the example given in Ref. [23], on-line microwave assisted oxidation yielded LODs of 2.5, 5.3, 3.3 and 5.9 ng ml21 AsB, DMAA, MMAA and As(V), respectively. 5.3.6.6 Slurries As discussed earlier, a slurry is a suspension of solid sample in a liquid medium. The advantages of slurry sample introduction include the ease of preparation, the non-requirement of powerful reagents such as hydroﬂuoric acid, no possibility of losing volatile elements and, for most analytes, minimal contamination. Fuller et al. [25] reported very early on the relative merits of ﬂame, electrothermal and ICP atomization techniques for the direct analysis of slurries. Several slurry preparation techniques have been reported, but the fundamental necessity is that it be representative of the sample, i.e., be homogeneous. This usually means that the powdered sample must be ground using either the bottle and bead method or in a micronizer. In the bottle and bead method, a sub-sample is placed in a small plastic bottle, a small volume (e.g., 5 ml) of aqueous dispersant are added together with 10 g of zirconia beads (2 mm diameter) and then the bottle is sealed and placed on a mechanical ﬂask shaker for a period of time that is dependent on the sample type. Blanks are prepared in the same way, but omitting the sample. The drawback with the technique is that the blanks tend to give a “worst case scenario”, because the beads have no sample to cushion the impact of the collisions between them during the grinding process. This means that the beads will grind themselves to a greater extent than when the sample is present. The concentration of the contaminants in the blank is therefore often slightly larger than that found in the samples. The process is, however, suitable for the determination of a great number of analytes, with the obvious exceptions being zirconium and hafnium (which is often a substantial contaminant in the beads). The micronizer uses agate rods to grind the sample instead of zirconia beads. This will give rise to a different set of contaminants (i.e., Na, Mn, Si, etc.). The choice of which

132

Detection methods for the quantitation of trace elements

grinding procedure to use will therefore, depend on the analytes to be determined and on the nature of the sample. The zirconia beads are fairly hard [measure of hardness (MOH) ¼ 8 þ ], whereas the agate rods are softer ðMOH ¼ 7Þ: If a particularly hard sample is to be slurried, then the zirconia beads are a more suitable grinding medium because otherwise the agate rods may end up actually being ground by the sample. For exceptionally hard samples, it may be necessary to use a tungsten carbide swing mill to affect grinding. After grinding is complete, the beads (or agate rods) may be removed by simple ﬁltration through a coarse Buchner funnel, without the presence of a ﬁlter paper. The beads may then be washed with more dispersant and the washings collected and combined with the sample. Apart from the problems associated with insufﬁcient grinding and contamination, another problem is that some samples are very soft and have a tendency to become squashed or ﬂattened during the grinding process rather than being smashed into smaller fragments. The overall effect, therefore, is that particle size is not reduced sufﬁciently. This problem is more common with organic based samples such as plants, etc. The dispersant used will depend on the nature of the sample. For inorganic matrices such as soils, rocks, ceramics and other refractory materials, sodium hexametaphosphate or sodium pyrophosphate is suitable. For more organic based samples such as plant material, blood, food samples, etc., then Triton X-100 or aerosol OT are more appropriate. In either case, it is necessary to inspect the ground sample under a microscope to ensure that the particles are sufﬁciently dispersed, i.e., they have not ﬂocculated together. If sample particles do ﬂocculate together to form an agglomeration, then they will act as a much larger particle and the slurry will no longer be homogeneous. Slurries may be aspirated into either ﬂame [26] or plasma-based instruments, introduced to ET-AAS instruments or they may even be analyzed using a hydride generation technique. If the slurry is to be aspirated into a ﬂame or a plasma via a conventional nebulizer/spray chamber assembly, it is necessary to ensure that the particle size is extremely small and that, ideally, the particle size distribution covers only a small range. If this is the case, then the sample particles will act in a similar manner to aerosol droplets, enabling calibration against standards prepared in the aqueous dispersant. If the slurry particles are too large, the nebulizer and spray chamber select against them and they will preferentially be passed to waste so that the sample that reaches the atom cell is not representative of the whole, leading to inaccuracy and poor precision. In addition, the larger the particle size, the more difﬁcult it will be to ensure homogeneity, i.e., even if the slurry is stirred, the larger droplets will sink to the bottom of the container at a faster rate than the smaller ones. The fundamental parameters required for slurry nebulization into plasmas were discussed in a paper by Goodall et al. [27]. These authors determined that an upper particle size diameter of 2.0 –2.5 mm was necessary for accurate results to be obtained, but the maximum particle size that yielded accurate results was

133

L. Ebdon et al.

also dependent upon the sample density. For a sample having a density of 1 g cm23, a particle size of 2.9 mm could still be transported efﬁciently to the plasma; however, for a sample with a density of 7 g cm23, the size had to decrease to 1.5 mm. For very refractory samples, even if the particle size of the slurry is sufﬁciently small to pass through the nebulizer/spray chamber assembly and reach the plasma, complete dissociation may not occur and an inaccurate concentration value will be determined. To overcome such problems, alternative gases have been used. Ebdon and Goodall [28] introduced hydrogen to the nebulizer gas ﬂow to yield more accurate results when slurries of refractory certiﬁed reference materials (CRMs) were analyzed. This was attributed to the increased thermal conductivity of the hydrogen improving the energy transfer from the toroidal part of the plasma to the annulus, thereby increasing the rotational temperature and, hence, improved dissociation of the particles. A review of slurry nebulization into plasmas has been prepared by Ebdon et al. [29]. Even when the slurry is to be analyzed by ET-AAS, sample homogeneity must be maintained. Since the particulate material of slurries will settle with time, it is necessary to agitate the slurries vigorously to ensure complete homogeneity before the sample is introduced. Failure to do this will lead to exceptionally poor precision and accuracy. Therefore, hand-held pipettes are often used so that sample introduction takes place immediately after homogenization. The introduction of slurries using an autosampler is a possibility provided that there is a mechanism by which homogeneity is ensured. Miller-Ihli developed an ultrasonic probe that is inserted into the autosampler cups to mix the slurry and hence ensure homogeneity [30]. Using such a device, the analysis of slurries can become completely automated. A review of the slurry sampling for ET-AAS applications between 1990 and 2000 has been presented by Cal-Prieto et al. [31]. For many slurry types and some biological liquids such as blood, it may be necessary to introduce air or oxygen during the pyrolysis stage to ensure a more efﬁcient oxidative combustion process that decomposes the organic material more efﬁciently and hence helps reduce interferences arising from smoke. In addition, for the blood samples, it will also prevent the build-up of a carbonaceous residue that will, in time, start to obscure the light beam. If a reactive gas is introduced during the pyrolysis stage, it is normally necessary to use a second pyrolysis stage with just an inert gas passing through the tube to remove all traces of the air before atomization. Failure to do this will lead to accelerated tube wear. Precision for slurry analysis by ET-AAS is dependent upon the homogeneity of the slurry, but could be as low as 3–5% RSD. Occasionally, the slurry will be mixed with nitric acid or some other reasonably strong reagent to help leach the analytes from the solid matrix into the liquid phase. This will often have the effect of increasing accuracy of the analysis, because some of the analyte is in solution and will therefore act in

134

Detection methods for the quantitation of trace elements

a similar manner to the standard. Additionally, the particle size is likely to be decreased, enabling more efﬁcient transport of these to the atom cell. An example of a procedure that has used acid leaching as an aid to slurry nebulization into inductively coupled plasma-mass spectrometry (ICP-MS) has been published by Persaud et al. [32]. If the sample is biological in origin, the analytes may well be at a low concentration. It is sometimes possible to place the sample in a mufﬂe furnace and then char it at 4508C for several hours until only ash remains. This ash may then be slurried in the normal way. Such a pre-treatment will enable an effective pre-concentration to be achieved since, on ashing, many biological samples will lose 90% of their mass and hence a larger quantity of sample may be introduced before problems associated with excessive amounts of dissolved/suspended solids occur. The usefulness of the dry-ashing pre-concentration technique is, however, analyte selective and will be inappropriate for very volatile analytes such as mercury, cadmium and possibly lead and zinc. 5.4 5.4.1

ELECTROTHERMAL AAS Introduction

Electrothermal AAS shares the same fundamental principles as ﬂame AAS, the major differences being the atom cell and the method of sample introduction. In ET-AAS, the liquid sample is dispensed into a graphite tube, which is heated resistively, undergoing a temperature programme that ﬁrst dries the sample, pyrolyzes it so that as many matrix concomitants (i.e., potential interferences) are removed as possible, and then heats it to a temperature that is sufﬁciently high to vaporize and atomize the analyte so that it can absorb the HCL light beam. There is then usually a cleaning stage to prevent analyte carry over between samples. The temperature of each of these stages is dependent upon the analyte of interest. The drying temperature should be sufﬁcient to ensure smooth evaporation of the solvent. If it is too high, the sample may froth and spit out of the tube, decreasing precision. The pyrolysis temperature should be high enough to remove as many interferences as possible, but not too high so that the analyte is lost through volatilization. This temperature can range between , 2508C for mercury through to 17008C for very refractory analytes such as erbium. The atomization temperature should be sufﬁcient to ensure complete atomization of the analyte whilst not being excessively high so as to cause accelerated tube wear. A temperature between 1200 and 28008C may be used, depending on the analyte and on the capability of the instrument. The tube is protected from atmospheric oxidation by purging the entire system with argon, although nitrogen may be used for some analytes, excepting those that form a refractory nitride. The gas usually surrounds the graphite tube and ﬂows at a rate of 1–3 l min21. Many modern instruments also have a ﬂow of argon (200 –300 ml min21) internally through the tube to aid

135

L. Ebdon et al.

the ﬂushing of smoke and solvent vapor from it. This internal ﬂow is normally switched off during atomization to prevent dilution of the analyte atoms and from ﬂushing the atoms out of the light beam too rapidly. The speed of analysis is much poorer compared with ﬂame AAS, as a typical ET-AAS temperature programme can exceed 2 min and frequently three replicates will be analyzed per sample. It is therefore unlikely that many more than 10 samples can be analyzed per hour. The initial cost of the instrumentation is substantially higher than simple ﬂame spectrometers, with the least expensive electrothermal (also called graphite furnace) instruments being double the price. More complex instrumentation will cost much more. The running costs also tend to be higher, with the graphite tubes costing up to US $70 each, as well as the supply of argon gas. The lifetime of the tube depends largely on how corrosive the sample is and what analytes are of interest (i.e., how high the atomization temperature needs to be). The advantages of using ET-AAS rather than many other detection techniques include the requirement of only a very small volume of sample. Typically, other instrumentation requires at least 0.5 –1 ml of sample unless dilution is performed (which may put the analyte below the LOD of the technique) or unless specialized sample introduction methods are used, e.g., pulse nebulization, etc. The typical injection volume for ET-AAS is 10 –30 ml, so even if triplicate measurements are made, less than 100 ml would be sufﬁcient. In addition, the sensitivity is 100 –1000 times superior to ﬂame AAS and for many elements it is also superior to inductively coupled plasma-atomic (optical) emission spectrometry (ICP-OES). 5.4.2

Conventional ET-AAS

There are several types of tube available commercially but most are manufactured from some type of graphite (although there are a few applications where metal atomizers have been used). Many of the applications of this latter type of atomizer have been reviewed by Nobrega et al. [33]. Of the graphite-based tubes, electrolytic (electro)graphite is the least expensive of the materials, but is very porous and samples can soak into the graphite lattice leading to interactions between the graphite and the analytes. For analytes such as chromium and other refractory carbide forming elements, this can be problematic. Pyrolytic graphite is much less porous (more dense) and is far less reactive than electrographite. Therefore, there is less interaction between the tube and the analytes and the lifetime is extended. The tube can either be coated with pyrolytic graphite or some may even be manufactured totally from it. There are also several different styles of tube available commercially. Some are heated longitudinally from the ends, leading to a temperature gradient along the tube with the middle being hottest. This is not a favorable scenario, since the analyte may be atomized from the hot central region of the tube and

136

Detection methods for the quantitation of trace elements

then condense at the cooler ends. Other tubes are heated transversely from the sides and do not exhibit a temperature gradient. Slightly lower atomization temperatures may be used without fear of condensation problems. Some tubes have an in-built platform onto which the sample is introduced. The platform ensures that the sample is heated both by the hot internal gas (convectively) and radiatively rather than by conduction from the tube walls, facilitating a process known as isothermal (or stabilized temperature) operation. This leads to fewer interference problems (these will be discussed in a later section). Sample is usually dispensed as a liquid into the graphite tube or onto the platform. An autosampler can dispense the sample to the same part of the graphite tube in a more reproducible way than a hand-held pipette. If the sample is placed in the same place more reproducibly, better precision should be obtained, especially for longitudinally heated tubes. Ideally, the sample should be dispensed from the same height each time, so that the sample drop is not disturbed by the autosampler introduction arm. If it were disturbed, it may spread over a larger area of the tube, again leading to impaired precision. In general, precision obtained with ET-AAS determinations are 1–3% RSD if an autosampler is used and 3–5% RSD if the sample is dispensed using a handheld micropipette. Studies on the behaviour of various arsenic species in ETAAS have ben reported [34]. Interferences are far more problematic for ET-AAS than for ﬂame spectroscopy. Although true spectral interferences are equally as rare, chemical interferences and non-speciﬁc absorption (smoke) problems are exacerbated. The presence of some chemical species, e.g., chlorides, often increases the volatility of the analyte and may lead to loss at a lower pyrolysis temperature than occurs for aqueous standards. If a temperature optimization experiment is performed on standards and these “optimum” temperatures are used during the analysis, signiﬁcant losses of the analyte may occur from the samples, leading to erroneous data. Other interferences have already been discussed, such as the formation of refractory carbides and nitrides (if nitrogen is used as the inert gas). Careful optimization of the temperature programme can overcome some of these problems, especially if matrix modiﬁers (also known as chemical modiﬁers) are used. Chemical modiﬁers are reagents that are introduced with the sample that interact either with the analyte, stabilizing it thermally, or with the matrix, to make it relatively more volatile. This means that a higher pyrolysis temperature may be used before analyte loss occurs through volatilization. If a higher pyrolysis temperature can be used, then more potential interferences may be removed. An assortment of chemical modiﬁers has been used; including phosphate based ones, magnesium nitrate (this helps occlude analytes within its matrix, hence preventing their loss) and a mix of magnesium nitrate and palladium nitrate. In general, their use can extend the temperature range used during the pyrolysis stage by 300 –6008C. A review of many of the interferences encountered in ET-AAS was published by Slavin and Manning [35]. Slavin developed the stabilized temperature platform

137

L. Ebdon et al.

furnace (STPF) concept. This was a protocol based on recommended precautions to take to minimize the effects of interferences [36] and included the use of an appropriate matrix modiﬁer, integration of signals (area rather than peak height measurements), isothermal operation, rapid heating during atomization and the use of one of the more powerful background correction systems, such as the Zeeman effect. Virtually all modern analyzes made by ET-AAS use most or all of the recommendations of the STPF concept. Modern autosamplers achieve far more than simply delivering the sample to the atom cell. They may be used to perform on-line dilution, they mix the sample with appropriate matrix modiﬁers and they may be programmed to run quality control standards, re-calibrate if necessary, etc. As noted previously, isothermal operation using a platform usually helps decrease the extent of interferences because the analyte is vaporized from the platform when the temperature of the tube wall is otherwise higher. There is, therefore, less chance that the analyte will re-condense on a cooler part of the tube or recombine with cooler gas phase species, hence becoming unavailable for atomic absorption. An alternative to the use of a platform is probe atomization. This requires a specialized tube style that has a small slot machined into its side. It is through this slot that a mechanically operated graphite probe is inserted. The sample is introduced onto the probe, which is then inserted into the heated furnace through the slot wherein the sample is then dried and pyrolyzed in the normal way. The probe is then withdrawn from the tube, which is then heated to the atomization temperature, and the probe re-inserted. Again, the analytes will be radiatively and conductively heated within the tube and will therefore be less vulnerable to interferences. As discussed previously, the majority of samples introduced to ET-AAS are in the form of a liquid. However, the introduction of slurries is also possible. The relative advantages and pitfalls of slurry atomization have been discussed earlier (Section 5.3.6.6). Solids may be analyzed directly using ET-AAS if specialized tubes are used. The solid material (a few milligram) is usually weighed into a sample boat and then this is placed through a slot into a specialized tube. The furnace programme may then be run, although a drying stage may not be necessary. The technique is not common because precision can be poor. The technique is dependent upon the sample being completely homogeneous and, if only a few milligrams of sample are weighed into the boat, then homogeneity issues are of paramount importance and these may directly inﬂuence precision. There is also the possibility of some of the sample blowing from the boat prior to insertion into the atom cell. Using this technique, a precision of 10% RSD would normally be regarded as being good. The limits of detection (quoted as a concentration) obtainable using ET-AAS are relatively meaningless unless the injection volume is stated. Typically, 10– 30 ml is injected, but inevitably, 30 ml will yield a better LOD than a smaller volume. Instead, the LOD quoted as an absolute mass is usually given. As an

138

Detection methods for the quantitation of trace elements

alternative, the characteristic concentration (or mass), i.e., the concentration (or mass) that gives rise to 0.0044 absorbance (absorbance-s for integrated measurements) is also often quoted. In general, it is possible to say that for most elements, the concentration LOD for ET-AAS is often approximately 2–3 orders of magnitude lower than ﬂame AAS. The exceptions are for elements that form refractory carbides. A list of approximate characteristic masses obtainable by ET-AAS is given in Table 5.2. As well as many of the standard pre-concentration techniques discussed in Section 5.3.5 and in other chapters of the book, preconcentration is also possible using ET-AAS. If a sample aliquot of 20 ml is introduced and dried in the normal way, a second aliquot may then be introduced on top of it. If this sample introduction and drying cycle continues for four or ﬁve aliquots and then the normal pyrolysis and atomize stages are performed, an effective ﬁvefold pre-concentration may be achieved. It should be noted, of course, that there is also a ﬁvefold increase in the amount of matrix present, so unless it is a very simple matrix, such as fresh water, or unless the matrix is easily removed during the pyrolysis stage, then severe interferences may result. Although this is a very time-consuming process, the presence of an autosampler can achieve this unattended and the analyst is free to perform other tasks. It has been emphasized previously that a typical ET-AAS cycle can take in excess of 2 min to achieve. Occasionally, this may be decreased if the method of “hot injection” is used. This is achieved when the sample is introduced at a slow rate into a furnace that has been pre-heated to 120 –1308C, such that the solvent evaporates as soon as it is introduced. This can reduce the overall time of analysis dramatically. Occasionally, higher drying temperatures may be used, e.g., 4008C, although this is rare. Methods for minimizing the time required for ET-AAS determinations have been summarized by Halls [37]. Using these accelerated programmes, the overall analysis time per replicate may occasionally be decreased to 20 –30 s. 5.4.3

Multi-element ET-AAS

The majority of ET-AAS instruments is capable of detecting only one analyte at any given time. However, instrumentation is now available that uses an echelle spectrometer and a solid-state detector capable of multi-element determinations that offers considerable savings in terms of time, costs and sample and reagent consumption. The one drawback is that the analytes to be determined must have similar physico-chemical properties. This is a result of the temperature programme used being a compromise, rather than an optimum, for any individual analyte. This means that analytes that are quite volatile and that require the same chemical modiﬁer, such as As, Se, Te and Ge, may be determined together. It would, however, be inappropriate to combine one or more of these in the same determination as a much more refractory analyte such as chromium, that requires a different chemical modiﬁer and a very different temperature programme for optimum sensitivity/reduction of

139

L. Ebdon et al. TABLE 5.2 Characteristic mass for ET-AAS under optimum conditions using the most sensitive line Analyte

Characteristic mass (pg)

Analyte

Ag Al As Ba Be Bi Ca Cd Co Cr Cs Cu Dy Er Eu Fe Ga Gd Ge Hf Hg Ho In Ir K La Li Lu Mg Mn

0.7 5 10 17 0.5 9 0.6 0.2 4.2 1.5 11 6 45 100 25 2 4.5

Mo Na Nb Nd Ni P Pb Pd Pr Pt Rb Ru Sb Sc Se Si Sn Sr Ta Tb Te Th Ti Tl Tm U V W Y Zn

9 150 7 135 0.4 4 0.2 0.6

Characteristic mass (pg) 7 0.1

5 2200 6 9 70 1 15 10 14 15 10 2 4 9 50 15

22

0.15

interferences. At present, usually only three or four elements are determined simultaneously, although there are facilities for up to six. 5.4.4

Chemical vapor generation – ET-AAS

Chemical vapor generation has been coupled with ET-AAS. Generation phase interferences using this technique are basically the same as for any vapor

140

Detection methods for the quantitation of trace elements

generation determination. If the tube of the ET-AAS system is treated with a semi-permanent modiﬁer such as iridium, zirconium or tungsten, and heated to 300 –8008C, hydrides of selenium, arsenic and several other analytes may be collected quantitatively [38]. The technique is known as “in-atomizer trapping” or “in situ trapping”. The subject of in situ trapping has been reviewed by Matusiewicz and Sturgeon [39]. Since typically 5–12 ml of sample is consumed for continuous vapor generation determinations, effectively the analyte from 5 ml of sample rather than 20 ml is deposited in the tube and, hence, the sensitivity is greatly improved. For batch HG-ET-AAS determinations, a sample loop of 500 ml has been used which yielded LODs of 0.82, 0.04, 0.26 and 0.29 mg l21 for As, Bi, Sb and Se, respectively [38], representing an improvement of over 10-fold compared with conventional ET-AAS on the same instrument. Precision at the 5 mg l21 level was typically less than 3.5% RSD. The use of this method will limit the linear dynamic range accordingly. As well as improving the limits of detection, this technique also separates the analytes from potential matrix interferences. This means that lengthy drying and pyrolysis stages are not required. As a result, the time required for HG-ET-AAS is not dissimilar to conventional ET-AAS.

5.4.5

Speciation

On-line speciation analysis using liquid chromatography and ET-AAS as a detection system is relatively rare, because the atom cell is often required to be heated continuously at the atomization temperature. Since chromatograms may take several minutes to be complete, this leads to very rapid tube wear and great expense. Examples do exist, however, where very rapid temperature programmes are used that have achieved on-line speciation [40]. The majority of speciation analyzes undertaken using ET-AAS as a detector have therefore been off-line, wherein fractions (typically 0.5 ml aliquots) of the eluant are collected at the end of the chromatographic column, which are then subjected to normal ET-AAS temperature programmes so that any analytes present may be determined. The concentration of the analyte in each fraction is then plotted so that a composite chromatogram is obtained. Since the separation and detection stages are not coupled directly, there is a greater chance of contamination and mislabeling of a particular fraction. This could potentially lead to great error if a fraction containing an analyte species is present, because the transient signal obtained from this species would be “moved” to a different retention time. In addition, closely eluting species may not be fully resolved, and hence would appear as only one peak. There are also major difﬁculties in optimizing the chromatography. Despite these drawbacks, numerous examples have appeared in the literature [41].

141

L. Ebdon et al.

5.5

5.5.1

INDUCTIVELY COUPLED PLASMA-ATOMIC EMISSION SPECTROMETRY Introduction

Inductively coupled plasma-atomic (optical) emission spectrometry is often the method of choice of most laboratories when several analytes need to be determined in a batch of samples. This is because the technique can detect analytes in either a rapid sequential manner or, for some instrumentation, detection of several analytes can be simultaneous. The cost of instrumentation varies, but is typically in the range US $50,000–80,000. 5.5.2

Theory and interferences

5.5.2.1 Theory The basic theory of emission from an ICP is identical to that for ﬂame emission except that the ICP is an atom cell consisting of a very high temperature (6000– 10,000 K) ionized gas. The theory behind the formation of the ICP is discussed in detail elsewhere [42,43]. Since the plasma is at such a high temperature, any sample entering it will be desolvated; molecules will be dissociated forming atoms and, depending on the individual analyte’s ionization energy, these will become (partially) ionized. The atoms and/or ions then become thermally excited and emit light, the wavelengths of which may be separated from other wavelengths by an appropriate line isolation device and then detected. 5.5.2.2 Interferences Since the ICP is such a good excitation source, there are many species, both naturally present in the plasma and those that are introduced to it with the sample that emit light. The resulting emission spectrum can be far more complex than that produced using ﬂame techniques and the chances of line co-incidence are much greater. The line isolation devices used in ICP-OES, therefore, tend to be more highly resolving than those required for the ﬂame techniques. Despite the improved resolution, interferences are still common. The choice of which analytical line to use is therefore governed by both the potential interferences and by the sensitivity required. A more comprehensive discussion of interference effects in ICP-OES has been given in Ref. [42], and will therefore, only be dealt with brieﬂy here. Overlap from other spectral lines (atomic, ionic and molecular) is common. The high temperature of the plasma causes species that are normally not problematic in ﬂame spectroscopy to emit light. The high temperature exacerbates the problem because it causes line broadening, and broader lines are more likely to lead to spectral overlap than narrow ones. The argon that forms the plasma emits at approximately 200 different wavelengths and these emissions, together with the emission from

142

Detection methods for the quantitation of trace elements

assorted molecular species derived from water, entrained gases and the sample matrix, e.g., OH, N2þ, NH and NO, which produce a series of molecular bands that are spread throughout the wavelength range, can clutter the emission spectrum considerably. Line overlap by concomitant metallic species may also exist. This problem is especially severe when line rich elements such as iron, the lanthanides or uranium are present at an appreciable concentration in the sample. A background emission continuum is also present, the intensity and characteristics of which will vary, depending on the solvent loading, the solvent type and the matrix elements. Stray light, i.e., light that unintentionally reaches the detector, may also be a problem. This often arises from imperfections in the dispersing device, but many modern instruments suffer from this problem far less because of the quality of the optics. It should still be noted, however, that an analyte line that has a low intensity may be interfered with by a nearby very strongly emitting species. Most modern instruments have a software library giving a list of the relevant lines of the analytes, together with the potential interferences that may be experienced at each line. Inspection of the line tables and prior knowledge of the sample chemistry usually enables an analyst to pick a suitable “interference free” wavelength. Many instruments also enable background correction methods to be used in an attempt to compensate for any interference effects. There are a number of correction methods that may be used and these are discussed elsewhere [42]. 5.5.3

Instrumentation

5.5.3.1 RF generators The radio frequency (RF) generator may be of several types, e.g., crystal controlled or free running, 27.12 or 40.68 MHz. A far more detailed discussion on RF generators has been given elsewhere [42]. Both 27.12 and 40.68 MHz generators are used commercially and both normally produce RF power at up to 2000 W, although for normal usage, a power of between 1000 and 1500 W is typical. In general, the 40 MHz generators are regarded as being more stable, to couple more efﬁciently and to produce a lower background signal. Therefore, slightly improved LODs may be achieved for these instruments compared with those obtained from an instrument equipped with a 27 MHz generator. 5.5.3.2 Torches The plasma is formed in a torch, which is a concentric arrangement of quartz tubes that permits delivery of independently adjustable ﬂow rates of argon to one end which is located in the RF load coil. The plasma is formed from argon gas ﬂowing at a rate typically between 11 and 15 l min21. This ﬂow is called the coolant or plasma gas ﬂow. The auxiliary (intermediate) gas ﬂow (typically 1 l min21) prevents the plasma from sitting too low in the torch and melting the innermost tube (the injector). The nebulizer (carrier) gas ﬂow passes through

143

L. Ebdon et al.

the injector and punches a hole through the ﬁreball, forming an annular, doughnut-shaped plasma. Technically, this is termed the annulus and the surrounding ﬁreball the torus. Several types of torch exist, but most now conform to the basic Fassel style, which consumes substantially less gas (14– 17 l min21) when compared with the much larger Greenﬁeld style torch (typically 12 –38 l min21 argon and 20–70 l min21 nitrogen). Similarly, the powers required to operate the Fassel torch are 1.0 –1.5 kW, compared with several kW for the Greenﬁeld torch. The advantage of the Fassel style torch is that it is less expensive to operate, but its drawback is that it is less robust than the Greenﬁeld torch and is less tolerant of gases other than argon. Many torches are demountable or partially demountable. This usually means that the coolant and auxiliary tubes are ﬁxed, but that a different injector may be introduced. The shape and bore of the injector of the torch may have a large effect on the stability of the signal. Wider bore injectors (e.g., 2 or even 2.5 mm) are less likely to block than normal injectors (1.5 mm) if samples with a high dissolved salt content are introduced. If the injector is too wide, however, problems may be experienced in “punching” the plasma and the plasma may simply extinguish. Some injectors are made from ceramic or alumina and are therefore more resistant to hydroﬂuoric acid than quartz ones. Some injectors taper gently from wide to narrow bore and these are less likely to become blocked than injectors that have a step reduction in bore. A demountable or partially demountable torch therefore gives the analyst more freedom of choice to use an appropriate injector type. Several variations of the Fassel style torch exist. These include low ﬂow torches that are much smaller (i.d. 13 mm compared with 18 mm for a conventional torch) and operate at a lower power (,1 kW) and with a lower consumption of gas (8 l min21) [44]. Micro-torches that operate at even lower power and gas ﬂow also exist [45]. These torches are reported to offer similar sensitivity to their larger counterparts, but are more easily blocked because of their smaller diameter injectors. 5.5.3.3 Radial and axial plasmas The majority of instruments use a radial conﬁguration wherein the plasma is viewed from the side. Axial instruments have the torch turned at a right angle so that it lies horizontally and the plasma is viewed end-on. There is a great deal of discussion as to which orientation offers the best performance. Some workers state that the axial instruments offer improved limits of detection by a factor of nearly 10, because the light from a much larger area may be detected. However, others have stated that they are prone to far more interferences, because they are less “optically thin” than the radial instruments. This means that the emitted light has to pass through a much larger distance, wherein it may be absorbed by other analyte atoms/ions, or that effects from molecular interferences are greater. A recent paper has addressed this problem and stated that self-absorption effects in axially viewed plasmas are partially controllable

144

Detection methods for the quantitation of trace elements

by careful optimization of the operating conditions [46]. The relative merits of both axial and radial instruments have been discussed by Brenner and Zander [47]. For axially viewed plasmas, a shear gas is often required to prevent thermal damage to the collection optics. It has the additional effect of removing the ICP tail ﬂame that, under normal circumstances, would be rich in interfering species. 5.5.3.4 Wavelength isolation and detection systems Traditionally, ICP spectrometers utilize a monochromator and photomultiplier tube arrangement similar to AAS instrumentation. The monochromators, however, tend to have a much longer focal length and are of higher resolving power than those used for AAS. The higher resolution is required because the high temperature of the plasma excites many more species than does a ﬂame and hence the emission from ion lines and some molecular species may become problematic if low-resolution monochromators were to be used. As its name suggests, instruments that utilize a monochromator can only interrogate one wavelength at any one time, and must scan over several wavelengths sequentially if more than one analyte is to be determined. The speed with which it can achieve this will govern the overall time required for analysis. Also, the accuracy and repeatability with which it ﬁnds each wavelength will have a large effect on the accuracy and precision of the analysis. Fortunately, once optimized, most modern instruments tend not to drift signiﬁcantly (unless physical parameters, such as the room temperature, change). Polychromators have been developed commercially that may determine several analytes simultaneously. Here, several PMTs are arranged at intervals around a circle (known as a Rowland Circle) and as the light emitted from the plasma is diffracted from the grating, the wavelengths are separated and each PMT may detect one particular wavelength. Since the PMTs are not easily moved, the instrument is usually prepared in the factory to determine only speciﬁc wavelengths that the customer requires. Therefore, although simultaneous analyte determinations are possible using such instrumentation, it is cumbersome and extremely inﬂexible. Many modern instruments use an echelle-based spectrometer and specialized charge transfer device detectors. These may be either charge coupled devices, segmented charge coupled devices or charge injection devices. The theory behind their operation is beyond the scope of this chapter, but may be found in Refs. [48,49]. These devices function as an “electronic photographic plate” and are therefore truly simultaneous and may be used to determine several analytes together with suitable background correction points at once. There are a few drawbacks associated with their use, including the possibility of “blooming”, which occurs when an analyte may be so concentrated that the individual pixels detecting that wavelength may become saturated so that the charge spills over into adjacent pixels, thereby giving erroneously high signals for other analytes. Modern electronics have gone a long way in overcoming this

145

L. Ebdon et al.

problem. Also, these instruments do not have the ﬂexibility of PMT based spectrometers, because they may have only a limited number of lines that can be detected. However, this number reaches into the hundreds and, therefore a suitable line should be available for most analytes. 5.5.3.5 Sample introduction systems Nebulizers and spray chambers The nebulizer/spray chamber assembly performs the same functions as those in the ﬂame systems, i.e., to form an aerosol and then segregate large droplets from smaller ones. There are, however, a very wide variety of nebulizers and spray chambers. A description of many types has been given recently by Thomas [50]. Although this paper describes sample introduction for ICP-MS analyzes, the principles and most of the instrumentation are identical. The processes occurring within them have been discussed in two papers by Sharp [51,52]. Some nebulizers, such as the Meinhard style ones, are self-aspirating, i.e., they draw liquid samples up in a similar fashion to ﬂame AAS nebulizers, and others require the sample to be pumped to them. Some nebulizers are easily blocked by the presence of dissolved or suspended solids, whereas others are far more tolerant. Examples of dissolved solids tolerant nebulizers include the crossﬂow, the Ebdon, the Burgener, the Hildebrand grid and assorted specialized pneumatic ones. The Ebdon, crossﬂow and some of the pneumatic ones are also tolerant of suspended solids such as those found in slurries. The choice of nebulizer will depend largely on the application. Some nebulizers are manufactured from inert polymers and are therefore more resistant to corrosive samples, such as those containing hydroﬂuoric acid. As well as acting as a droplet size ﬁlter, for those nebulizers that require sample to be pumped to them, the spray chamber acts as a pump noise dampener. A typical ICP nebulizer/spray chamber assembly will have a transport efﬁciency of 1–2%. Much more than this is likely to lead to severe plasma perturbation and its possible extinction. The problem is exacerbated by the aspiration of organic solvents. As discussed previously, these tend to have lower viscosity, lower surface tension and higher volatility (higher vapor pressure), leading to transport efﬁciencies substantially higher than 1–2%. Many of the more modern generators (especially the more robust 40 MHz ones) may be able to cope with the increased solvent loading, however, many of the older ones cannot and plasma extinction occurs. Boorn and Browner discussed introduction of organic solvents to ICPs [53]. If larger volumes of solvent are likely to reach the plasma, a desolvation device should be used. These come in several different forms, including membrane drier tubes [54], desolvation devices made in-house [55] and commercial equipment. Many of these devices decrease the amount of solvent reaching the plasma, whilst not signiﬁcantly decreasing the analyte transport efﬁciency. The reduction in the solvent loading often leads to greater stability and, hence, improved LODs.

146

Detection methods for the quantitation of trace elements

An ultrasonic nebulizer increases transport efﬁciency to approximately 25–30%. These usually have built in heating and cooling devices to desolvate the aerosol and prevent plasma perturbation. Such a device, by increasing analyte transport and decreasing the solvent loading, improves LODs by typically 10-fold. The disadvantages of the nebulizer include its cost (approximately US $10,000) and the need to optimize the operating conditions carefully. Failure to optimize the heating and chilling temperatures is likely to lead to inconsistent nebulization and a noisy signal, which degrades the limits of detection. The whole device may be used without the need of a spray chamber. Spray chambers are available in assorted shapes and sizes. Their function is to separate large aerosol droplets from smaller ones and to act as a noise dampener. The efﬁciency with which the spray chamber achieves the latter function is often dependent on its internal volume. Larger spray chambers, such as the Scott double pass, dampen the noise quite effectively, whereas the numerous reduced volume ones are less effective. Conversely, the larger volume Scott style has a greater internal surface area and regions of dead volume, i.e., areas where the nebulizer gas ﬂow does not rapidly ﬂush any sample entering it away. Some analytes may exhibit a much longer memory effect or “washout” period in such a spray chamber, because it may become adsorbed to the glass walls (e.g., for lead) or may simply become trapped in an area of dead volume. For routine analysis, this problem is little more than annoying, because the main result is that a longer washout period is required between samples, which obviously extends the analysis time and increases the cost of analysis. However, when transient signals are obtained, especially those arising from chromatography, the memory effects can broaden the analyte peaks to the extent where they may start to merge. This is obviously undesirable, as confusion between different analyte species may result. Broader peaks also lead to lower signal-to-noise ratios and inferior limits of detection. The broadening effects are reduced for low volume spray chambers such as the cyclone and single pass styles. If a high quality liquid chromatography pump is used, the pump noise should be minimal and so the reduced volume spray chambers often offer the best resolution and sensitivity with adequate noise characteristics. Some corrosion resistant spray chambers manufactured from polymers (e.g., Ryton) are also available. Many spray chambers come with a jacket surrounding them, through which a cooling ﬂuid is pumped to maintain the spray chamber at a constant temperature and improve stability. The cooling ﬂuid is usually water, although anti-freeze may be used at a temperature of 25 to 2 108C to decrease the vapor pressure of organic solvents. This may help to decrease plasma perturbation by minimizing plasma loading. Sample throughput will depend on the type of instrument used. If a sequential spectrometer is used, then the determination of each analyte may take 20 s and so if 10 analytes are to be determined, then the analysis time for

147

L. Ebdon et al.

one sample may be 3–5 min; leading to a sample throughput of approximately 12–20 samples per hour. Inevitably, the sample throughput will be dependent on the number of analytes, with throughput increasing with a decreasing number of analytes. If a simultaneous spectrometer is used, then the same time will be required for one analyte as for 10, and so sample throughput is likely to be greatly increased. Assuming that the instrument has been set up properly, with the analyte lines “trimmed” so that measurement is made from the top of the peak rather than the rapidly sloping sides, then precision can be 1–5% RSD. The limits of detection for many analytes using optimal conditions are given in Table 5.3. It should be noted, however, that since the ICP has a deﬁnite structure and each region has a different temperature, ionizing properties, etc., then each analyte has an optimal set of conditions which yield the best sensitivity. The most critical parameters that govern analyte sensitivity are the viewing height, the nebulizer (carrier) gas ﬂow rate, the power and, to a lesser extent, the auxiliary (intermediate) gas ﬂow rate. If several analytes are to be determined, then compromise conditions will probably have to be used. Since compromise conditions are the “best overall”, but may not actually be optimal for any given analyte, the LODs obtained will be inferior to those shown in the table. Occasionally, other specialized nebulizers are used, e.g., the thermospray, the electrospray and the direct injection nebulizers. The theory of these is beyond the scope of this chapter, but may be found in Refs. [56,57]. Each of these nebulizers produces a very ﬁne aerosol and usually operates at low ﬂow rate (typically 10–50 ml min21). They can therefore be placed at the base of the plasma torch, omitting the spray chamber. Transport efﬁciency to the plasma is virtually 100%, but plasma extinction is prevented because only a similar absolute volume of sample reaches the plasma in any given time period as for a conventional nebulizer. Other sample introduction methods There is a plethora of alternative sample introduction methods for ICP spectrometry, including those described for ﬂame spectrometry, i.e., FI and chromatography. A typical sample ﬂow rate for HPLC is 1– 2 ml min21, which is compatible with the sample uptake of an ICP. However, when organic solvents are to be introduced, a desolvation device may be required to prevent plasma extinction. Similarly, for HPLC applications that use a mobile phase containing a high dissolved salt content, nebulizers and torch injectors that are tolerant of this must be used. The use of ICP-OES as a detector for elemental speciation studies has been described recently [58]. Liquid chromatography is normally coupled with ICP spectrometry via the nebulizer and spray chamber assembly, although the electrospray, thermospray and direct injection nebulizers have also been used. Gas chromatography has occasionally been coupled with ICP spectrometry, but few routine applications exist. The problem (as with GC –AAS coupling) is the transfer of the analyte to the atom cell in a

148

Detection methods for the quantitation of trace elements TABLE 5.3 Limits of detection for ICP-OES under optimum conditions with conventional nebulization Analyte

LOD (mg l21)

Analyte

LOD (mg l21)

Ag Al As Ba Be Bi Ca Cd Co Cr Cs Cu Dy Er Eu Fe Ga Gd Ge Hf Hg Ho In Ir K La Li Lu Mg Mn

3 1.5 12 0.07 0.2 12 0.03 1.5 5 4 3200 2 0.3 0.7 0.3 1.5 6.5 3 13 4 8.5 0.5 18 4 10 0.02 0.6 0.05 0.1 0.3

Mo Na Nb Nd Ni P Pb Pd Pr Pt Rb Ru Sb Sc Se Si Sn Sr Ta Tb Te Th Ti Tl Tm U V W Y Zn

4 1 4 2 6 18 14 7 0.8 20 3 6 18 0.4 37 5 15 0.02 9 5 27 17 0.6 16 1.5 18 2 17 0.2 0.9

sufﬁciently hot form to prevent condensation. Coupling of a heated transfer line to the ICP torch can be problematic because any metal components within the transfer line may act as an aerial for the RF power, leading to a potential hazard. The transfer line must normally be placed as far up the torch as possible to prevent analyte condensation, whilst ensuring that the end of it does not melt and that potential hazards are avoided. A further problem is that the

149

L. Ebdon et al.

transport gas ﬂow rate typical of GC separations is not sufﬁcient to punch a sample channel into the plasma. A make up gas is therefore usually required. This too often requires heating to prevent analyte condensation. Sample throughput is obviously dependent upon the length of time required for the chromatography, but for many HPLC and GC applications, only 3–8 samples per hour may be analyzed. Precision is also dependent on the method of sample introduction, but often lies in the region of 3 –10% RSD, and may be improved if an appropriate internal standard is used. Chemical vapor generation is a popular method of sample introduction for ICP spectrometry as well as ﬂame spectrometry and the beneﬁts of this approach are the same. The one problem that may be encountered is the production of excess hydrogen as a by-product of the hydride generation reaction. As discussed previously, some instruments are relatively intolerant of gases other than argon and so, if large quantities of hydrogen enter the plasma, perturbation may occur. The use of an automated continuous hydride generator is highly recommended, as is careful optimization of both the reagent concentrations, to minimize the excess hydrogen produced, and the instrumental operating conditions. A recent example of HG-ICP-OES is presented by Overduin and Brindle [59]. Some workers have coupled chromatography with HG prior to ICP-OES detection [60]. Again, this would improve the sensitivity when compared with HPLC –ICP-OES. Just as with HG-AAS, sample pretreatment may be necessary to transform some species into a form that produces a hydride. Electrothermal vaporization has also been used to introduce assorted sample types to ICP-OES instrumentation. The principles of ETV are the same as those described previously. Again, it is a useful technique when only limited sample volume is available. It may be used to analyze liquid samples, slurries and solid samples directly. The same heating programs are used, i.e., a dry, a pyrolysis and a vaporization stage followed by a high temperature cleanup step. For ETV–ICP-OES (and -MS), the vaporization stage does not have to atomize the analyte. As long as the temperature is sufﬁciently high to vaporize the analyte, either as an atom or as a compound, it may be transported in a ﬂow of inert gas to the plasma. Since this is at a temperature of 6000 –10,000 K and the sample arrives in a dry form, the plasma has more energy available for atomization and excitation, so it will dissociate the vast majority of analyte compounds. If adequate pyrolysis temperatures are used, the analyte is separated from the majority of the matrix, thereby facilitating interference free determination. Two reviews of the process of ETV–ICP have been published; an early one by Carey and Caruso [61] and a more recent one that compares ETV with laser ablation (LA) [62]. Sample throughput is again dependent on the type of instrumentation used and the number of analytes to be determined. Since the signal obtained is in the form of a transient, for sequential instruments, a number of replicate analyzes will have to be performed because the instrument will not have sufﬁcient time to scan to more than one

150

Detection methods for the quantitation of trace elements

wavelength before the signal returns to the baseline. Since each replicate analysis may take up to 2 min, and three replicates are normally required per element, then if three elements are to be determined, each sample will take 20 min. This may be longer if solids are weighed directly into the graphite atomizer. Precision should be roughly equivalent to that obtained with conventional ET-AAS. Again, LODs should be measured as an absolute amount rather than as a concentration. In general, sensitivity is improved because transport efﬁciency of the analyte to the atom cell is substantially larger than for conventional solution sample nebulization, although condensation of the analytes within the transfer line may occasionally decrease the transport efﬁciency. Direct sample insertion (DSI) and in-torch vaporization (ITV) are off-shoots of ETV. Sample (either liquid or solid) is dispensed onto the tip of a probe, usually made of graphite or a refractory metal. The probe is then inserted directly up the injector of a specialized torch towards the plasma. Sample transport efﬁciency is close to 100%, but the analyte is not separated from the matrix and determinations are thus more prone to interferences. The technique was reviewed in 1990 by Karanassios and Horlick [63] and again in 1999 by Sing [64]. In LA, a laser beam is focused either onto, or just above, the surface of a sample. The laser vaporizes a small area of the sample and the vapor is transported in a stream of inert gas to the plasma. The laser may be focused onto extremely small areas (, 0.1 mm) and hence may be used, for example, to analyze ﬂuid inclusions in geological materials. If the laser is aimed at the same spot on some types of sample, then depth-proﬁling is possible, i.e., the top 0.1 mm of surface is analyzed, followed by 0.1 mm below that, etc. This may be of use for some sample types where the depth may be correlated directly with age. There are several problems associated with quantitative analysis using LA –ICP-OES. Since only a very small area of sample is vaporized, if a bulk analysis of a sample has to be performed, it is essential that it is homogeneous, otherwise accuracy and precision will be affected. Since the laser radiation will interact with different types of sample to different extents, it is necessary for calibration to be performed using materials that have an identical matrix. Failure to calibrate properly will lead to the results being, at best, semiquantitative. Laser ablation may also be used to “map” the surface of a sample, i.e., to determine how an analyte concentration varies over the surface of the sample. Laser ablation produces transient signals so, as with ETV sample introduction, a sequential instrument will not be able to determine typically more than one analyte at any given sample site, whereas a simultaneous instrument could potentially do several. A review of the interaction of laser radiation with samples has been given by Darke and Tyson [65]. A more recent review comparing ETV and LA has been made by Kantor [62]. Accuracy and precision of the technique will depend on the sample homogeneity and on how

151

L. Ebdon et al.

closely the standards are matrix matched to the samples. These issues are discussed in more detail by Russo in a later chapter in this book. Solid samples may be introduced directly to the plasma by several techniques, including slurry sampling, ETV, DSI and LA. In addition to these methods, some instruments come with a solid sampling (SS) accessory. The requirement for this is that the sample must conduct electricity and therefore metallurgical samples, including steels, brass, other alloys, wires and even coal ﬂy ash, may be analyzed. An arc or a spark is used to ablate material from the surface of the sample and the dry aerosol produced is transported to the plasma in a stream of argon. The sample may be in the form of rods, powders or briquettes. The technique has a few variants but most deliver a precision of 0.2 –1% RSD for a concentration of 1%. Custom accessories have even been produced commercially that enable the determination of wear metals in lubricating oils [66]. A direct current arc has been reported as giving a precision of 3–10% RSD at the 1% concentration level. One of the drawbacks with this method of sample introduction is the need for very closely matrix-matched standards. 5.5.4

Figures of merit

As noted throughout the text, sample throughput and precision will depend on the instrumentation used, the number of analytes to be determined and the sample introduction method. Assuming a liquid sample (or a digested solid sample) is to be analyzed using conventional nebulization, then a modern simultaneous instrument may analyze 25 –30 samples per hour and use only 1–5 ml of sample. A sequential instrument will analyze fewer than this and will consume substantially more sample. Both types of instrumentation should provide analytical results with a precision of 1–5% RSD. The linear range should extend to at least ﬁve orders of magnitude for ICP-OES determinations, although it should be noted that for some applications, e.g., chromatography, the chromatographic column may become overloaded at higher concentrations of analyte. 5.6 5.6.1

INDUCTIVELY COUPLED PLASMA-MASS SPECTROMETRY Introduction

Inductively coupled plasma-mass spectrometry was developed in the 1970s and commercial instrumentation was available in the 1980s. It is a coupling of an ICP with mass spectrometric detection. The principles behind the sample introduction and the processes of plasma formation and of desolvation, dissociation, atomization and ionization within the plasma are the same as those described previously. It has become a very popular method of analysis because it has several advantages over other techniques, i.e., it is extremely

152

Detection methods for the quantitation of trace elements

rapid, with simultaneous and quasi-simultaneous instruments being available; it offers improved sensitivity over many of the other techniques for most analytes and the mass spectrum produced from any sample is far more simple than that obtained from an emission instrument. There are several different types of ICP-MS instrument. Each of these will be discussed in more detail later. The one major drawback is the cost, ranging between US $80,000 and US $400,000, depending on the type. A schematic diagram of a general ICP-MS instrument is shown in Fig. 5.1. 5.6.2

Theory

As discussed previously, the sample introduction systems used for ICP-MS can be identical to those for ICP-OES. Similarly, the processes occurring within the plasma are also identical. It is worth noting, however, that since all ICP-MS instruments detect the analytes according to a mass-to-charge ratio ðm=zÞ, for any signal to be detected, the analytes must become ionized within the plasma. The extent of ionization will depend on several factors, but most importantly on the ﬁrst IP of the analyte. The plasma consists of ionized argon that has a ﬁrst IP of 15.76 eV. Therefore, any element that has an IP less than this will be at least partially ionized. Cesium, having a ﬁrst IP of 3.89 eV, will be 100% ionized, but arsenic has a ﬁrst IP of 9.81 eV and will be only 30– 40% ionized. Fluorine, with a ﬁrst IP of 17.42 eV, will not be ionized and therefore cannot be determined directly by ICP-MS (using an argon plasma).

Fig. 5.1. Schematic diagram of an ICP-MS instrument.

153

L. Ebdon et al.

Once the ions have been formed, they must pass from atmospheric pressure through several chambers of increasingly high vacuum to the mass separation and detection stages. Several different types of mass ﬁlters and detectors exist, which will be discussed in a later section (Section 5.6.3.1). A more detailed account of how the ions pass from the plasma through the expansion chamber and ion lens system to the mass ﬁlter and detector may be found elsewhere [2,67 –69]. 5.6.3

Instrumentation

5.6.3.1 Mass ﬁltration A discussion of ICP-MS instrument types has been given in a recent book chapter [70]. In addition, several tutorials have also been published recently that have discussed the various types of mass ﬁlters [71– 73]. The most common mass analyzer is the quadrupole. Four rods (often made from molybdenum) are arranged in a set of two pairs in a square orientation. Two rods have a DC voltage on them and the other two have RF voltage. The magnitude of the voltages will allow one m=z to pass through the rods towards the detector, whilst ensuring that all ions of other m=z collide with one of the rods, hence preventing them from being detected. A short while later (often , 1 ms) the magnitude of the voltages changes and an ion of different m=z is allowed to pass to detection. A quadrupole instrument is therefore not truly simultaneous, but instead is so rapidly sequential as to be regarded as being quasi-simultaneous. This device is relatively inexpensive and robust, but has a relatively poor resolution, i.e., between 0.7 and 1.0 atomic mass units (AMU). This means that it is more prone to spectral interferences than other mass analyzers. The problem of interferences is discussed in a later section (Section 5.6.5). Other, more highly resolving ICP-MS instruments are available commercially, although at much greater cost. An example is double focussing magnetic sector instrumentation. The principle of operation of this instrumentation has been discussed by Thomas [72]. These instruments have a resolving power of up to 10,000 compared with a quadrupole-based instrument that has a resolving power of only 300. This large improvement in resolution will enable distinction to be made between some interfering polyatomic ions and analyte ions (see Section 5.6.5). The resolution of the instrument can be set by the analyst so that individual interferences can be overcome. The resolution required obviously depends on how close the interfering species is in mass to the analyte. A few examples include 34Sþ and 16O18Oþ that may be separated using a resolution of 1300 and 75Asþ and 40Ar35Clþ, which require a resolution of 7725. It should be noted, however, that the higher the resolution required, the lower the sensitivity. It is therefore advisable to use the lowest resolution necessary to achieve interference free determination. As well as specializing in the reduction of interferences, it can be used to gather extremely precise isotope ratio data. If used in low resolution and for interference free analytes, a

154

Detection methods for the quantitation of trace elements

precision of 0.01–0.05% is obtainable. In addition, the sensitivity at low resolution can be at least an order of magnitude superior to quadrupole instruments. Some instruments of this type use several detectors. These multicollector instruments have the capability of detecting and measuring multiple ions simultaneously and are regarded as being capable of producing the ultimate in precision for isotope ratio measurements. The time-of-ﬂight ICP-MS (ICP-TOF-MS) is the most recent development of ICP-MS instrumentation, being commercially available since 1998. Although still relatively immature, it does have several potentially important advantages over other instrument types. This instrumentation permits truly simultaneous detection and therefore has advantages when measuring transient signals; it produces high precision isotope ratios and can decrease the amount of time required to complete an analysis. A more complete discussion of the theory behind ICP-TOF-MS, the instrumentation, and its relative merits, can be found in the literature [73– 75]. Examples of the use of ICP-TOF-MS instruments include the determination of isotope ratios [76] and the detection of rare earth elements in seawater after FI matrix elimination and pre-concentration [77]. 5.6.3.2 Reaction cells The use of reaction cells has been discussed recently by Thomas [78]. These devices are placed between the ion lens system and the mass ﬁlter. As the ions from the plasma enter, a quadrupole, hexapole or an octopole cell helps focus them towards the reaction cell gas (usually helium or hydrogen). As the ions and the reaction gas collide, the polyatomic ions fracture, leading to a decrease in the interference observed [79,80]. The use of reaction cell technology has improved the analytical capabilities of quadrupole based ICP-MS instruments in terms of both interference and, for some analytes, limits of detection. Even for analytes that are renowned for being difﬁcult to determine using ICP-MS, such as Fe, LODs signiﬁcantly below 1 ng ml21 may be obtained when a reaction cell is used to overcome the interference caused by 40Ar16Oþ on 56Feþ. 5.6.3.3 Detectors There are several types of detector available for ICP-MS instruments. The channel electron multiplier is a horn shaped device that is coated with a semiconductor. As an analyte ion impinges on the surface, an electron is ejected which is accelerated down towards the other end of the tube, but on its way down, collides with the wall of the tube ejecting secondary electrons. Each of these is also accelerated towards the other end of the tube and they collide with the wall producing further electrons. An avalanche effect is therefore built up. The number of electrons reaching the pre-ampliﬁer at the far end of the tube is proportional to the number of analyte ions impinging on the detector, i.e., the concentration of the analyte. The discrete dynode electron multiplier functions in a very similar way, but instead of using a continuous tube like dynode, as in

155

L. Ebdon et al.

the channel multiplier, it uses a series of discrete dynodes. The Faraday cup may be used when ultra-trace detection limits are not required. A more detailed description of the different detector types and how they may be used either in combination, or singly, to achieve a linear range spanning nine orders of magnitude, has been given by Thomas [81]. 5.6.4

Different types of analysis

Analysis using ICP-MS offers a wide variety of options. It may be used in the normal, fully quantitative analysis mode, or as a detector for chromatographic separations where perhaps only one or two target elements may be determined. Other time-resolved functions include serving as a multi-element detector for ETV, LA and FI analyzes. Semi-quantitative analysis is also achievable wherein a mass response curve is prepared using one mixed standard of perhaps six elements, each at either 10 or 100 ng ml21. The response from each of these elements is calculated and the line of best ﬁt between each of the points plotted. The software will “assume” that for the same concentration, any other analyte will have a response on that line of best ﬁt. The method is semiquantitative, because only an estimate of the analyte’s concentration can be made, although it is normally accurate to within a factor of two. A suite of nearly 70 elements can have their concentrations estimated within 10–30 s. The method is especially useful when a sample of completely unknown characteristics must be analyzed. A semi-quantitative analysis will enable the analyst to identify an appropriate concentration range for the standards prior to a fully quantitative analysis or it may be used to identify suitable internal standards. Isotope ratio measurements for isotope dilution (ID) is regarded as being a deﬁnitive method of analysis. The subject of isotope ratio measurements has been discussed in great detail in a recent book [82]. Although possessing a number of advantages, the greatest drawback is cost; the price of pure isotopes or even isotopically enriched metals can be prohibitive. For species speciﬁc ID, the isotopically enriched compound will probably have to be prepared in-house and the cost of enriched isotopes is high. Once prepared, the isotopically enriched compound should be analyzed using an assortment of instrumental methods, such as nuclear magnetic resonance, so that an estimate of its purity can be made. A discussion of ID methods for trace metal speciation has been published recently [83]. 5.6.5

Interferences

There are several types of interference that may occur in ICP-MS analyzes. Those attributable to sample transport effects are the same as for ICP-OES analyzes. The presence of 0.5% dissolved solid and high concentrations of acid in a sample will not nebulize with the same efﬁciency as a standard prepared in

156

Detection methods for the quantitation of trace elements

2% nitric acid, and hence a different signal may be obtained for the same concentration of analyte. Additionally, space charge interferences may arise when the ion current in the sampled analyte beam exceeds the capacity of the ion lens systems to maintain focusing and the transmission efﬁciency is changed. This usually occurs as a consequence of the presence of high concentrations of concomitant elements and tends to favor the transmission of higher mass analytes from the beam. These effects may be partially overcome by the use of at least one internal standard. As usual, the internal standard should not be present naturally at a signiﬁcant concentration in the sample and should match, as closely as possible, the ionization energy and mass of the analyte. If a range of analytes is to be determined, e.g., 65Cuþ and 66Znþ, 111Cdþ and 208Pbþ, then it may be necessary to have up to three internal standards, one at the lower m=z range such as 59Coþ, one in the middle, such as 115Inþ and one at the higher m=z range, such as 205Tlþ. The use of more than one internal standard may lead to greater long-term instrument stability. Since the mass response curve (i.e., the signal obtained per unit concentration over the mass range) may change with time, the use of a single internal standard, such as 115 Inþ, may be insufﬁcient. For example, if after 50 samples have been analyzed, a standard containing 100 ng ml21 of analytes is analyzed as a check standard, it may be found that the concentrations range from 70 ng ml21 at the lower mass range up to 130 ng ml21 at the higher end. If two or more internal standards are used, instrumental drift can be diminished to , 10% over a whole day’s work. There are several types of spectroscopic interference. The most common is that of polyatomic interferences. This occurs when two (or more) atoms form a molecule that has nominally the same mass as the analyte. The vast majority of this type of interference occurs below m=z 80 (the argon dimer) and these often contain argon (from the plasma) combined with an ion present in the matrix of the sample. Examples include the interference from 40Ar35Clþ on 75Asþ, 32 16 S O2þ on 64Znþ, 40Ar16Oþ on 56Feþ and 23Na40Arþ on 63Cuþ. A far more complete list is given in a review by Evans and Giglio [84]. In a paper by Nonose and Kubota [85], the interferences observed in quadrupole and in high resolution ICP-MS instruments are compared. Isobaric interferences occur when two analytes have isotopes of nominally the same mass, such as for 113Cdþ and 113Inþ. However, most elements have at least one isotope free from such interference. Doubly charged ion interferences also occur, but the only element that suffers from this to any signiﬁcant degree is barium, because it is the only commonly determined analyte that has a second IP , 15:76 eV: The overall effect is that the signal for 138Baþ decreases whilst the signal for 138Ba2þ (which is equivalent to 69Gaþ) increases. The extent of ionization may be different between samples and standards and so either/or the Ba and Ga determination may be affected. Another type of interference arises due to the formation of metal oxides. This is similar to polyatomic interferences and occurs mainly for the rare earth elements, with

157

L. Ebdon et al.

the lower mass analytes, that are usually there at appreciably higher concentration than the ones with higher m=z, combining with oxygen to give an elevated signal at M þ 16, e.g., 141Pr16Oþ interfering with 157Gdþ. There are several means that may be used to overcome spectroscopic interferences. The easiest is to use an alternative isotope that does not suffer from interferences, although some elements are mono-isotopic. Sometimes, it is possible to perform some chemistry on the sample prior to analysis such that the interferences are separated from the analytes. An example has been the use of a FI technique with a micro-column of a chelating resin to retain the analytes, whilst potential interferences were washed to waste by an appropriate buffer [86]. Some of the other alternative sample introduction methods also succeed in separating the analyte from potential interferences, such as chemical vapor generation and ETV. The introduction of alternative gases has also been demonstrated to overcome some interferences. The introduction of 4% v/v nitrogen to the nebulizer gas ﬂow has been shown to markedly reduce the interference from 40Ar35Clþ on 75Asþ [87]. The mechanism by which this works is uncertain, but a concomitant increase in the signal at m=z 51 (14N37Clþ) and at 49 (14N35Clþ) would appear to indicate that a favorably competitive reaction is occurring. Hydrocarbon gases have also been shown to be beneﬁcial for many analytes [88,89]. As well as chemical methods of interference removal, instrument manufacturers have also produced several hardware and software methods. The software based methods are mathematical algorithms that rely on correction factors. For instance, the extent of 40Ar35Clþ interference on 75Asþ can be estimated by taking into account Se signals at m=z 77 and 82. The Se isotope at m=z 77 is also interfered with by chloride (40Ar37Clþ), but the isotope at m=z 82 is not. Since each isotope’s theoretical relative abundance is known, any deviation from this known ratio can be measured and a correction made. Hardware modiﬁcations and accessories offer a more reliable method of overcoming interferences. The collision cell and the dynamic reaction cell have been successfully used to overcome interferences (see Section 5.6.3.2). The use of high resolution mass analyzers also overcomes the vast majority of common interferences. Most polyatomic interferences exist only for quadrupole-based instrumentation. This arises because even though the interferences and analyte ions do have a slightly different mass (e.g., 75Asþ actually has an m=z of 74.926 whereas 40Ar35Clþ has m=z 74.932), as discussed previously, the quadrupole only has unit mass resolution and can therefore not distinguish between the two. Magnetic sector instruments are capable of much higher resolution and can distinguish between the two masses, hence eliminating the interference. 5.6.6

Sample introduction techniques

In general, the principles, advantages, drawbacks and applications of the assorted sample introduction techniques are the same for ICP-MS as for ICP-

158

Detection methods for the quantitation of trace elements

OES. As well as increased sensitivity, the one big advantage of ICP-MS over some ICP-OES instrumentation is that it is simultaneous (or at least far more rapidly sequential). This means that for sample introduction techniques that produce transient signals, such as LA, ETV, FI and chromatography, more than one element may be determined at any one time. For liquid chromatography utilizing a mobile phase with a high dissolved salt content, coupling with ICPMS may lead to additional problems. As well as potentially blocking the nebulizer (if an appropriate one is not used) and the injector of the torch, blocking of the oriﬁce of the sampler cone may also occur. Inevitably, this will lead to signiﬁcant signal drift, until blockage is complete, at which point no signal will be obtained. A similar problem arises for the introduction of organic solvents. The solvent will pyrolyze within the plasma and will produce large quantities of soot. Whereas in ICP-OES this soot will pass harmlessly to waste via the fume extraction system, with ICP-MS instrumentation it may clog the sampler cone. In addition, the ion lens system also becomes dirty and the instrument will have to be dismantled so that it can be cleaned. The problems arising from soot deposition can be overcome by introducing oxygen (3 –5% v/v) into the nebulizer gas ﬂow. This turns the pyrolysis into an oxidative combustion process, and so the soot is burned off as carbon dioxide, thereby preventing sampler cone blockage. It should be noted, however, that the amount of oxygen used is critical. If too much oxygen is introduced, then the nickel sampler cone itself becomes oxidized away. This process can occur rapidly and a new cone can become unusable within a few minutes. Platinum or platinum tipped sampler cones are also available, and these tend to be more robust and resilient to oxidative attack, but are obviously substantially more expensive. Some workers have coupled GC with ICP-MS [90], but the coupling requires a heated transfer line that has to be constructed to enable safe and simple coupling. A commercial GC –ICP-MS instrument has now been produced, so the overall coupling is more robust. Some workers have also coupled capillary electrophoresis (CE) with ICPMS. The ﬂow rate through a CE instrument is typically at the low ml min21 level, or perhaps even nl min21. A specialized coupling is therefore required to make sure that the ﬂow rate of the CE and the uptake rate of the ICP-MS are compatible, often achieved using a micro-ﬂow nebulizer or a DIN [91]. Occasionally, a gas inlet is used to prevent suction from the nebulizer destroying the chromatographic separation by drawing the sample through at an accelerated rate. Since the injection volume is exceptionally low (again at the nl or ml range), the concentrations detected are normally at the mg ml21 range, although the absolute amount is at the pg or fg level. There are therefore very few applications for this coupling and its use is far from routine. Other sample introduction methods, including LA, ETV and chemical vapor generation, share the same advantages and disadvantages as discussed for analyzes by ICP-OES.

159

L. Ebdon et al.

5.6.7

Figures of merit

The limits of detection obtainable by conventional nebulization ICP-MS are usually at least two orders of magnitude lower than those obtainable by ICPOES. The LOD will depend on a number of factors, including the ionization energy of the particular analyte, the number of isotopes the analyte has (if an element has six or seven isotopes, the signal will be split between these and hence sensitivity will be less than for an element with only one isotope), the acquisition (integration) time and potential interferences. In addition, the operating parameters and the type of instrumentation used will also have a great effect. For a quadrupole based ICP-MS, approximate LODs are shown in Table 5.4. At a low-resolution setting, a magnetic sector instrument may improve these by at least an order of magnitude. Obviously, for sample introduction methods that give increased transport efﬁciency to the atom cell (ETV and chemical vapor generation), the LODs shown in the table can also be improved by over an order of magnitude. Under standard conditions, the linear dynamic range spans ﬁve or six orders of magnitude. If, however, both pulse counting and analogue modes are used, then the linear range may be extended to eight or even nine orders of magnitude. It should be noted though that the standards still have to be prepared in an appropriate range for the individual analytes within the sample. Sample throughput will again depend on the method of sample introduction and the time of acquisition/number of replicate measurements, but for conventional nebulization and a 10 s acquisition (integration time) for each of three replicates, potentially up to 50 or 60 samples may be analyzed in an hour. The number will also be affected by the speed of washout from the spray chamber, and so a fast clearing spray chamber will enable more samples to be analyzed per hour than a slower one. Since the instruments are simultaneous (or quasi-simultaneous), a large number of analytes may be determined simultaneously and an enormous amount of data may be collected in a short period of time. Precision will depend on the application. For conventional nebulization a precision better than 1% RSD may be obtained. For isotope ratio and ID measurements, precision would normally be expected to be better than 0.1% RSD. For other sample introduction techniques, such as LA or ETV, precision will depend on the homogeneity of the sample rather than the detection technique. 5.7 5.7.1

ATOMIC FLUORESCENCE SPECTROMETRY Introduction

Atomic ﬂuorescence spectrometry (AFS) is theoretically applicable to all of the commonly determined analytes. Modiﬁcations to standard ﬂame instruments

160

Detection methods for the quantitation of trace elements TABLE 5.4 Limits of detection for quadrupole ICP-MS under optimum conditions and using conventional nebulization Analyte

LOD (mg l21)

Analyte

LOD (mg l21)

Ag Al Au Ba Be Bi Ca Cd Co Cr Cs Cu Dy Er Eu Fe Ga Gd Ge Hf Hg Ho In Ir K La Li Lu Mg Mn

0.005 0.05 0.005 0.001 0.001 0.001 0.5 0.005 0.001 0.005 0.001 0.005 0.01 0.001 0.001 0.05 0.001 0.001 0.05 0.005 0.001 0.001 0.001 0.005 0.5 0.05 0.001 0.001 0.05 0.0004

Mo Na Nb Nd Ni P Pb Pd Pr Pt Rb Ru Sb Sc Se Si Sn Sr Ta Tb Te Th Ti Tl Tm U V W Y Zn

0.005 0.05 0.005 0.001 0.005 0.5 0.001 0.005 0.001 0.005 0.001 0.005 0.005 0.05 0.05 0.5 0.005 0.001 0.005 0.001 0.05 0.001 0.05 0.001 0.001 0.001 0.005 0.005 0.001 0.005

may be used to obtain one capable of detecting atomic ﬂuorescence. Despite its general applicability, in recent times AFS has been used almost entirely for the vapor generating elements. Indeed, commercial instrumentation has been produced that specializes in detecting As, Sb, Se and Te and another that detects Hg. The specialized commercial AFS detectors are relatively cheap, costing ,US $5000, although fully automated systems are also available at

161

L. Ebdon et al.

greater cost. Some research papers have been published concerning laser induced ﬂuorescence (LIF) or laser excited atomic ﬂuorescence (LEAF). These, however, are research methods at present and are not used routinely. A review of LEAF spectrometry has been published recently [92].

5.7.2

Theory

The theory of AFS may be found elsewhere in the literature [93]. Brieﬂy, radiation from an intense light source (line sources are used rather than continuum ones, although high intensity light emitting diodes that have a bandwidth of 20–40 nm may also be used) is used to excite the analyte which, upon relaxation back to a lower energy state, emits light of discrete wavelengths, depending on the transition involved. The intensity of the light source has a large impact on the sensitivity, as the ﬂuorescence intensity is proportional to the intensity of the source. Standard HCLs may be used, but greater sensitivity is obtained from boosted HCLs. A laser would provide the most intense source and a number of these have been used for this purpose, including standard YAGs, diodes, dye lasers and optical paramagnetic oscillators but, apart from diode lasers, most are difﬁcult to operate and costly to maintain. Atomic ﬂuorescence is exceptionally speciﬁc, ensuring that spectral interferences are minimal.

5.7.3

Instrumentation

The instrumentation used can be basically the same as for F-AAS, although the light source must be positioned at a right angle to the detector so that emission from the lamp is not detected as ﬂuorescence. Since AFS is so speciﬁc, it does not require a complex line isolation device such as a monochromator. Instead, simple ﬁlters will sufﬁce, although some high throughput multi-reﬂectance (interference) ﬁlters have also been used. These reportedly transmit 80% of the wavelengths of interest whilst virtually eliminating background noise. The atom cell in commercial AFS detectors is usually an argon/hydrogen diffusion ﬂame. This is a low temperature ﬂame that is used to dissociate the hydrides of these analytes. Both argon and hydrogen have low quenching crosssections for ﬂuorescence. For the commercial Hg detector, a simple quartz cell or open argon sheathed chimney is used. Since atomic vapor is introduced, there is no need for a heat source. Detection is usually with a PMT. It should also be noted that atomic ﬂuorescence has also been achieved using an ICP as an atom cell [94]. Although a commercial instrument was marketed brieﬂy, this too has only really been of research interest. A review of ICP-AFS has been produced by Greenﬁeld [95].

162

Detection methods for the quantitation of trace elements

5.7.4

Sample introduction

The majority of AFS techniques utilize chemical vapor generation to introduce the sample. The usual problems with chemical vapor generation are observed and therefore optimization of the generation chemistry to decrease interferences and to transform non-vapor forming species into ones that can is required. Often, a membrane drier tube is used to prevent the ingress of water vapor into the atom cell, since its presence may lead to light diffraction, quenching of the ﬂuorescence and possible interference. Chromatography, coupled with chemical vapor generation and AFS detection, has been used frequently as an alternative to ICP-MS detection, because the LOD is comparable whilst the overall cost of the instrumentation is substantially less. An example of HPLC – HG-AFS that also incorporated an on-line microwave transformation of inert species into forms that can generate a hydride has been published by Gomez-Ariza et al. [96]. Several studies have coupled chromatography directly with atomic ﬂuorescence. As an example, Puskel et al. used a specialized type of nebulizer (a hydraulic high pressure nebulizer) to introduce assorted selenium species [97]. 5.7.5

Interferences

Since the majority of applications of AFS utilize chemical vapor generation sample introduction, many of the interferences observed occur in the vapor generation step. Methods of overcoming these have been described previously. Once the vapor enters the atom cell, several types of interference may occur, including quenching by molecular gases (and other species), leading to a dramatic reduction in sensitivity. To minimize or prevent this, commercial AFS detectors use a gentle argon purge ﬂow. Similarly, if water vapor enters the atom cell, quenching or diffraction/scattering of the light may occur. The ideal ﬂame is the argon/hydrogen diffusion ﬂame, but this has a temperature that is too low to prevent chemical interferences and is another reason why the vapor generation technique is the preferred method of sample introduction. 5.7.6

Figures of merit

Atomic ﬂuorescence, especially when the sample is introduced by a chemical vapor technique, is exceptionally sensitive. For mercury, the detection limit is reported to be less than 1 ng l21 and for other analytes, such as As, Se, Sb and Te, the LOD is approximately 10 ng l21. The technique has a linear range spanning ﬁve orders of magnitude. Sample consumption will depend on the mode of chemical vapor generation used, i.e., batch or continuous; but is likely to be several milliliter. Precision is comparable to other common detection techniques and is typically better than 5% RSD.

163

L. Ebdon et al.

5.8

OTHER ATOMIC ABSORPTION, EMISSION AND FLUORESCENCE METHODS OF DETECTION

There are a number of other detection methods that have been used to determine trace analyte concentrations, although many are either research methods or have fallen virtually into disuse. 5.8.1

Microwave induced plasma

The microwave induced plasma (MIP) is the most commonly used of the other methods. This plasma is formed from microwave radiation and usually helium as the support gas, although other gases have also been used. The normal helium MIP is not a robust plasma and analytes must usually be introduced in a gaseous form since the plasma will be extinguished by the presence of any solvent. Recently, however, a very high power MIP (up to 1500 W) has been sustained while liquids were aspirated [98]. Since helium has much higher ionization energy than argon, the MIP is capable of detecting several analytes with greater sensitivity than argon based plasmas. Examples include the halogens (including ﬂuorine), sulfur and nitrogen. The MIP is used mainly in the atomic emission mode, although in the reference given above, a mass spectrometric detection method was used. In this latter mode, a helium-based plasma is useful because the argon polyatomic interferences observed in argon plasmas are largely eliminated. Detection limits were at the ng level. For MIP-AES, a number of systems have been used. These include different types of microwave cavity, e.g., Beenakker, slab-line and surfatron, and different types of line isolation and detection devices, e.g., Czerny –Turner monochromators, Rowland circle style polychromators and oscillating bandpass ﬁlters. The development of a commercial instrument that has coupled together GC and MIP-AES has ensured that this has become the most common method of sample introduction. The chromatography coupled with the MIP detection means that the vast majority of analyzes performed are speciation-based techniques. Speciation analyzes with MIPAES detection, and many of the fundamentals of the technique, have been reviewed recently [99]. The technique yields LODs in the range of 0.1 –5 pg s21 and linear ranges extend over four orders of magnitude. Sample throughput depends on the length of time required for the chromatography to be complete, but is unlikely to exceed 10 analyzes per hour. The sample throughput will also be dependent upon whether a temperature gradient was used to achieve the separation. If the chromatography is not isothermal, the GC oven will require time to cool to its starting temperature before another sample can be introduced. Precision is typically around 5% RSD.

164

Detection methods for the quantitation of trace elements

5.8.2

Direct current plasma

The direct current plasma (DCP) is an economical plasma to operate because it uses approximately 8 l min21 of argon and runs using a power of approximately 1000 W. However, it suffers very badly from interferences caused by easily ionized elements. Most applications require blanks, samples and standards to be spiked with high concentrations of lithium or barium to offset these effects. Although it is stable to the introduction of both aqueous and organic solvents, its use has declined almost to the point of non-existence. Commercial instrumentation produced nearly two decades ago used a similar line isolation device as found in many ICP instruments (an echelle spectrometer) and therefore had excellent resolution. Sample throughput and consumption is similar to that with the ICP. Precision is similar to that obtained with F-AES, but LODs are usually superior, especially for the hard to excite elements. In one relatively recent paper, the DCP was used to determine B in soils [100]. The linear dynamic range was reported as having ﬁve orders of magnitude and the LOD was 0.1 mg l21. 5.9 5.9.1

SECONDARY ION MASS SPECTROMETRY Introduction

Secondary ion mass spectrometry (SIMS) is based on the mass spectrometry of ionized particles that are emitted when a surface, usually a solid, is bombarded by energetic primary particles, which may be electrons, ions, neutrals or photons. The emitted or “secondary” particles will be electrons, neutral species, atoms or molecules or atomic and cluster ions. The large majority of species emitted are neutral, but it is the secondary ions that are detected and analyzed by the mass spectrometer. This is a process that provides a mass spectrum of a surface and enables a detailed chemical analysis of a surface or solid to be performed. The ﬁrst mention of sputtered secondary ions in the literature was made in 1910 by J.J. Thomson [101]. The ﬁrst regular secondary ion mass spectrometer was based on a patent by Herzog in 1942 [102,103], and the ﬁrst successful studies of surface compositions using mass-analyzed sputtered ions were made by several teams in the early 1950s [104,105]. An accelerated development of the ﬁeld was stimulated by new efﬁcient designs of narrow-beam primary ion columns [106] and of ion optics for “direct” imaging [107]. The late 1960s saw the emergence of the ﬁrst commercial instrumentation [108,109] and the coining of the SIMS acronym [110]. Static SIMS emerged as a technique of potential importance in surface analysis in the late 1960s and early 1970s as a consequence of the work of Benninghoven and his group in Mu¨nster [110]. Whilst the SIMS technique is basically destructive, the Mu¨nster group demonstrated that using a very low

165

L. Ebdon et al.

primary particle ﬂux density (,1 nA cm22), spectral data could be generated in a very short time scale compared to the lifetime of the surface layer. The information so derived would be characteristic of the chemistry of the surface layer because, statistically, no point on the surface would be impacted more than once by a primary particle during the analysis. Today, SIMS has an acknowledged place among the major techniques of surface analysis and microstructural characterization of solids. Proﬁling or other applications of SIMS that are not static are referred to as dynamic. Dynamic SIMS has found extensive application throughout the semiconductor industry where the technique had a unique capability to identify chemically the ultra-low levels of charge carriers in semiconductor materials and to characterize the layer structure of devices. Secondary ion mass spectrometry is particularly noted for its outstanding sensitivity of chemical and isotopic detection. Quantitative or semi-quantitative analysis can be performed for small concentrations of most elements in the periodic table, including the lightest. However, the high versatility of SIMS is mainly due to the combination of high sensitivity with good topographic resolution, both in depth and (for imaging SIMS) laterally. Its generally superior trace element sensitivity, capability for spatial resolution in three dimensions and for isotope measurements, as well as potential for identiﬁcation of chemical compounds in many cases, make SIMS the preferred method for the solution of an analytical problem. Deﬁciencies, however, still exist in the capability of SIMS for quantitative elemental analysis compared to other surface techniques (Auger, X-ray photoelectron spectroscopy, electron microprobe techniques, etc.). These deﬁciencies can be traced to the extreme dependence of relative and absolute secondary ion yields on several parameters. Among these the following are the most important: † † † † † † †

matrix effects; surface coverage of reactive elements; angle of incidence of primary beam with respect to the sample surface; angle of emission of detected ions; mass-dependent transmission of the mass spectrometer; energy band-pass of the mass spectrometer; dependence of detector efﬁciency on element.

Quantitative elemental SIMS analysis poses a twofold problem. Firstly, spectral interpretation, namely, the extraction of total detected isotopic ion currents assignable to elemental and molecular ions from a complete SIMS spectrum of the sample; secondly, spectral quantiﬁcation, namely the calculation of elemental concentration from total isotopic elemental (and molecular) ion currents. Difﬁculties in spectral interpretation are considerably reduced if high resolution mass analyzers ðM=DM . 3000Þ are used for mass analysis of secondary ions because most of the commonly occurring isotopic and molecular interferences (e.g., hydrocarbons, oxides and hydrides) can be resolved.

166

Detection methods for the quantitation of trace elements

5.9.2

Practical principles

A diagram of a SIMS instrument with a double-focusing mass analyzer is represented in Fig. 5.2. Secondary ion mass spectrometry is based on: † † †

bombardment of the sample surface by focused primary ions, with sputtering of the outmost atomic layers; mass spectrometric separation of the ionized secondary species (sputtered atoms, molecules, clusters) according to their mass-to-charge ratios; collection of separated secondary ions as quantiﬁable mass spectra, as indepth or along-surface proﬁles, or as distribution images of the sputtered surface.

The primary ions are normally produced by a duoplasmatron type of gas þ 2 þ þ þ source such as Oþ 2 , O , N2 , Ar ; by surface ionization as for Cs and Rb ; or by þ þ liquid-metal ﬁeld ion emission as Ga and In . The most common primary ions used are the oxygen ions, Csþ and Gaþ. The ions are accelerated and focused to a selected impact area on the specimen. The collision cascade following the incidence of a primary ion results in the implantation of the primary particle, reshufﬂing of some 50–500 matrix atoms, and emission of secondary particles, neutral or ionized. Secondary ions from the specimen are extracted into the mass spectrometer, which can consist of electric (ESA)/magnetic deﬂection ﬁelds or be of the quadrupole or time-of-ﬂight design (see Section 5.6.3.1). Secondary ions with a given mass-to-charge ratio and within a certain interval

Fig. 5.2. Diagram of double focusing SIMS (adapted from Ref. [111]).

167

L. Ebdon et al.

of kinetic energy are collected for pulse or current measurement, ion-optic imaging and data processing. The different ways of operating a SIMS instrument are presented in Fig. 5.3. In the microscope mode, a defocused primary ion beam (5 –300 mm) is used for investigating a large surface. In the microprobe mode, a focused primary ion beam (,10 mm) is used for investigating a very small portion of the surface and detecting inclusions in bulk material. The lateral resolution is deﬁned by the primary ion beam size. 5.9.3

Sensitivity and quantiﬁcation

Figure 5.4 shows schematically the types of analytical information that can be obtained by SIMS analysis. A SIMS spectrum normally shows mass peaks that are characteristic of the sputtered solid but affected by experimental factors. For instance, among these factors the following should be mentioned: type, intensity, energy and incidence angle of the primary ions; the transmission of the secondary ions and the selectivity for them in the mass analyzer; the type of detector. There are, effectively, two spectra: that of the matrix and that of the impurities. The task of analytical SIMS is to quantify the secondary ion currents, that is to convert the intensity of one or several peaks characteristic of an element to

Fig. 5.3. Operating modes of SIMS (adapted from Ref. [112]).

168

Detection methods for the quantitation of trace elements

Fig. 5.4. Analytical information obtainable from SIMS analysis (adapted from Ref. [113]).

its corresponding concentration ce . Assuming that a primary ion beam with a current density, ip ; strikes the sample; collision cascades are initiated, resulting in, among other things, the emission of secondary ions, which are partially detected with an instrument transmission, h, as a mass spectrum of ions from an analyzed area, A. The detected positive or negative current of an ionic species M at the mass number m will be: IM ¼ Ip SPM hM gM bM ce

ð5:1Þ

where Ip is equal to ip A and P is the probability that the particle (atomic or molecular) will emerge as the last step of the sputtering and recombination cascade. S is the sputtering yield (secondary particles per primary ions), gM is the positive or negative ionizability of M (ions per atom or molecule), and bM is the isotopic abundance of M in the element. 5.9.3.1 Absolute sensitivity In a situation where the prime goal is to detect trace elements of as low a concentration as possible, without consideration of sample consumption and analytical volume (e.g., in bulk analysis), the suitable ﬁgure of merit is the detected secondary ion current of an element E per unit of atomic concentration cðAÞ; that is the absolute sensitivity Sa ðEÞ : Sa ðEÞ ¼ N q ðEÞ=cðE

5:2Þ

where N q is the detected current (in counts per second) of element E in charge state q.

169

L. Ebdon et al.

5.9.3.2 Practical sensitivity The practical sensitivity, Sp ðEÞ; takes into account the fact that in different analytical situations different primary beam currents may be appropriate: Sp ðEÞ ¼ N q ðEÞ=Ip cðE

5:3Þ

This deﬁnition of practical sensitivity does not provide a ﬁgure of merit independent of material consumption. The same value would be obtained on different samples for the element E if, at the same primary beam current, the secondary ion currents of element E are identical, even if X sputters much faster than Y. 5.9.3.3 Useful yield If the amount of sample is limited or the sampling volume has to be small, the appropriate ﬁgure of merit is the useful yield, tu : It is deﬁned as the number of detected secondary ions/s, N q ðEÞ; of element E per number, NðEÞ; of sputtered E (atoms/s) from the same sampling volume: tu ðAÞ ¼ N q ðEÞ=NðEÞ ¼ Sp ðEÞ=Ytot

ð5:4Þ

Using the previously introduced ﬁgures of merit, the fundamental SIMS formula can be alternatively written as: N q ðEÞ ¼ Sa ðEÞcðEÞ ½cps ¼ Sp ðEÞNp cðEÞ ½cps ¼ tu ðEÞNp Ytot cðEÞ ½cps

ð5:5Þ

where Sa is measured in counts per second (cps) and dimensionless units have to be chosen for Sp and tu : When Sa ; Sp or tu are known, Eq. (5.5) provides a simple means for calculation of elemental concentration, cðEÞ; from the measured secondary particle current, N q : 5.9.3.4 Sensitivity factors Quantitation in SIMS can be achieved by external standards or by utilizing the concept of sensitivity factors. Under scrupulously reproducible conditions of analysis, and using external standards with composition and microstructures not too different from the analyzed samples, useful calibration factors may be obtained. However, long-term instabilities in analysis (instrumental drift, changes in primary beam conditions, vacuum effects, crystalline effects) make the use of absolute sensitivity factors hazardous. It is generally found to be both very feasible and more reliable to utilize the simultaneously measured ion current, IR ; of a matrix reference element, R. It has been found that relative sensitivity factors (RSFs) remain practically constant within quite wide ranges of concentrations, i.e., the differences are only weakly dependent on concentration. Excellent quantitation with RSFs has been reported, for example, for steels, binary alloys, glasses and semiconductors. The dominant sources of variation and irreproducibility in absolute and RSFs are connected with the ionizability of the elements.

170

Detection methods for the quantitation of trace elements

5.10

GLOW DISCHARGE MASS SPECTROMETRY

5.10.1 Introduction Glow discharge mass spectrometry (GD-MS) consists of the coupling of a glow discharge atomization/ionization source with a mass spectrometer. As noted earlier, the relative simplicity of mass spectra compared with optical spectra makes mass spectrometry an attractive alternative to optical spectrometry for trace element analysis. Moreover, mass spectrometry permits the coverage of essentially the entire periodic table and, since the spectral background can be very low, detection limits are usually 2–3 orders of magnitude better by mass spectrometry than for optical atomic emission using a glow discharge. For over 50 years glow discharges have been known as ion sources for mass spectrometry. The capability of generating a stable analyte ion population directly from a solid sample, thereby precluding the problems of dissolution, dilution and contamination that may arise for techniques requiring solution samples, makes the glow discharge an attractive ion source for elemental mass spectrometry of solids. The ability to obtain isotopic information across the periodic table down to ng g21 detection limits, along with the developments of improved mass spectrometers with more reliable data acquisition and control systems, has made GD-MS a powerful tool, not only for research laboratories but also for routine applications. A wide variety of analytical glow discharge geometries have been investigated as ion sources. Most GD sources, particularly the commercial versions, have used a direct insertion probe that permits certain ﬂexibility in sample shape, although pins or discs are normally used. In this conﬁguration, the sample serves as the cathode of the glow discharge system and the cell housing as the anode. Ions are sampled from the negative glow region through an exit oriﬁce. In Table 5.5, a comparison of the different sources is given. Hollow cathode glow discharges were coupled with a magnetic sector analyzer in preliminary investigations of analytical GD-MS [114,115]. Commercial instruments employ a modiﬁed coaxial cathode geometry [116,117]. This is also the most widely characterized glow discharge ion source. Whereas different glow discharge ion sources have not exhibited any signiﬁcant performance differences, different methods of powering the sources show speciﬁc performance differences. DC-powered sources are the most common, even though RF-powered sources have been studied [118] and applied as well as the pulsed sources [119]. The most widely used commercial GD-MS instrument is the VG9000 that consists of a DC-powered source, a double-focusing mass analyzer of the reverse Nier-Johnson geometry and Daly and Faraday cup detectors. Its cost is around US $600,000.

171

172 TABLE 5.5 Comparison of glow discharge sources Voltage (V)

Current (mA)

Pressure (Torr)

Cathode

Advantages

Disadvantages

Hollow cathode

250 –500

10– 100

0.1 –1.0

High sputter Intense ion beam Useable for powders

Charge exchange Complicated sample geometry

Grimm

500 –1000

25– 100

1 –5

23 mm deep cylinder with 5 mm diameter base 6.5 mm diameter circle

Only ﬂat samples

Jet-enhanced

900

28

2.5

12 mm diameter circle

Coaxial cathode

800 –1500

1 –5

0.2 –2.0

1.5 –2.0 mm diameter £ 4– 8 mm long rod

Depth proﬁling Easy for compacted powders High sputter rate Easy for compacted powders Useable for various sample shapes Ionization dominated by Penning process

Only ﬂat samples Higher discharge gas ﬂow rate Powders need to be converted into solid samples

L. Ebdon et al.

Source

Detection methods for the quantitation of trace elements

5.10.2 Glow discharge processes A glow discharge is a partially ionized gas consisting of approximately equal concentrations of positive and negative charges plus a large number of neutral species. It consists of a cathode and anode immersed in a low-pressure (< 0.1– 10 Torr) gas medium. Application of an electric ﬁeld across the electrodes causes breakdown of the gas (normally one of the rare gases is used, typically argon) and the acceleration of electrons and positive ions towards the oppositely charged electrodes. Detailed description of the phenomena can be found in Refs. [120 –122]. As an ion source for elemental mass spectrometry, the glow discharge is characterized by two attractive attributes, cathodic sputtering and Penning ionization, that are inherent to its operation. Cathodic sputtering generates a representative atomic population directly from the solid sample. Penning ionization selectively ionizes these sputtered atoms, permitting detection on the basis of their characteristic mass-to-charge ratios by mass spectrometry. 5.10.2.1 Atomization Cathodic sputtering is the phenomenon that makes a glow discharge useful in analytical spectrometry, providing the means of obtaining directly from a solid sample an atomic population for subsequent excitation and ionization. The sputtering involves directing an energetic particle onto a surface where, after collision, it transfers its kinetic energy in a series of lattice collisions. Atoms near the surface can receive sufﬁcient energy to overcome the lattice binding and be ejected, generally as neutral atoms with energies in the range of 5–15 eV. The bombarding particles are normally ions, easily accelerated by electrical ﬁelds. The sputter yield, deﬁned as the number of ejected atoms per bombarding ion, depends critically on the mass and energy of the incoming ions. Under the operating conditions of most analytical glow discharges, the sputter yield can be described as a function of kinetic energy and mass of the bombarding atom as well as of the lattice binding energy and mass of the target atoms. A related value is the sputtering rate, namely the number of target ions sputtered per unit time. This value is determined by the discharge operating current as well as the factors affecting the sputter yield. 5.10.2.2 Ionization The glow discharge sputtering can introduce into the plasma a representative population of the sample (cathode) ions. A fraction of them needs to be ionized for further elemental analysis by mass spectrometry. The discharge must then act as ionizing medium and must, of course, sustain itself. The fact that GD-MS does not utilize optical transitions of the analyte atoms, rather the mass-to-charge ratio of the atoms that have been ionized, shifts the emphasis from excitation mechanisms to ionization mechanisms, speciﬁcally

173

L. Ebdon et al.

simplifying, to some extent, the relationship between analyte signal and analyte concentration in the sample. Figure 5.5 shows schematically the processes in a GD [123] and in Table 5.6 ionization processes in glow discharges are summarized. Whereas we can assume that atomization does not differ signiﬁcantly between elements in a given matrix, we cannot assume the same to be true for ionization. Therefore, RSFs used for quantitative analysis are most likely controlled by differences in the probability of ionization among the elements. The RSF of an analyte element, E, is the ratio of its sensitivity to the sensitivity of some reference element. Sensitivity is deﬁned as the intensity (I) of the signal per unit of concentration (C): RSFE=R ¼ ðIE =CE Þ=ðIR =CR

5:6Þ

The RSFs consider the contributions arising from instrumental factors, such as ion transmission and sensitivity, and glow discharge processes, such as differential atomization and differential ionization. The dominant contribution is related to the glow discharge processes and varies from sample to sample. 5.10.2.3 Quantiﬁcation The mass spectrum obtained by GD-MS can be used directly for a semiquantitative measurement of the sample composition. One method is based on the ion-beam ratio (IBR) [124]. In this procedure, the ion signals for all

Fig. 5.5. Processes in a glow discharge (Reprinted from Spectroscopy Europe, 15(3) (2003) 15).

174

Detection methods for the quantitation of trace elements TABLE 5.6 Ionization processes in the glow discharge Electron impact ionizationa Penning ionizationb Associative ionizationb Symmetric charge exchangeb Asymmetric charge exchangeb

A þ e2 ! Aþ þ 2e2 Arm þ X ! Ar þ Xþ þ e2 Arm þ X ! ArXþ þ e2 Aþ þ A ! AþAþ Aþ þ B ! AþBþ

a

Collisions of the ﬁrst kind. Collisions of the second kind.

b

ionized sputtered species are summed and then the ratio of the ion signal for individual species is calculated, which corresponds to the concentration of the species in the bulk. Since the IBR depends on the sensitivity of the different elements, which varies by less than a factor of 10, this method can only provide reliable means for semiquantitative analysis. The signal intensity of the plasma species is inﬂuenced by several factors. Among these are: sample composition, matrix type, discharge power, cathode geometry, cooling effects, discharge gas, source pressure, ion transmission, the type of mass spectrometer and the detection system. Because of all this, for quantitative analysis, the use of standards is required for calibration. This can be performed in two ways. The ﬁrst consists of the construction of a calibration curve, based on a set of similar standards [125,126]. The second possibility is based on the analysis of a reference material as similar as possible in composition and behavior to the unknown sample, which allows the calculation of the RSFs [127]. Since suitable certiﬁed standards are not always available, powdered samples may be doped with an element of known concentration to be used as an internal standard.

5.10.3 Applications to trace element analysis GD-MS has taken the place of spark source mass spectrometry (SS-MS) for the analysis of trace elements in solid samples. In comparison to SS-MS, GD-MS presents many advantages, for instance, a simple source producing a stable supply of low energy ions characteristic of the sample and minimal matrix effects. In a DC powered GD-MS instrument, the samples must be conductive; therefore, bulk metals are the most ideal samples even though non-conductive samples can also be analyzed. In this case, the samples need to be mixed with a binder material [127] (Ag, Ti or Ta) or the technique of the secondary cathode can be applied [128]. Sample spectra may be obtained in a short time (min) and rapid qualitative analysis can be performed by the examination of the

175

L. Ebdon et al.

isotopic lines. Quantitative analysis can also be achieved, as previously explained. Semiconductors are another important category of bulk samples that can be analyzed by GD-MS. The electrical properties of these materials are critically dependent on the intrinsic and doped levels of impurities, making it essential to know, not only qualitatively their chemical composition, but also the concentration level of each element. Even extremely low concentrations of a speciﬁc element can alter semiconductor properties. GD-MS is always a surface analysis technique, even though it permits measurement of bulk concentrations. That is, the sputtering process central to the glow discharge acts as an atomic mill that regularly erodes away the surface of the bombarded sample. Whatever atoms are sputtered away from the surface are measured and, because GD-MS consumes signiﬁcant quantities of material (up to milligrams per minute), these sequential layer analyzes combine to yield an averaged composition that is typical of the bulk concentration. By slowing down the ablation process limiting measurement to a shorter duration, data indicative of the surface concentrations can be obtained. GD-MS and its optical analogue have found considerable application for the analysis of layered samples. Environmental samples can also be analyzed by GD-MS. In these cases the samples need to be compacted with or without conducting material [129]. Where a binder of conducting material is not added during the compaction of the samples, a secondary cathode has been used for their analysis [126]. 5.11

X-RAY FLUORESCENCE SPECTROMETRY

5.11.1 Introduction X-ray spectrometric techniques have been very useful in providing important information for theoretical physicists and have found increasing exploitation in the ﬁeld of material science characterization. Today, most stand-alone X-ray spectrometers use X-ray excitation sources rather than electron excitation. X-ray ﬂuorescence spectrometers typically use a polychromatic beam of short wavelength X-radiation to excite longer wavelengths, characteristic lines, from the sample to be analyzed. In modern X-ray spectrometers, single crystals are used to isolate a narrow energy band from the polychromatic radiation excited from the sample. This method is called “wavelength” dispersive spectrometry. The other possibility is to use a proportional detector to isolate a narrow energy band from those obtained from the excited sample. This method is called “energy” dispersive spectrometry. Since the relationship between emission wavelength and atomic number is known, isolation of individual characteristic lines permits the unique identiﬁcation of an element and elemental concentrations can be estimated from characteristic line intensities [130].

176

Detection methods for the quantitation of trace elements

X-ray ﬂuorescence spectrometry provides the means for the identiﬁcation of an element by measurement of its characteristic emission wavelength or energy. The quantitative estimation of an element is possible by ﬁrst measuring the emitted characteristic line intensity and then relating this intensity to the elemental concentration. A beneﬁt of using X-ray emission spectra for qualitative analysis is that, because the transitions arise from inner orbitals, the effect of chemical combination or valence state is almost negligible. 5.11.2 Instrumentation Several different types of sources have been employed for the excitation of characteristic X-radiation, including those based on electrons, X-rays, g-rays, proton and synchrotron radiation. By far, the most common source today is an X-ray photon source. This source is used in primary mode in the wavelength and primary energy dispersive system, and in secondary ﬂuorescer mode in secondary target energy dispersive spectrometers. An X-ray detector is a transducer for converting X-ray photon energy into voltage pulses. Detectors work by a process of photoionization, in which interaction between the incoming X-ray photon and the active detector material produces a number of electrons. The current produced by these electrons is converted to a voltage pulse by capacitors and resistors, such that one digital voltage pulse is produced for each entering X-ray photon. The most important characteristics of the detector are proportionality and linearity [131]. In the case of wavelength dispersive spectrometers, a gas ﬂow proportional counter is generally employed for the measurement of longer wavelengths, and a scintillation counter is generally used for shorter wavelengths. Neither of these two detectors has sufﬁcient resolution to separate multiple wavelengths on its own, and has to be employed with an analyzing crystal. However, in the case of energy dispersive spectrometry, where no dispersing crystal is used, a detector of higher resolution must be used, generally the Si(Li) detector [132]. 5.11.3 Matrix effects In the conversion of net line intensity to analyte concentration, it may be necessary to correct for absorption and/or enhancement effects. As for absorption, primary and secondary absorption needs to be considered. Primary absorption occurs because all atoms of the sample matrix absorb photons from the primary source. Since there is a competition for these primary photons by the atoms making up the sample, the wavelength distribution of intensity of the photons available for excitation of a given analyte element may be modiﬁed by other matrix elements. Secondary absorption refers to the absorption of characteristic analyte radiation by the specimen matrix. As characteristic radiation is released from the sample in which it is generated, it is absorbed by all matrix elements in amounts relative to their mass attenuation coefﬁcient.

177

L. Ebdon et al.

Enhancement effects of the total absorption occur when a non-analyte matrix element emits a characteristic line of energy just in excess of the absorption edge of the analyte element. This means that the non-analyte element is able to excite the analyte, giving characteristic photons in addition to those produced by the primary continuum. This results in an enhanced signal from the analyte [133]. 5.11.4 Quantitative and trace analysis The simplest challenge for quantitative analysis is the determination of a single element in a known matrix. In this case, calibration curves can be obtained. When the matrix is unknown, quantitative analysis is based on the use of internal standards, addition of standards and use of a scattered line from the Xray source. The most complicated case is the analysis of all, or most, of the elements in a sample whose matrix is unknown. In this case, a full qualitative analysis would be required before any attempt is made to quantitate the matrix elements. Once the qualitative composition of the sample is known, one of the following three techniques can be applied: type standardization, inﬂuence coefﬁcient methods, fundamental parameter techniques. The last two methods require a computer for their application [134,135]. One of the problems with any X-ray spectrometer system is that the absolute sensitivity (i.e., the measured cps per% of analyte element) decreases signiﬁcantly as the lower atomic number region is approached. This is, above all, due to the fact that the ﬂuorescence yield decreases with the atomic number, the absolute number of useful long-wavelength X-ray photons from the radiation source decreases with increasing wavelength and absorption effects become more severe with increasing wavelength of the analyte. The X-ray ﬂuorescence method is particularly applicable to the qualitative and quantitative analysis of low concentrations of elements in a wide range of samples as well as to the analysis of elements at higher concentration in a small amount of sample. Moreover, X-ray ﬂuorescence is often used as a nondestructive qualitative method for a multi-element content evaluation prior to quantitative analysis with another method. X-ray spectrometric methods based on total reﬂection geometry [136] have gained widespread strength in the past decade, principally because of their detection power (92–10 pg, 100 pg ml21 relative), quasi matrix independent calibration (internal standard), multi-element capability and non-destructive nature. For trace element analysis the sample is prepared on a totally reﬂecting sample carrier as a small quantity of residue from solutions or ﬁne-grained suspensions from the evaporation of a solvent and forms a layer of a few microns thick. It is thus a micro technique, achieving its performance advantage as a consequence of the attenuation of background due to the high angle reﬂection geometry employed.

178

Detection methods for the quantitation of trace elements

5.12

UV/VISIBLE SPECTROPHOTOMETRIC AND CHEMILUMINESCENCE TECHNIQUES

5.12.1 UV/Visible spectrophotometric techniques Many metals form compounds or complexes that give rise to an ultraviolet or visible absorbance spectrum. It is therefore sometimes possible to react the metal ion within a liquid sample (or a solid sample that has previously undergone a digestion or dissolution stage) with a reagent that will selectively change the optical properties of the sample. Many of the basic components required for such an instrumental based technique are similar to those in an atomic absorption instrument. However, the light source is different. Rather than using a HCL that produces a spectrum containing a series of discrete lines, a continuum source produced by a tungsten –halogen lamp, for example, is used that has a broad band of light output. For UV applications (,340 nm), a deuterium lamp is usually used as a source. The sample cell is frequently a cuvette, manufactured from glass or plastic (for wavelengths above 340 nm) or quartz for the UV region. The sample may simply be poured into the cuvette for absorbance readings to be made. Instead of a cuvette, gas cells are also available. The monochromator is usually fairly basic, but operates in the same way as the atomic absorption spectrometers. Again, detection is usually via a photomultiplier tube. The theory behind molecular absorption is analogous to atomic absorption, i.e., analyte molecules absorb light and electronic transitions occur within the molecule. A greater emphasis on the theory may be found in the literature [137]. It should be noted, however, that molecular absorbance occurs over a wide band of wavelengths rather than a very narrow line, as in atomic absorption. This means that, in many cases, even if detection is not at exactly the most intense wavelength, some results may be obtained, although the system will not give optimal sensitivity. As with all absorbance techniques, the Beer–Lambert law (Eq. (5.7)) applies and, since this law is true over only a relatively small concentration range, the calibration will be linear over a range of perhaps two orders of magnitude. A ¼ 1CL

ð5:7Þ

where A is absorbance, 1 is the molar extinction coefﬁcient (also called the molar absorptivity constant), C is the concentration and L is the path length. It can be seen from Eq. (5.7) that, in addition to concentration, the absorbance is dependent on the path length and the molar extinction coefﬁcient. The path length is usually 1 cm for most liquid cuvettes, but may be substantially larger for gas cells. The molar extinction coefﬁcient is a number whose numerical value is different for each metal –complex system, but is regarded as being a constant for each. The higher the value of 1, the greater

179

L. Ebdon et al.

the absorption of light by the compound formed, and hence the better the sensitivity. One of the major problems with UV/Visible spectrophotometric methods is that of interferences. Often, more than one metal ion will combine with a complexing agent and the absorption spectra from the two species may overlap. This is shown diagrammatically in Fig. 5.6. It can be seen that even a small contribution from one interfering species could potentially lead to inaccuracy and since the interferences are additive, the presence of several concomitants could lead to large errors. Assuming that all of the potential interferences within a sample are known, it is possible to use simultaneous equations or mathematical algorithms to correct for the relative contributions from each. An example is the simultaneous determination of chromium and manganese in steel samples [1]. It should also be noted that pH may affect the absorption proﬁle of the analyte complex and it is thus often necessary to buffer the samples and standards to the same pH. If the analyte forms a very stable compound with another concomitant, then it may not be available to form the complex and a low analytical result would occur. It may occasionally be necessary to treat the sample in some way to selectively bind the interfering species or to destroy it. The linear range for these methods usually covers no more than two orders of magnitude. The LOD will depend on the molar extinction coefﬁcient, but for some of the more sensitive methods may be as low as 10–50 ng ml21. Sample throughput can be very rapid, with up to 5–10 samples per minute being determined. Although gaseous samples may be analyzed, the norm is for the sample to be in a liquid form. Since the method of detection is through light absorption, it is necessary for the samples to be free from particulate matter. This may be achieved either by centrifugation or by ﬁltering. Since the light beam in most instruments will pass through the bottom third of the cuvette, a volume of 1 ml is usually sufﬁcient for the measurement step. It is necessary to

Fig. 5.6. Overlap of absorption spectra.

180

Detection methods for the quantitation of trace elements

remember that the sample must often be combined with a known volume of complexing reagent and buffer prior to the measurement step, and therefore the actual amount of sample used may frequently be less than 1 ml. The UV/Visible instrumentation described thus far has concerned standalone spectrometers that may also be used to scan over a wavelength range so that absorption maxima may be identiﬁed. If a ﬂow-through UV/Visible detector is used, then transient signals may be detected. Such instrumentation may be used as a detector for chromatography, but is predominantly used for automated high throughput analyzes, e.g., with FI techniques. If the sample, compleximetric reagent and buffer are mixed on-line and allowed to pass through the detector, a transient signal will be obtained which may be recorded using either a chart recorder (if peak height is to be measured) or an integrator. The same problems arise with interferences, but the major advantage of this approach is that the instrumentation is more portable and may be taken into the ﬁeld (assuming an adequate power supply is available) or aboard a ship. Therefore, savings in time are made and the disadvantages associated with the transport of samples back to a laboratory (with possible loss of analyte through adsorption or breakage of storage vessels, contamination from preserving agents, etc.) may be overcome. An example of such a method has been published by Hernandez et al. [138]. If a micro-column of resin is used to entrap the analyte prior to mixing with the compleximetric agent, then matrix removal and pre-concentration may be achieved. Sample consumption will be dependent upon the size of the sample loop, but is typically 100 –200 ml per injection, but this will increase markedly if a pre-concentration technique is used. If several analytes require determination and they all form a complex with a particular reagent, it may be possible to determine them using liquid chromatography coupled with UV/Visible detection. A recently published paper has outlined the determination of several analytes (Th, V, Bi, U, Hf and Zr) using a 10 cm column of neutral polystyrene loaded with dipicolinic acid and 1 M potassium nitrate as an eluent [139]. This application yielded LODs of substantially less than 1 mg l21 for several analytes. A similar study reported the determination of a selection of transition and heavy metal ions using a polystyrene –divinyl benzene column loaded with 4-chlorodipicolinic acid and an eluent of 0.25 mM chlorodipicolinic acid in 1 M potassium nitrate at pH 2.2 [140]. 5.12.2 Molecular ﬂuorescence and chemiluminescence detection Molecular ﬂuorescence detection is less commonly used because it is necessary to combine the analyte with a suitable reagent capable of ﬂuorescing, whilst ensuring that other metal species present in the sample do not. Despite these disadvantages, the technique can give LODs in the ng ml21 range, is more selective than absorption and, when a chromatographic technique is used to separate potential interferences from the analyte, excellent selectivity may be achieved and sample consumption need only be 50 –200 ml. A recent example of

181

L. Ebdon et al.

such an application compared ICP-MS and ﬂuorescence detection for Al species that had been separated using chromatography [141]. Chemiluminescence is a form of emission, but is produced from the energy originating from a chemical reaction rather than the absorption of light. Since it is related to ﬂuorescence, it possesses the same analytical advantages, including long linear ranges and very good sensitivity. Most methodologies use either FI or chromatography as a sample introduction method. As usual, FI methods tend to be very inexpensive and easily portable, although for this type of work they are also usually designed to determine only single analytes. Again, the sample must be in a liquid form, so soils and other solid materials must ﬁrst be digested. Typically, the analyte is isolated on a mini- or micro-column of a resin. This will often mean that the sample must be buffered to a speciﬁc pH and reagents added to ensure that other concomitant species are not retained since, on elution, these may react with the chemiluminescent reagents and potentially interfere. An example of a FI method has been published by Achterberg et al. [142], who determined Cu in seawater. This application also emphasized a method for overcoming another common problem experienced with this type of methodology. The presence of humic acids or other chelating compounds will usually mean that the analyte will complex with these in preference to the resin in the micro-column, resulting in poor analyte retention and its passage directly through the system undetected. The use of UV photolysis (possibly in the presence of hydrogen peroxide) prior to analysis can help destroy the organic material and will therefore render the Cu (or other analytes) available for detection. In this particular application, the UV digestion was performed on-line and, although less efﬁcient than batch irradiation, was sufﬁcient to enable successful determination whilst being substantially more rapid. The use of a UV digestion will inevitably increase the length of time required for the analysis, but even with a digestion time of 5 or 6 min, 5–10 samples per hour may be analyzed. As with all FI techniques, preconcentration is a possibility, and the pre-concentration process may limit the sample throughput further. An example of simple FI–chemiluminescence detection without the need for UV digestion has been reported by Bowie et al. [143]. These workers determined Fe in seawater by reducing Fe(III) to Fe(II) using sulphite, and then retaining/pre-concentrating the whole Fe content of the sample on a column of 8-hydroxyquinoline. On elution, the Fe was reacted with luminol for the chemiluminescence detection. Limits of detection were found to be sub-nM, precision was 3.2% RSD and the whole analytical cycle took 3 min, enabling a sample throughput of close to 20 samples per hour. The portability of the technique was demonstrated by the shipboard determination of Fe in the Atlantic. Chemiluminescence may also be used as a means of detection for liquid chromatography. The relative advantages of this are the same as those discussed for UV/Visible detection, with the added bonus of the extra sensitivity. Several papers have reported the determination of trace metals

182

Detection methods for the quantitation of trace elements

using chemiluminescence as a method of detection. Included amongst these is the determination of silver [144]. After elution from a cation exchange column, the silver was detected by a novel post-column reaction system involving the oxidation of luminol by peroxodisulfate. The system yielded a LOD of 0.5 mg l21. A similar paper by the same authors managed to speciate Cr(III) and Cr(VI) in fresh waters [145]. This system enabled LODs of 0.05 and 0.1 mg l21 to be obtained for Cr(III) and Cr(VI), respectively, whilst yielding a linear range of 0.1 –500 mg l21. Precision at the 10 mg l21 level was 5% RSD. Multi-element determinations are also possible using such systems. An example that yielded mg l21 LODs for several analytes in 15 min has been reported [146]. 5.13

ELECTROCHEMICAL METHODS

5.13.1 Differential pulse anodic stripping voltammetry A liquid sample is placed in a sample cell along with a suitable buffer, and then a hanging mercury drop electrode, a reference electrode (usually a saturated calomel electrode) and a platinum wire counter electrode are immersed in the sample. The sample is purged with an inert gas to de-oxygenate it because oxygen causes an interference effect. A negative voltage may then be applied to the mercury drop. The sample is stirred magnetically, and some of the positively charged metal ions in the sample will diffuse to the mercury drop and plate onto and diffuse into it. The period during which this occurs is termed the plating time. After this, a brief relaxation period occurs, followed by a period in which the potential applied to the mercury drop becomes increasingly less negative. Each analyte ion will be stripped or oxidized from the mercury drop at its own reduction potential and will re-enter the solution. As they re-enter the liquid phase, they are detected by the counter and standard electrodes and will appear as a series of peaks. The area under each of the peaks is proportional to the concentration of that particular ion in the sample. It is the length of the plating time that determines the overall sensitivity of the analysis. Short plating times will be insufﬁcient for many of the analyte ions within the sample to become signiﬁcantly accumulated in the mercury drop, whilst longer times will enable greater sensitivity to be obtained, but at the expense of decreased sample throughput. The overall sensitivity will be limited by the contamination within the buffer system and by time constraints. Detection limits substantially below 1 ng ml21 may readily be obtained for Cd, Cu, Pb and Zn using a plating time of just a few minutes, but plating times of over an hour are known, which yield exceptionally low LODs. The technique is multi-elemental, with approximately 20 metallic ions being detected. Despite this, the four aforementioned ions are most commonly determined by this method. Another advantage of the technique is that it is capable of determining different species of an element. An example has been the determination of assorted tin species,

183

L. Ebdon et al.

although an ion exchange procedure was also required for full speciation because many of the tin species produced only one peak [147]. Since only a very small portion of the analyte ions in the sample plate onto the mercury drop, the method is very vulnerable to errors caused by differences in plating times, stirring rates, temperatures and current density between samples and standards. It is this last potential problem that often leads to the method of standard additions being used to calibrate the process. The plating time is obviously the limiting factor for sample throughput, but if three standard additions are made to each sample, then rarely more than 5–10 samples may be analyzed per hour. Sample consumption is typically 1–10 ml, depending on the volume of the sample cell and on the volume of buffer/diluent added. Since many instruments possess a spiking port, the standard additions may be made to the same sub-sample. Precision is typically ,5% RSD. Care must be taken with the stirring of the sample, as the hanging mercury drop is easily dislodged. If stirring is too vigorous and the drop is dislodged it may be necessary (depending on whether plating has begun) to start the analysis again with a new sub-sample. For this reason, some workers prefer glassy carbon working electrodes and co-plating mercury from mercury (II) chloride added to the sample solution to produce the thin ﬁlm mercury electrode. Another drawback with the technique is that the presence of organic matter, that may complex with many of the ions, will lead to a decrease in the plating efﬁciency (i.e., the analytes are kept in solution as a complex and are not available for analysis). This will potentially lead to an underestimate of the true analyte concentration. It is possible to differentiate between the free ions and those complexed with organic matter if an analysis is ﬁrst made on an untreated sub-sample to yield the concentration of the free ions. A second analysis made on another subsample that has been treated with UV radiation, destroying the organic matter will yield the “total” concentration of the analytes. The presence of high concentrations of some metal ions may also lead to overlap of the peaks or to the formation of intermetallic compounds.1 5.13.2 Cathodic and adsorptive stripping voltammetry This is analogous to ASV, and may usually be achieved using the same instrumentation. Cathodic stripping voltammetry (CSV) is used less for metal ion determinations than ASV simply because there are fewer negatively charged metal ions, but metals may be adsorbed as their complexes (e.g., Ni with dimethylglyoxime). The obvious exceptions are the metalloids, e.g., arsenic, selenium, etc., and some of the transition elements (i.e., those that 1 Methods for the determination of the metals aluminum, cadmium, chromium, cobalt, iron, lead, nickel, uranium, vanadium and zinc in marine, estuarine and other waters by stripping voltammetry or concentration and atomic absorption spectrophotometry, HMSO, London, 1987.

184

Detection methods for the quantitation of trace elements

form negatively charged complexes). This technique shares many of the relative advantages and disadvantages of ASV. Again, speciation is possible with As(III) and As(V) species having been determined [148]. 5.13.3 Ion selective electrodes Ion selective electrodes (ISE) are so termed because they are selective for a speciﬁc ion. Some texts describe them as ion speciﬁc electrodes, but this is misleading because they are not speciﬁc. Most suffer from interferences in that they respond to the presence of other species in solution, although to a lesser extent than to the ion they are designed to detect. Examples of these interferences include Hþ, Csþ, Liþ and Kþ on the Na ISE and Zn2þ, Pb2þand Mg2þ on the Ca ISE. Additionally, the presence of organic matter, such as humic acids that may complex with the analytes of interest, may prevent them from being detected and hence, an underestimate of the true concentration will be made, although it should be noted that the electrode is truly responding to activity. Several of these electrodes exist, but they do not exist for every metallic element. Their response is based on the Nernst equation (Eq. (5.8)). E ¼ Eu þ

2:303 RT log ½ion zF

5:8Þ

where R is the gas constant, T is temperature, z is the charge of the species of interest, F is the Faraday constant and [ion] is the concentration of the analyte. As can be seen from Eq. (5.8), the response is dependent upon temperature and on the charge of the species under investigation, although at 258C and for a singly charged analyte, a change of 59 mV per decade of concentration is obtained. For a doubly charged analyte, e.g., Ca2þ at the same temperature, a change of 29.5 mV per decade should be generated if the electrode is Nernstian in response. Calibration curves can be plotted on semi-log paper, and can cover ﬁve or six orders of magnitude. Detection limits will depend on the individual ISE, but are often at the ng ml21 range; with copper, for example, the ISE is linear over the range 1028 –0.1 M (i.e., 6.4 £ 1024 –6350 mg l21). The electrodes are readily portable and may easily be used in conjunction with a data logger for unattended operation in the ﬁeld. Assuming the temperature of the water body is monitored simultaneously, a simple algorithm can be used to correct the data. It should be noted however, that any changes in ionic strength of the sample may lead to interferences (depending on the ISE). Therefore, for some ISEs, most samples (and standards) are mixed with total ionic strength adjustment buffer (TISAB), a pH buffer that also contains an inert salt to ﬁx the ionic strength. Normally, liquid samples are required and so solid samples have to be extracted, digested or dissolved such that the analytes are present in a solution. The volume of sample can be low (5 ml or less), but one of the advantages of the

185

L. Ebdon et al.

technique is that it does not consume any of the sample. Assuming that measures are taken to prevent contamination, the sub-sample used for this analysis may be used for other analyzes. The technique is also relatively rapid; once immersed in the sample, a brief equilibration time is allowed so that a stable signal is obtained, but this still allows two determinations to be made per minute. An example in which ISEs have been used to determine metal ions has been published by Vazquez et al. [149]. In this paper, Na, K and Ca were determined in wood pulp suspensions and the results compared with those obtained by ICP-OES and XRF.

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23

186

Vogel’s Textbook of Quantitative Chemical Analysis. Revised by J. Mendham, R.C. Denney, J.D. Barnes and M.J.K. Thomas, 6th ed., Prentice-Hall, 2000. In: L. Ebdon, E.H. Evans, A. Fisher and S.J. Hill (Eds.), An Introduction to Analytical Atomic Spectrometry. Wiley, Chichester, 1998. B. Welz and M. Sperling, Atomic Absorption Spectrometry, 3rd edn. Wiley-VCH, Weinheim, 1999. R.D. Beaty and J.D. Kerber, Concepts, Instrumentation and Techniques in Atomic Absorption Spectrophotometry. Perkin Elmer, Norwalk, USA, 1993. J. Dedina and D.L. Tsalev, Hydride Generation Atomic Absorption Spectrometry. Wiley, Chichester, 1995. X.C. Duan, R.L. McLaughlin, I.D. Brindle and A. Conn, J. Anal. At. Spectrom., 17 (2002) 227. A.G. Howard, J. Anal. At. Spectrom., 12 (1997) 267. P. Carrero, A. Malave, J.L. Burguera, M. Burguera and C. Rondon, Anal. Chim. Acta, 438 (2001) 195. R. Cornelis, X.R. Zhang, L. Mees, J.M. Christensen, K. Byrialsen and C. Dyrschel, Analyst, 123 (1998) 2883. S.P. Quinaia and M.D.C.E. Rollemberg, J. Braz. Chem. Soc., 8 (1997) 349. J.L. Gomez-Ariza, M.A.C. de la Torre, I. Giraldez, D. Sanchez-Rodas, A. Velasco and E. Morales, Appl. Organomet. Chem., 16 (2002) 265. C.C.Y. Chan and R.S. Sadana, Anal. Chim. Acta, 270 (1992) 231. T.L. Deng, Y.W. Chen and N. Belzile, Anal. Chim. Acta, 432 (2001) 293. R.C. de Campos, P. Grinberg, I. Takase and A.S. Luna, Spectrochim. Acta, Part B, 57 (2002) 463. Z.L. Fang, S.K. Xu and G.H. Tao, J. Anal. At. Spectrom., 11 (1996) 1. H. Matusiewicz, Spectrochim. Acta, Part B, 52 (1997) 1711. H. Matusiewicz and M. Kopras, J. Anal. At. Spectrom., 12 (1997) 1287. L. Ebdon, S. Hill and R.W. Ward, Analyst, 112 (1987) 1. L. Ebdon, S. Hill and R.W. Ward, Analyst, 111 (1986) 1113. J. Szpunar Lobinska, C. Witte, R. Lobinski and F.C. Adams, Fresenius’ J. Anal. Chem., 351 (1995) 351. R. Morabito, P. Massanisso and P. Quevauviller, TrAC Trends Anal. Chem., 19 (2000) 113. L. Ebdon, S.J. Hill and P. Jones, Analyst, 110 (1985) 515. K.J. Lamble and S.J. Hill, Anal. Chim. Acta, 334 (1996) 261.

Detection methods for the quantitation of trace elements 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58

M.B. Amran, F. Lagarde and M. Leroy, Mikrochim. Acta, 127 (1997) 19. C.W. Fuller, R.C. Hutton and B. Preston, Analyst, 106 (1981) 913. P. Vinas, N. Campillo, I.L. Garcia and M.H. Cordoba, Anal. Chim. Acta, 283 (1993) 393. P. Goodall, M.E. Foulkes and L. Ebdon, Spectrochim. Acta, Part B, 48 (1993) 1563. L. Ebdon and P. Goodall, J. Anal. At. Spectrom., 7 (1992) 1111. L. Ebdon, M. Foulkes and K. Sutton, J. Anal. At. Spectrom., 12 (1997) 213. N.J. Miller-Ihli, J. Anal. At. Spectrom., 4 (1989) 295. M.J. Cal-Prieto, M. Felipe-Sotelo, A. Carlosena, J.M. Andrade, P. Lopez-Mahia, S. Muniategui and D. Prada, Talanta, 56 (2002) 1. A.T. Persaud, D. Beauchemin, N.E. Jamieson and R.J.C. McLean, Can. J. Chem. Rev. Can. Chim., 77 (1999) 409. J.D. Nobrega, M.M. Silva, P.V. Deoliveira, F.J. Krug and N. Baccan, Quim. Nova, 18 (1995) 555. V.I. Slaveykova, F. Rastegar and M.J.F. Leroy, J. Anal. At. Spectrom., 11 (1996) 997. W. Slavin and D.C. Manning, Prog. Anal. At. Spectrosc., 5 (1982) 243. W. Slavin, D.C. Manning and G.R. Carnrick, At. Spectrosc., 2 (1981) 137. D.J. Halls, J. Anal. At. Spectrom., 10 (1995) 169. J. Murphy, G. Schlemmer, I.L. Shuttler, P. Jones and S.J. Hill, J. Anal. At. Spectrom., 14 (1999) 1593. H. Matusiewicz and R.E. Sturgeon, Spectrochim. Acta, 51B (1996) 377. F. Laborda, M.V. Vicente, J.M. Mir and J.R. Castillo, Fresenius’ J. Anal. Chem., 357 (1997) 837. B. Mitrovic and R. Milacic, Sci. Total Environ., 258 (2000) 183. A. Montaser and D.W. Golightly (Eds.), Inductively Coupled Plasmas in Analytical Atomic Spectrometry. VCH, Weinheim, Germany, 1987. In: P.W.J.M. Boumans (Ed.), Inductively Coupled Plasma Emission Spectroscopy, Part II: Applications and Fundamentals. Wiley, New York, 1987. R.N. Savage and G.M. Hieftje, Anal. Chim. Acta, 123 (1981) 319. A.D. Weiss, R.N. Savage and G.M. Hieftje, Anal. Chim. Acta, 124 (1981) 245. M. Chausseau, E. Poussel and J.M. Mermet, Fresenius’ J. Anal. Chem., 370 (2001) 341. I.B. Brenner and A.T. Zander, Spectrochim. Acta, Part B, 55 (2000) 1195. R.S. Pomeroy, CTD Detectors in Atomic Emission Spectrometry. VCH, New York, 1994. R.B. Bilhorn, P.M. Epperson, J.V. Sweedler and M.B. Denton, Appl. Spectrosc., 41 (1987) 1125. R. Thomas, Spectroscopy, 16 (2001) 38. B.L. Sharp, J. Anal. At. Spectrom., 3 (1988) 613. B.L. Sharp, J. Anal. At. Spectrom., 3 (1988) 939. A. Boorn and R.F. Browner, Anal. Chem., 54 (1982) 1402. A. Canals, L. Gras and H. Contreras, J. Anal. At. Spectrom., 17 (2002) 219. W.R.L. Cairns, L. Ebdon and S.J. Hill, Fresenius’ J. Anal. Chem., 355 (1996) 202. R. Gotz, J.W. Elgersma, J.C. Kraak and H. Poppe, Spectrochim. Acta, Part B, 49 (1994) 761. T.S. Conver, J.F. Yang and J.A. Koropchak, Spectrochim. Acta, Part B, 52 (1997) 1087. L. Ebdon and A. Fisher, The use of ICP-AES as a detector for elemental speciation studies. In: D. Barcelo (Ed.), Chapter 8 in Elemental Speciation. New Approaches

187

L. Ebdon et al.

59 60 61 62 63 64 65 66 67 68

69 70

71 72 73 74 75 76 77 78 79 80 81 82

83 84 85 86 87 88 89 90

188

for Trace Element Analysis, By J.A. Caruso, K.L. Sutton and K.L. Ackley. Vol. XXXIII of Wilson and Wilson’s Comprehensive Analytical Chemistry, Elsevier, Amsterdam, 2000. S.D. Overduin and I.D. Brindle, J. Anal. At. Spectrom., 16 (2001) 289. R.T. Gettar, R.N. Garavaglia, E.A. Gautier and D.A. Batistoni, J. Chromatogr. A, 884 (2000) 211. J.M. Carey and J.A. Caruso, Crit. Rev. Anal. Chem., 23 (1992) 397. T. Kantor, Spectrochim. Acta, Part B, 56 (2001) 1523. V. Karanassios and G. Horlick, Spectrochim. Acta, Rev., 13 (1990) 89. R. Sing, Spectrochim. Acta, Part B, 54 (1999) 411. S.A. Darke and J.F. Tyson, J. Anal. At. Spectrom., 8 (1993) 145. Thermo Jarrell Ash literature for ICAP 61E instrument. R. Thomas, Spectroscopy, 16 (2001) 26. E.H. Evans, J.J. Giglio, T.M. Castillano and J.A. Caruso, Inductively Coupled and Microwave Plasma Sources for Mass Spectrometry. Royal Society of Chemistry, Cambridge, 1995. In: K.E. Jarvis, A.L. Gray and R.S. Houk (Eds.), Handbook of Inductively Coupled Plasma Mass Spectrometry. Blackie Academic, 1992. G. O’Connor and E.H. Evans, Fundamental aspects of ICP-MS. In: S.J. Hill (Ed.), Inductively Coupled Plasma Spectrometry and Its Applications. Shefﬁeld Academic Press, Shefﬁeld, 1999. R. Thomas, Spectroscopy, 16(10) (2001) 44. R. Thomas, Spectroscopy, 16(11) (2001) 22. R. Thomas, Spectroscopy, 17(1) (2002) 36. M. Guilhaus, Spectrochim. Acta, Part B, 55 (2000) 1511. S.J. Ray and G.M. Hieftje, J. Anal. At. Spectrom., 16 (2001) 1206. M. Barbaste, L. Halicz, A. Galy, B. Medina, H. Emteborg, F.C. Adams and R. Lobinski, Talanta, 54 (2001) 307. S.N. Willie and R.E. Sturgeon, Spectrochim. Acta, Part B, 56 (2001) 1707. R. Thomas, Spectroscopy, 17 (2002) 42. M.A. Dexter, P.K. Appelblad, C.P. Ingle, J.H. Batey, H.J. Reid and B.L. Sharp, J. Anal. At. Spectrom., 17 (2002) 183. P. Leonhard, R. Pepelnik, A. Prange, N. Yamada and T. Yamada, J. Anal. At. Spectrom., 17 (2002) 189. R. Thomas, Spectroscopy, 17 (2002) 34. F. Vanhaecke, L. Moens and P. Taylor, Use of ICP-MS for isotope ratio measurements. In: S.J. Hill (Ed.), Inductively Coupled Plasma Spectrometry and Its Applications. Shefﬁeld Academic Press, Shefﬁeld, 1999. S.J. Hill, L.J. Pitts and A.S. Fisher, TrAC Trend Anal. Chem., 19 (2000) 120. E.H. Evans and J.J. Giglio, J. Anal. At. Spectrom., 8 (1993) 1. N. Nonose and M. Kubota, J. Anal. At. Spectrom., 16 (2001) 560. L. Ebdon, A.S. Fisher, P.J. Worsfold, H. Crews and M. Baxter, J. Anal. At. Spectrom., 8 (1993) 691. S. Branch, L. Ebdon, M. Ford, M. Foulkes and P. O’Neill, J. Anal. At. Spectrom., 6 (1991) 151. S.J. Hill, M.J. Ford and L. Ebdon, J. Anal. At. Spectrom., 7 (1992) 1157. L. Ebdon, M.J. Ford, R.C. Hutton and S.J. Hill, Appl. Spectrosc., 48 (1994) 507. A.W. Kim, M.E. Foulkes, L. Ebdon, S.J. Hill, R.L. Patience, A.G. Barwise and S.J. Rowland, J. Anal. At. Spectrom., 7 (1992) 1147.

Detection methods for the quantitation of trace elements 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109

110 111

112

113

114 115 116 117 118 119 120 121

K.A. Taylor, B.L. Sharp, D.J. Lewis and H.M. Crews, J. Anal. At. Spectrom., 13 (1998) 1095. P. Stchur, K.X. Yang, X.D. Hou, T. Sun and R.G. Michel, Spectrochim. Acta, Part B, 56 (2001) 1565. G.F. Kirkbright and M. Sargent, Atomic Absorption and Fluorescence Spectroscopy. Academic Press, London, 1974. A. Young, L. Pitts, S. Greenﬁeld and M. Foulkes, J. Anal. At. Spectrom., 18 (2003) 44. S. Greenﬁeld, J. Anal. At. Spectrom., 13 (1994) 565. J.L. Gomez-Ariza, M.A.C. de la Torre, I. Giraldez, D. Sanchez-Rodas, A. Velasco and E. Morales, Appl. Organomet. Chem., 16 (2002) 265. E. Puskel, Z. Mester and P. Fodor, J. Anal. At. Spectrom., 14 (1999) 973. P. Read, H. Beere, L. Ebdon, M. Leizers, M. Hetheridge and S. Rowland, Org. Geochem., 26 (1997) 11. B. Rosenkranz and J. Bettmer, TrAC Trends Anal. Chem., 19 (2000) 138. K. Malekani and M.S. Cresser, Commun. Soil Sci. Plant Anal., 29 (1998) 285. J.J. Thomson, Phil. Mag., 20 (1910) 752. R.F.G. Herzog, Patent DRP H172192 IXa/42h, 1942. R.F.G. Herzog and F.P. Viehbo¨ck, Phys. Rev., 76 (1949) 855L. R.E. Honig, J. Appl. Phys., 29 (1958) 549. R.E. Honig, Int. J. Mass Spectrom. Ion Processes, 66 (1985) 31. H.J. Liebl and R.F.G. Herzog, J. Appl. Phys., 34 (1963) 2893. G. Slodzian, Thesis Univ. Paris, 1963; R. Castaing, B. Jouffrey and G. Slodzian, C. R. Acad. Sci., 251 (1960) 1010. H.J. Liebl, J. Appl. Phys., 38 (1967) 5277. J.R. Rouberol, J. Guernet, P. Deschamps, J.-P. Dagnot and J.-M. Guyon de la Berge, Proc. 5th Int. Conf. X-Ray Opt. Microanal., Springer-Verlag, Heidelberg, 1969, pp. 311– 318. A. Benninghoven, Z. Phys., 230 (1970) 403. G. Tamborini, The development of the SIMS technique for the analysis of radionuclides in microparticles from environmental materials, PhD Thesis (in French), University of Paris-Sud, Orsay, France, 1998. G. Tamborini, Latest developments in particle analysis, High Performance Trace Analysis — Environmental Sampling (HPTA-ES), Nuclear Inspectors Training Course, European Commission, Luxembourg 2001. G. Tamborini, Analysis of Uranium and actinides by SIMS, European Microbeam Analysis Society, 8th Workshop on Modern Developments in Microbeam Analysis, Chiclana de la Frontera, Spain, May 2003. W.W. Harrison and C.W. Magel, Anal. Chem., 46 (1974) 461. B.N. Colby and C.A. Evans, Anal. Chem., 46 (1974) 1236. W.A. Mattson, B.L. Bentz and W.W. Harrison, Anal. Chem., 48 (1976) 489. W.W. Harrison, C.M. Barshisk, J.A. Klingler, P.H. Ratliff and Y. Mei, Anal. Chem., 62 (1990) 943A. C.R. Schick, P.A. De Palma and R.K. Marcus, Anal. Chem., 68 (1996) 2113. W.W. Harrison and W. Hang, J. Anal. At. Spectrom., 11 (1996) 835. W. Harrison, Glow discharge mass spectrometry, In: F. Adams, R. Gijbels and R. van Grieken (Eds.). Inorganic Mass Spectrometry. Wiley, 1988. L. King and W.W. Harrison, Glow discharge mass spectrometry. In: R.K. Marcus (Ed.), Glow Discharge Spectroscopies. Plenum Press, New York, 1993.

189

L. Ebdon et al. 122

123

124 125 126 127 128

129 130 131 132 133

134 135 136 137 138 139 140 141 142 143 144 145 146 147 148 149

190

W.W. Harrison, C. Yang and C. Oxley, Mass spectrometry glow discharge. In: R.K. Marcus and J.A.C. Broekaert (Eds.), Glow Discharge Plasma in Analytical Spectroscopy. Wiley, 2003. L.O. Actis-Dato, Etude par spectrome´trie de masse a` de´charge luminescente de la diffusion d’impurete´s chimiques dans des couches d’oxyde de zirconium, PhD thesis, Feb 2000, University Paris-Sud, Orsay, France. R.W. Smithwick, J. Am. Soc. Mass Spectrom., 3 (1992) 79. C.L. Yang, M. Mohill and W.W. Harrison, J. Anal. At. Spectrom., 15 (2000) 1255. L. Aldave de las Heras, E. Hrnecek, O. Bildstein and M. Betti, J. Anal. At. Spectrom., 11 (2002) 1011. M. Betti, J. Anal. At. Spectrom., 11 (1996) 855. A. Bogaerts, W. Schelles and R. van Grieken, Analysis of nonconducting materials by glow discharge spectrometry, In: R.K. Marcus and J.A.C. Broekaert (Eds.), Glow Discharge Plasma in Analytical Spectroscopy. Wiley, 2003. M. Betti, S. Giannarelli, T. Hiernaut, G. Rasmussen and L. Koch, Fresenius’ J. Anal. Chem., 355 (1996) 642. In: R.E. Van Grieken and A.A. Markowicz (Eds.), Handbook of X-ray Spectrometry. Marcel Dekker, New York, 1993. G. Lachance and F. Claisse, Quantitative X-ray Fluorescence Analysis: Theory and Applications. Wiley, Chichester, 1995. E.P. Bertin, Introduction to X-ray Spectrometric Analysis. Plenum Press, New York, 1978. P.J.M. Van Espen and K.H.A. Janssens, in: R.E. Van Grieken and A.A. Markowicz (Eds.), Handbook of X-ray Spectrometry. Marcel Dekker, New York, 1993, pp. 181– 294. W.K. De Jongh, X-Ray Spectrom., 2 (1973) 151. J.W. Criss and L.S. Birks, Anal. Chem., 40 (1968) 1080. G. Tolg and R. Klockenkamper, Spectrochim. Acta, Part B, 48 (1993) 111. F.W. Fiﬁeld and D. Kealey, Principles and Practice of Analytical Chemistry. 4th edn., Blackie, London, 1995. O. Hernandez, F. Jimenez, A.I. Jimenez and J.J. Arias, Analyst, 121 (1996) 169. J. Cowan, M.J. Shaw, E.P. Achterberg, P. Jones and P.N. Nesterenko, Analyst, 125 (2000) 2157. M.J. Shaw, P. Jones and P.N. Nesterenko, J. Chromatogr. A, 953 (2002) 141. B. Fairman, A. Sanz-Medel, P. Jones and E.H. Evans, Analyst, 123 (1998) 699. E.P. Achterberg, C.B. Braungardt, R.C. Sandford and P.J. Worsfold, Anal. Chim. Acta, 440 (2001) 27. A.R. Bowie, E.P. Achterberg, R.F.C. Mantoura and P.J. Worsfold, Anal. Chim. Acta, 361 (1998) 189. P. Jones and H.G. Beere, Anal. Proc., 32 (1995) 169. H.G. Beere and P. Jones, Anal. Chim. Acta, 293 (1994) 237. P. Jones, T. Williams and L. Ebdon, Anal. Chim. Acta, 217 (1989) 157. K.M. Ochsenkuhn, M. Ochsenkuhn-Petropolou, F. Tsopelas and L. Mendrinos, Mikrochim. Acta, 136 (2001) 129. G. Henze, W. Wagner and S. Sander, Fresenius, J. Anal. Chem., 358 (1997) 741. M. Vazquez, K. Mikhelson, S. Piepponen, J. Rama, M. Sillanpaa, A. Ivaska, A. Lewenstam and J. Bobacka, Electroanalysis, 13 (2001) 1119.

This page intentionally left blank

SECTION - 1

Chapter 6

Wet digestion methods Henryk Matusiewicz

6.1

INTRODUCTION AND BRIEF HISTORY

Sample (matrix) digestion plays a central role in almost all analytical processes, but it is not often recognized as an important step in analytical chemistry, with primary attention being directed to the determination step. This sense of priorities is reﬂected all too conspicuously in the equipment and investment planning of many analytical laboratories. However, a welcome trend in recent years points toward fuller recognition of the true importance of sample digestion (decomposition, dissolution) in the quest for high-quality analytical results and valid conclusions. Wet digestion with oxidizing acids is the most common sample preparation procedure. Many of the sample preparation methods currently in use were actually developed during the 19th century. In the early 1800s, Berzelius developed test tubes, separatory funnels, and platinum crucibles; in 1831, he ﬁrst made use of the conversion of SiO2 to SiF4 by reaction with HF for analytical purposes. In 1834, Henry and Zeise developed methods for the gravimetric determination of sulfur as sulfate in organic samples. Their method called for the sample to be digested with fuming nitric acid or aqua regia and fused with potassium hydroxide or potassium nitrate. The ﬁrst published wet digestion reagent was chloric acid from HCl þ KClO3, as described in 1838 by Duﬂos [1] as well as by Fresenius and Babo [2] in 1844. The classical wet digestion reagent HNO3 þ H2SO4 (the most important and most versatile of the so-called wet-oxidation mixtures) was investigated by Danger and Flandin in 1841 [3], for the destruction of organic matter. The use of pure concentrated HNO3 in a closed system under elevated temperatures and pressure is well known since 1860 from Carius [4]. Kjeldahl [5] digested organic biological material in 1883 with boiling concentrated H2SO4 in an open system. Hydrogen peroxide was introduced by Classen and Bauer [6] in 1884, and HClO4 was used at elevated temperatures by Stcherbak [7] in 1893. Relatively new is Van Slyke’s [8] mixture of H2SO4 þ H3PO4 þ KIO3 þ K2Cr2Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

193

H. Matusiewicz

O7 (1954). In 1955, Polley and Miller [9] introduced a mixture of 50% H2O2 þ conc. H2SO4 as a most powerful oxidizing reagent. Rediscovery of the high oxidizing power of OHz radicals (Fenton’s reagent H2O2/Fe2þ) [10] for biological materials in 1961 and 1968, by Sansoni et al. [11,12] led to a technique for wet digestion at temperatures below 1108C. Since the beginning of the 1970s, a large increase of general interest in different digestion techniques and in publications dedicated especially to wet digestion methods has been evident. This chapter gives an overview of wet digestion methods and recent developments and applications of the digestion of different materials. Other sample preparation methods, such as chemical extraction and leaching, solubilization with bases, enzymatic decomposition, thermal decomposition, and anodic oxidation, are beyond the scope of this contribution and will not be discussed here. 6.2

NOMENCLATURE

For some methods of analysis, it may be required that the analytical sample be in a liquid form—the sample solution. Thus, standard procedures to convert solid (or solid containing) samples to solutions prior to detection are required. However, this conventional designation is often imprecise or even misleading with respect to the actual mechanism of the process. Several very different names are sometimes applied to a single technique, which presents a considerable obstacle for anyone (particularly a non-specialist) interested in acquiring a quick overview of systems applicable to a speciﬁc task. The terms decomposition (of organic materials), dissolution (of inorganic materials), destruction, digestion, ashing, mineralization, acid-digestion, wet-ashing, and even oxidative acid digestion all refer to this process. In this chapter, the general expression will be digestion, which is speciﬁed as wet digestion; therefore, wet digestion will be the term used for obtaining the resulting acid solution. It should be mentioned that guidelines for terms used in sample digestion are provided by the International Union of Pure and Applied Chemistry (IUPAC) Analytical Chemistry Division [13,14]. 6.3

BIBLIOGRAPHY

There are numerous publications giving useful information on the digestion (dissolution and/or decomposition) of any conceivable combination of matrix and analyte. Some comprehensive books and general review articles (and references cited therein) contain material pertinent to either organic [15–18] or inorganic [19– 22] matrices; others, to both [23–29]. Within the scope of this chapter, a comprehensive discussion on digestion techniques is not feasible. For more comprehensive information, the following

194

Wet digestion methods

reviews and books are available: books by Sˇulcek and Povondra [20], by Bock [23] and by Krakovska´ and Kuss [29] are dedicated solely to digestion methods. Other books deal exclusively with a single technique: microwave-assisted sample preparation [30,31], this topic has also been reviewed elsewhere [32– 39]; even the literature on the use of microwave-assisted digestion procedures for subsequent sample analysis by means of electrothermal atomic absorption spectrometry (ET-AAS) is reviewed [40]. In 1997, the establishment of a site on the World Wide Web (WWW) for information transfer and education in the areas of sample preparation and microwave chemistry (http://www. sampleprep.duq.edu/sampleprep) was announced. Recommended guidelines for sample preparation (methods of digestion) of different matrices are also available from the Encyclopedia of Analytical Chemistry [41]. Although it is very difﬁcult to refer to every paper published in this area, the enlisted bibliography of this chapter gives comprehensive coverage of advance of the topic made to date. To follow the latest development and new applications in this ﬁeld, the reader may consult the annual reviews in Analytical Chemistry and Journal of Analytical Atomic Spectrometry. Relevant material is to be found under the headings “Sample preparation”, “Sample digestion”, and “Sample dissolution”, for the appropriate topics. Literature cited herein is not intended to be comprehensive, but has been selected with a view to relevance, as a pertinent review or seminal topic paper, or for its potential application, novel developments, and progress in wet digestion techniques. 6.4

REAGENTS AND VESSEL MATERIALS FOR WET DIGESTION PROCEDURES

Sample wet digestion is a method of converting the components of a matrix into simple chemical forms. This digestion is produced by supplying energy, such as heat; by using a chemical reagent, such as an acid; or by a combination of the two methods. Where a reagent is used, its nature will depend on that of the matrix. The amount of reagent used is dictated by the sample size which, in turn, depends on the sensitivity of the method of determination. However, the process of putting a material into solution is often the most critical step of the analytical process, because there are many sources of potential errors, i.e., partial digestion of the analytes present, or some type of contamination from the vessels of chemical products used. It is beyond the scope of this contribution to discuss all possible systematic errors; therefore, further details on how to avoid systematic errors during sample digestion that cannot be referred to in detail here are discussed by Knapp and Schramel in Chapter 2 of this book. The majority of wet digestion methods (total decomposition and strong attack) involves the use of some combination of oxidizing acids (HNO3, hot conc. HClO4, hot conc. H2SO4) and non-oxidizing acids (HCl, HF, H3PO4, dilute

195

H. Matusiewicz

H2SO4, dilute HClO4) and hydrogen peroxide. All of these acids are corrosive in nature, especially when hot and concentrated, and should be handled with caution to avert injury and accidents. Concentrated acids with the requisite high degree of purity are available commercially, but they can be puriﬁed further by sub-boiling distillation [42]. Detailed discussion of the properties and applications of these reagents may be found elsewhere [20,22–25]. Wet digestion has the advantage of being effective on both inorganic and organic materials. It often destroys or removes the sample matrix, thus helping to reduce or eliminate some types of interference. The physical properties of the common mineral acids used in sample preparation are summarized in Table 6.1. Most wet digestion procedures are conducted under conditions that, in terms of temperature or reagents used, must be considered as extreme. Thus, the material of which the ﬂasks, crucibles, etc., are made must be chosen carefully according to the particular procedure to be employed. The material from which the digestion device is fabricated is also a frequent source of elevated blanks. Elements can be either dissolved from the material or they can be desorbed from the surface. Very important in this respect is the nature of the material. The suitability of materials may be estimated according to the following criteria: heat resistance and conductance, mechanical strength, resistance to acids and alkalis, surface properties, reactivity, and contamination, whereby the speciﬁc characteristics of the organic and inorganic material must also be given special consideration. Table 6.2 shows preferred materials for digestion vessels. The apparatus and containers that are used for the wet digestion procedures must be scrupulously cleaned and tested for any possible contamination. Usually it is sufﬁcient to boil the ﬂasks in concentrated nitric acid followed by rinsing several times with ultrapure water before use. In cases where this procedure is not adequate, one of the most powerful cleaning procedures is steaming the vessels with nitric or hydrochloric acid with assembly in a microwave-heated sealed Teﬂon vessel [43]. This procedure is particularly recommended for quartz, borosilicate glass, and polytetraﬂuorethylene (PTFE) vessels. To generalize this section, nitric acid is an almost universal digestion reagent and the most widely used primary oxidant for the decomposition of organic matter, because it does not interfere with most determinations and it is available commercially in sufﬁcient purity. Hydrogen peroxide and hydrochloric acid can usefully be employed in conjunction with nitric acid as a means of improving the quality of a digestion. Hydrochloric acid and sulfuric acid may interfere with the determination of stable compounds. Mixtures with hydrochloric acid are generally used for samples containing principally inorganic matrices, and combinations with hydroﬂuoric acid are used to decompose silicate insoluble in the other acids. Safety considerations are particularly important when using perchloric acid.

196

TABLE 6.1 Physical properties of common mineral acids and oxidizing agents used for wet digestion

Nitric acid Hydrochloric acid Hydroﬂuoric acid Perchloric acid Sulfuric acid Phosphoric acid Hydrogen peroxide

Formula

HNO3 HCl HF HClO4 H2SO4 H3PO4 H2O2

Molecular weight

63.01 36.46 20.01 100.46 98.08 98.00 34.01

Concentration w/w (%)

Molarity

68 36 48 70 98 85 30

16 12 29 12 18 15 10

Density (kg/l)

Boiling point (8C)

Comments

1.42 1.19 1.16 1.67 1.84 1.71 1.12

122 110 112 203 338 213 106

68% HNO3, azeotrope 20.4% HCl, azeotrope 38.3% HF, azeotrope 72.4% HClO4, azeotrope 98.3% H2SO4 Decomposes to HPO3

Wet digestion methods

Compound

197

198

TABLE 6.2 Preferred materials for wet digestion vessels Chemical name

Working temperature (8C)

Borosilicate glass

SiO2a

,800b

Fused quartz

SiO2c

,1200

Glassy carbon

Graphite

,500

PE PP PTFE

Polyethylene Polypropylene Polytetraﬂuorethylene

,60 ,130 ,250

107 150

,0.02 ,0.03

PFA FEP TFM

Perﬂuoralkoxy Tetraﬂuorperethylene Tetraﬂuormetoxil

,240 ,200

166 158

,0.03 ,0.01 ,0.01

a

SiO2 content between 81 and 96%. Softens at a temperature of 8008C. c SiO2 99.8%. b

Heat deﬂection temperature (8C)

Water absorption (%)

Comments Ordinary laboratory glass is not suitable for use in wet digestion procedures For all procedures involving wet digestion of organic material, quartz is the most suitable material for vessels Glassy carbon is used in the form of crucibles and dishes for alkaline melts and as receptacles for wet digestion procedures

PTFE is generally used only for digestion vessels in pressure digestion systems

H. Matusiewicz

Material

Wet digestion methods

6.5

WET ACID DIGESTION (DECOMPOSITION AND DISSOLUTION) PROCEDURES

The task of preparing samples with acid treatment to release the elements of interest from the sample matrix and transfer them to a liquid matrix for subsequent analysis is commonplace in many laboratories. A variety of techniques are employed—from ambient-pressure wet digestion in a beaker on a hot plate (or hot block) to specialized high-pressure microwave heating. Dissolution is usually deﬁned as the simple process of dissolving a substance in a suitable liquid at relatively low temperature, with or without a chemical reaction. The term decomposition denotes a more complex process that is usually performed at higher temperature and/or at increased pressure, with the aid of reagents and special apparatus. A clear distinction between these terms cannot, however, be made. Table 6.3 gives an overview of the wet digestion methods, one of the oldest and still most frequently used techniques, for organic and inorganic samples. The intent is not to present the procedural details for the various sample matrices, but rather to highlight those methods that are unique to each technique and sample. 6.5.1

Open systems

Open vessel acid digestions, one of the oldest techniques, are undoubtedly the most common method of sample decomposition or dissolution of organic and TABLE 6.3 Schemes for wet digestion methods Digestion technique

Required reagents

Application

Open systems Conventional heating Microwave heating Ultraviolet digestion

HNO3, HCl, HF, H2SO4, HClO4, H2O2 HNO3, HCl, HF, H2SO4, HClO4, H2O2 H2O2, K2S2O8

Inorganic/organic Inorganic/organic Waters, slurries

Closed systems Conventional heating Microwave heating

HNO3, HCl, HF, H2O2 HNO3, HCl, HF, H2O2

Inorganic/organic Inorganic/organic

Flow systems Conventional heating UV on-line decomposition Microwave heating

HNO3, H2SO4, H2O2 H2O2, K2S2O8 HNO3, H2SO4, H2O2

Inorganic/organic Waters, slurries ? Inorganic/organic

Vapor-phase acid digestion

HNO3, HCl, HF, H2O2

Inorganic/organic

199

H. Matusiewicz

inorganic sample materials used in chemical laboratories. This very inexpensive technique is of inestimable value for routine analysis because it can easily be automated; all the relevant parameters (time, temperature, introduction of digestion reagents) lend themselves to straightforward control. Thousands of different methods and minor variations on these methods have been described in the literature. The main advantage of wet digestion (ashing) over dry ashing is its speed. However, systems of this type are limited by a low maximum digestion temperature, which cannot exceed the ambient-pressure boiling point of the corresponding acid or acid mixture. For instance, the oxidizing power of nitric acid with respect to many matrices is insufﬁcient at such low temperatures (boiling point 1228C). One possible remedy is the addition of sulfuric acid, which signiﬁcantly increases the temperature of a digestion solution. Whether or not this expedient is practical depends on the matrix and the determination method. High-fat and high-protein samples are generally not subject to complete digestion at atmospheric pressure. Other disadvantages relate to the risk of contamination through laboratory air, the necessarily rather large amounts of required reagents (very often employing expensive reagents), and the danger of losses of trace elements. Losses can be kept low by using an excess of acid (mainly nitric) combined with a reﬂux condenser and by optimization of temperature and duration. Nevertheless, systems operated at atmospheric pressure are preferred from the standpoint of workplace safety. 6.5.1.1 Conventional heating (thermally convective wet digestion) The conventional approach to wet digestion, which has proven its worth over many years, entails a system equipped with heated conventional source (Bunsen burner, heating plate, sand bath, etc.) operating either at a ﬁxed temperature or in response to a temperature program. Acid digestions are often accomplished in any vessel, usually in glass or PTFE (beaker, conical ﬂask, etc.) with or without a reﬂuxing condenser. However, when a sample is decomposed by open wet digestion, reﬂuxing is compulsory. The necessary apparatus has been described by Bethge [44]. Open block digestion systems have been popular in sample analysis over the past decades, but have consistently suffered from the major drawback of their sensitivity against corrosion and subsequent risk of contamination. Therefore, block digestion systems (hotplate techniques) have not been considered state-of-the-art technology in trace and ultratrace sample preparation. Graphite block digestion systems are becoming more frequently considered. These systems overcome the deﬁciencies of the traditional systems, made from stainless steel or aluminum, because the block is manufactured from graphite and typically coated with a ﬂuoro-polymer to prevent the possibility of metallic contamination from the surface of the system during the handling of the samples. Graphite block systems present an alternative to the current mainstream technology of open- and closed-vessel (“classical” or microwave-assisted) digestion systems, as they allow large

200

Wet digestion methods

numbers of samples to be digested simultaneously, thus overcoming one of the major weaknesses of closed-vessel systems. Commonly employed digestion agents include nitric acid, sulfuric acid, hydroﬂuoric acid, perchloric acid, and hydrogen peroxide, as well as various combinations of these. Most applications of wet digestion involve aqueous or organic matrices, such as surface waters, waste water, biological and clinical samples, food samples, as well as soil, sediment and sewage sludge, coal, high-purity materials, and various technical materials. More recently, open systems have progressed: the usual digestion ramps consist of several vessels equipped with reﬂux condensers to limit possible volatilization losses of some analytes and to avoid the evaporation of the reactive mixture. Such assembling is entirely satisfactory for ensuring concurrent digestions of large series of samples. Modern commercially available Hach Digesdahl Digestion Apparatus (Hach Comp., USA) is designed to digest organic and mineral samples for subsequent analysis. 6.5.1.2 Microwave heating (microwave-assisted wet digestion) The most innovative source of energy for wet digestion procedures is microwaves. Because the heating takes place inside the digestion mixture, microwave digestion is more efﬁcient than with conventional means of heating. Using microwaves, both the speed and the efﬁciency of digestion for some types of samples considered difﬁcult to solubilize are often improved. Additionally, automation becomes possible with some instrumentation. This technique is discussed in detail in Chapter 8 and will therefore only be brieﬂy summarized here. Several different names are applied to this technique. An example of incorrect terminology involves uncritical use of the expression “microwave digestion” for acid digestion with microwave excitation. Although this technique makes use of microwave radiation, the direct effects of this radiation are of minor importance, at most. Microwaves cannot rupture molecular bonds directly because the corresponding energy is too low to excite electronic or vibrational states. Rotational excitation of dipoles and molecular motion associated with the migration of ions are the only processes that are observed in this microwave ﬁeld [45]. For this reason, an expression such as “microwave-assisted digestion” is preferable and recommended. Since Abu-Samra et al. [46] reported on the application of microwave techniques to wet digestion of biological samples in 1975 (the ﬁrst paper published on microwave-assisted digestion), there has been a rapid development in microwave-assisted digestion for elemental analysis. Recent reviews [26–41] detail the application of microwave-assisted digestion to a wide variety of sample types, such as geological, biological, clinical, botanical, food, environmental, sludge, coal and ash, metallic, and synthetic materials and mixed samples and present speciﬁc experimental conditions as a function of the matrix to be digested. The earliest attempts at microwave-assisted digestion were performed using home appliance microwave ovens. This was necessary because commercial devices were not available at the time. The use of domestic

201

H. Matusiewicz

microwave ovens in laboratory experiments should be discouraged because of safety and performance. Microwave-assisted digestion in open systems at atmospheric pressure (focused microwaves using open vessels ﬁtted with reﬂuxing facilities made of borosilicate glass, quartz, or PTFE) is generally applicable only with simple matrices or for strictly deﬁned objectives, and the results are reproducible only if the speciﬁed digestion parameters are strictly observed. The performance of the focused-microwave-assisted systems and a wealth of applications have been reviewed by White and Mermet [30,31] and very recently by No´brega et al. [47]. Focused-microwave-assisted sample preparation is a suitable strategy for dealing with high masses of organic samples (up to 10 g). Losses may be encountered with mercury and possibly also with organometallic compounds (e.g. those containing arsenic, antimony, or tin). Addition of sulfuric acid is essential in order to achieve a sufﬁciently high digestion temperature using atmospheric pressure equipment, where the boiling point of the acid establishes the maximum digestion temperature, although it is important to remember that the presence of sulfate interferes with many procedures for metal determination (e.g. graphite furnace atomic absorption spectrometry or electrochemical techniques). New equipment, the non-pressurized digestion systems (STARPlus Systems, CEM Corp., USA; QLAB6000 System, Questron Technologies Corporation, Canada) are at the frontier of this ﬁeld. They are designed for routine use and can easily be automated. In addition, automated evaporation-to-dryness can be accomplished in an open vessel by attaching a pump to evacuate the fumes while the container is heated and multiple methods for different samples can be simultaneously applied owing to the possibility of operating each reaction vessel independently. All relevant parameters, such as reagent volume, digestion time, applied power, temperature, and addition of reagent composition, lend themselves to straightforward control. Although non-pressurized microwave systems are limited by a low maximum digestion temperature, which cannot exceed the ambient-pressure boiling point of the acid (or the acid mixture), they provide the best option with regard to the safety of personnel, because no overpressure can occur. Moreover, non-pressurized microwaveassisted digestion is suitable for on-line digestions in continuous-ﬂow systems (cf. Section 6.5.3.2). A compact apparatus in which a speciﬁc position can be irradiated by microwaves (MW) and ultrasound (US) simultaneously has been developed [48]. The combination of these two types of irradiation, electromagnetic (2.45 GHz) and mechanical (20 KHz), and their application to physical processes like digestion appears interesting. The MW–US reactor has been designed for atmospheric pressure decomposition and dissolution of biological (olive oil) and chemical products (refractory mineral material Co3O4) in nitric acid and hydrogen peroxide. Simultaneous MW and US irradiation is shown as a new technique for atmospheric pressure digestion of solid and liquid samples suitable for chemical and biological analysis.

202

Wet digestion methods

6.5.1.3 Ultraviolet digestion (photolysis) Ultraviolet (UV) digestion is utilized mainly in conjunction with uncontaminated or slightly contaminated natural water matrices (aqueous solutions), such as sea, surface, fresh, river, lake, ground, estuarine and coastal water. Liquids or slurries of solids are decomposed by UV radiation (light) in the presence of small amounts of hydrogen peroxide, acids (mainly HNO3) or peroxodisulphate (i.e. beverages, special industrial waste water, water of sewage treatment plants, soil extracts) [49]. Dissolved organic matter (DOM) and complexes of the analyte elements are decomposed to yield free metal ions. The corresponding digestion vessel should be placed in the closest possible proximity to the UV lamp (low- or high-pressure) to ensure a high photon ﬂux. In photolysis, the digestion mechanism can be characterized by the formation of OHz radicals from both water and hydrogen peroxide that is initialized by the aid of the UV radiation [49]. These reactive radicals are able to oxidize, to carbon dioxide and water, the organic matter present in simple matrices containing up to about 100 mg/l of carbon. Complete elimination of the matrix is, of course, possible only with simple matrices or by combining photolysis with other digestion techniques [50]. The method does not oxidize all organic components possibly present in water; chlorinated phenols, nitrophenols, hexachlorobenzene and similar compounds are only partly oxidized. Effective cooling of the sample is essential, because losses might otherwise be incurred with highly volatile elements. Hydrogen peroxide addition may need to be repeated several times to produce a clear sample solution. Modern UV digestion systems are commercially available (see Ref. [49], Table 1). 6.5.2

Closed systems

During the last few decades, methods of wet sample preparation using closed vessels have become widely applied. Closed systems offer the advantage that the operation is essentially isolated from the laboratory atmosphere, thereby minimizing contamination. Digestion of the sample is essentially ensured by a common wet digestion procedure, which is performed under the synergistic effects of elevated temperature and pressure; digestion occurs at relatively high temperature due to boiling-point elevation. The pressure itself is, in fact, nothing more than an undesirable—but unavoidable—side effect. These techniques are generally much more efﬁcient than conventional wet digestion in open systems, the loss of volatile elements is avoided, any contribution to blank values may be reduced and the digestion of more difﬁcult samples is possible. The principal argument in favor of this form of digestion is the vast amount of relevant experience acquired in recent decades. The literature is a treasure trove of practical information with respect to virtually every important matrix and a great number of elements. Closed system digestion is particularly suitable for trace and ultratrace analysis, especially when the supply of sample is limited.

203

H. Matusiewicz

Because the oxidizing power of a digestion reagent shows a marked dependence on temperature, an arbitrary distinction should be made between low (simple) pressure digestion and high-pressure digestion. Low-pressure digestions (,20 bar) are limited to a temperature of ca. 1808C, whereas with high-pressure apparatus (. 70 bar) the digestion temperature may exceed 3008C. 6.5.2.1 Conventional heating (thermally convective wet pressure digestion) The expression “pressure digestion” is another—very timely—example of incorrect terminology, giving the impression that pressure is critical for the digestion process. Indeed, it is a relatively high boiling temperature that ensures more effective digestion, not the associated high pressure. However, it is important to note that the pressure buildup, amongst other things, does introduce some dangers to the application of these methods. These methods are better described if they are grouped under the title, “wet digestion methods in closed systems”. However, the conventional designation should, in the author’s opinion, nevertheless be retained in this case, if only because of its wide acceptance; attempting to rename the procedure now would introduce more confusion than clarity. The digestion of inorganic and organic substances in sealed tubes was the method ﬁrst proposed for pressure digestion at the end of 19th century, and some of these applications are still difﬁcult to replace by other digestion methods. The use of sealed glass tubes goes back to Mitscherlich [51] and Carius [4,52], often referred to as the Carius’ technique, ﬁrst described in 1860. Carius undertook digestion of organic materials with concentrated nitric acid at 250– 3008C. The sample and the acid were mixed in a strong (thick)-walled quartz ampule and sealed. The ampule was transferred to a “bomb canister” and heated in what was called a “bomb oven” for several hours, after which it was cooled, opened, and the contents analyzed. Carius tube digestion involves the generation of internal pressure in excess of 100 bar (atm) at 2408C. The modern redesign and employment of the Carius combustion tube for the digestion of some refractory materials was accomplished at the US National Bureau of Standards during the 1940s [53–55]. Extreme care should be used in the handling and venting of pressurized tubes. A discussion of Carius tube design for minimizing losses and hazards by explosion has been provided by Gordon [54]. For safety, any stainless steel sleeve jacket (along with solid CO2 pellets, to maintain equal pressure across the tube wall when heated) that is large enough to contain the Carius tube will sufﬁce as an external pressure vessel [56]. With the development of the so-called Carius tube, the ﬁeld of closed-vessel digestion was born. Digestion in autoclaves with metal inner reaction vessels was originally proposed in 1894 by Jannasch [57], but was not widely employed because of a number of drawbacks (such as strong corrosion of the platinum vessel). Seventy years later, May and Rowe [58] designed a new type of

204

Wet digestion methods

autoclave with an inner lining made of Pt–Ir alloy (platinum-lined crucible and bomb), which is more resistant than platinum alone, for digestion with hydroﬂuoric acid. It is difﬁcult to construct an autoclave with an inner metal reaction vessel and it is thus very expensive. Therefore, this technique has not been used extensively in analytical practice and thus the apparatus is not manufactured commercially. Extensive use of pressure digestion in analytical procedures began in 1960, as a result of the considerable technological progress in the manufacture of organic polymers. Convectively heated pressure vessel systems have proved to be the most valuable systems for guaranteeing complete, or almost complete, digestion of solid samples because they provide elevated digestion temperatures (about 200 –2308C [59]). Most sample vessels (containers) for use in thermally convective pressure digestion are constructed from PTFE [60 –62], PFA [63], or PVDF [64], although special quartz vessels with PTFE holders [65] or glassy carbon vessels [66] are available for trace analysis purposes. The sample vessel is mounted in a stainless steel pressure autoclave and then heated, usually in a laboratory drying oven, furnace or heating block, to the desired temperature. Because of the necessity to examine numerous samples, mechanized multisample pressure digestion systems able to process rather large sample numbers of the same matrix type were developed [67]. A cooling circuit can be ﬁtted into the metal casing (jacket) to permit rapid manipulation of the solution formed immediately after removing the “digestion bomb” from the oven or heating block [68]. Dissolution can be also accelerated by mixing the reactants, preferably by using a stirring bar (covered with PTFE) [69]. An alternative design has been proposed by Uchida et al. [70], wherein a small screw cap vial for sample digestion is placed inside the Teﬂon digestion vessel. This digestion system (Teﬂon double vessel) can reduce the risk of sample leakage and contamination with extraneous materials. To improve dealing with pressure –temperature evaluation and the carbon balance for some materials, a system with a Teﬂon-lined membrane pressure meter and a thermocouple was designed [71]. Recently, a digestion vessel for use with a convection oven was proposed [72], which has an unusual design in which the vessel consists of three nested structures: an innermost PTFE container of 30 ml capacity, an intermediate PTFE container of 100 ml capacity, and an outer stainless steel shell. It should be stressed here that digestion in a Teﬂon-lined pressure vessel using one or a mixture of acids does not result in complete decomposition (see Section 6.5.5), because of the limited temperature. Pressure digestion systems are all feasible below ca. 2008C, but above this temperature PTFE begins to “ﬂow”, rendering it unsuitable for use in high-pressure applications, and consequently at higher temperatures. All thermally initiated digestions have the disadvantage that a considerable amount of time is consumed in preheating and cooling the digestion solutions and sample vessel [73], the limited sample size, and the inability to visually check the progress of the digestion.

205

H. Matusiewicz

The contributions of Langmyhr, Bernas, To¨lg, and co-workers are worth mentioning with regard to the commercialization of the digestion vessels or “digestion bombs”, as they are often called. Today, there are a number of digestion bombs covering the whole market range, including: the popular Parr acid digestion bombs (Parr Instrument Company, USA), Uniseal decomposition vessels (Uniseal Decomposition Vessels Ltd, Israel), stainless steel pressure vessels with Teﬂon inserts (Berghof Laborprodukte GmbH, Germany), the pressure decomposition system CAL 130FEP (CAL Laborgera¨te GmbH, Germany), and the pressure digestion system (PRAWOL, Germany). To avoid the problem of loss of mechanical stability at high temperatures, vessels made of quartz are now being used in a new pressure digestion system [74,75]. The introduction of a high-pressure ashing (HPA) technique by Knapp [76] has not only reduced the effective digestion time but also opened the way to digestion of extremely resistant materials, such as carbon, carbon ﬁbers, mineral oils, etc. In academic terms an HPA currently represents the highest standard in pressurized wet digestion techniques, combining the advantages of the Carius technique with easy and safe handling. Essentially complete digestion can be accomplished with the vast majority of samples so far investigated. Nitric acid alone is a sufﬁciently powerful reagent in many cases. High-pressure digestion is conducted in quartz vessels, with a maximum digestion temperature as high as 3208C at a pressure of ca. 130 bar. For dissolution requiring HF, glassy carbon vessels are used instead of quartz. The quartz vessel (or glassy carbon) is stabilized during the digestion process by subjecting it to an external pressure roughly equivalent to or higher than that developed internally. Because the pressure within the vessel is lower than the pressure applied, the vessel is protected from explosion. A perfected system of wet digestion under high temperature and pressure developed by Knapp is commercially available, the HPA-S High Pressure Asher system (Anton Paar GmbH, Austria). Very recently, again in respect to complete digestion of organic waste materials, a potent digestion technique was developed [77] based on a prototype of an HPA device using infrared heating (IR-HP-asher). High-pressure digestion is conducted in six quartz vessels inside a steel autoclave, with a maximum digestion temperature as high as 3008C at a pressure of 130 bar. The novelty of this approach lies in the design of an HPA system with IR heating. In comparison to open vessel digestion, closed-vessel digestion methods have many advantages, but there is one disadvantage—complex and expensive vessel designs. A new technique—pressurized wet digestion in open vessels— combines the advantages of closed-vessel sample digestion with the application of simple and inexpensive open vessels made of quartz or PFA [78]. The vessels are placed in a High Pressure Asher HPA, which is adapted with a Teﬂon liner and partially ﬁlled with water. The vessels (in principle any shape of vessel can be used) are loaded with sample material and digestion reagent and are simply covered with PTFE stoppers and not sealed. The vessels are transferred to

206

Wet digestion methods

a specially adapted HPA and digested at temperatures up to 2708C. The digestion time is ca. 90 min, and cooling to room temperature requires a further 30 min. As metal autoclaves are expensive, a pressure vessel without an outer metal casing has been designed. The technique of digestion at slightly increased pressure has been found to be very useful for routine laboratory analysis, primarily because of its simplicity. Almost an unlimited number of samples can be digested simultaneously. The vessel can be sufﬁciently well sealed by using a screw cap [79]. Volatile components are not lost during heating and the laboratory atmosphere is thus not contaminated by acid vapors. All-Teﬂon thick-walled PTFE vessels (bombs) have been used in the dissolution of refractory oceanic suspended matter using HCl, HNO3, and HF [80]. Translucent Nalgene-sealed bottles have been proposed for the “wet pressure digestion” of biological materials (ﬁsh, bird, plant tissue) using a combination of HClO4 and HNO3 [81]. A method utilizing a pressure digestion technique for real sample matrices using linear polyethylene bottles has been proposed [82]. Vessels of polyethylene are transparent, permitting observation of the whole digestion process and reduction of the reaction time to a minimum. A complete digestion of fatty material with slight overpressure (,4 bar) was possible in a closed system completely made from quartz [83]. A closed PTFE bomb (30 ml capacity, screw-cap vessel machined from molded, stress-relieved Teﬂon-TFE rod) was designed for the digestion of materials using a conventional heating (drying) oven [84]. 6.5.2.2 Microwave heating (microwave-assisted pressurized wet digestion) Closed-vessel microwave-assisted digestion technology has been acknowledged as one of the best solutions for clean chemistry applications and has a unique advantage over other closed-vessel technologies. The vessels used for microwave acid digestion are either low-pressure or high-pressure bombs. The current generation of microwavable-closed vessels consists of a two-piece design, liners and caps composed of high-purity Teﬂon or PFA with casings (outer jacket) made of polyetherimide and polyetherethereketone or other strong microwave transparent composite material. Their practical working temperature is 2608C (softening point of Teﬂon), and their pressure limit is 60–100 bar. Closed-vessel digestion is ideal for those samples that are being dissolved in HNO3 and/or HCl. However, for those digestions where H2SO4 is required, such as digestions of petroleum products, there is little advantage in using the regular closed-vessel approach because the boiling point of H2SO4 (3308C) exceeds the temperature available in a Teﬂon-lined vessel. The use of closed-vessel microwave-assisted digestion techniques minimizes the analytical blank by minimizing the amount of reagents used and by controlling the digestion environment, as well as through augmentation of the operator’s skills. Microwaves only heat the liquid phase, while vapors do not absorb microwave energy. The temperature of the vapor phase is therefore lower

207

H. Matusiewicz

than the temperature of the liquid phase and vapor condensation on cool vessel walls takes place. As a result, the actual vapor pressure is lower than the predicted vapor pressure. This sort of sustained dynamic, thermal nonequilibrium is a key advantage of microwave technology, as very high temperatures (and, in turn, short digestion times) can be reached at relatively low pressures. The inspiration for pressure digestion studies came from a US Bureau of Mines report [85], which described how rapid dissolution of some mineral samples had been achieved using a microwave oven to heat samples and an acid mixture contained in polycarbonate bottles. To overcome the presence of “hotspots” in the oven, which result in uneven heating, they designed a polypropylene rack to ﬁt on top of the standard microwave carousel. Although sealed polycarbonate bottles were used as pressure vessels, the plastic quickly became opaque and brittle (the melting point of polycarbonate is 1358C). Smith et al. [86] substituted Teﬂon PFA ﬂuorocarbon resin (a tetraﬂuoroethylene with a fully ﬂuorinated alkoxy side-chain) vessels for polycarbonate because of its superior chemical and mechanical properties. Buresch et al. [87] used low pressure-relief type containers made of PTFE or quartz. Alvarado et al. [88] exploited modiﬁed thick-walled Pyrex glass test tubes ﬁtted with polypropylene screw caps as pressurizable vessels. Kojima et al. [89] modiﬁed a Teﬂon digestion bomb by using a double Teﬂon vessel with a polypropylene jacket to permit leak-free and safe digestion of samples. A closed-vessel microwave digestion system was described [90]. In situ measurement of elevated temperatures and pressures in closed Teﬂon PFA vessels during acid decomposition of organic samples was demonstrated. Temperature and pressure monitoring permitted controlled digestions, studies of digestion mechanisms, and the development of transferable standard microwave sample preparation methods. Laboratory-made all Teﬂon bombs, used for low- or medium-pressure work, are also appropriate for microwave-heated digestion purposes [91], some ﬁtted with pressure-relief holes, valves, or membranes (rupture discs). Low-volume microwave-assisted digestion methods have found applications for studies involving small sample sizes where loss of sample in large digestion equipment is inevitable. Small quantities of tissue (5 –100 mg dry weight) are digested in high-purity nitric acid by use of a modiﬁed Parr microwave acid digestion bomb with modiﬁed Teﬂon liner [92]. The use of low-volume (7 ml) Teﬂon-PFA closed vessels designed for the preparation of small-sized (,100 mg dry mass) biological tissues has been described [93]. In order to prevent excessive pressure rises during closed microwave acid digestion of fairly large (1 g) samples having high organic content, an openvessel pre-digestion technique under reﬂux was designed to allow the escape of oxidation products, such as carbon dioxide, without incurring evaporation losses of acid or analytes. Following pre-digestion, the vessels were capped and subjected to microwaves to complete the digestion under pressure [94].

208

Wet digestion methods

In an attempt to minimize the delay in opening Teﬂon pressure vessels following microwave acid digestion, and thus signiﬁcantly reduce sample preparation time, digestions with the pressure vessels immersed in liquid nitrogen and the use of liquid nitrogen as a pre- and post digestion coolant were applied [95]. In other developments, a special type of Teﬂon bomb was constructed in which the vapor pressure can be maintained at a moderate level (up to 5 bar) by means of an internal quartz or Teﬂon cooling spiral (a water cooling spiral is inserted into a closed space through the cover). During operation, reﬂux of the condensed acid and water vapors continuously renews the liquid phase over the sample [96]. Several microwave-heating conﬁgurations were presented by Pougnet et al. [97,98] based on 500 or 1200 W, 2.45 GHz fundamental-mode microwave waveguide cavities, which heat pressure vessels currently used in laboratories for sample digestion and other applications. The capsule concept was reviewed in detail by Le´ge`re and Salin [99,100]. The sample is handled in an encapsulated form until it is in the digestion solvent. The operation of the capsule-based microwave-assisted digestion system proceeds in several steps, during which temperature and pressure are monitored. The heating in such a system, as in all microwave bomb systems, is from the solution outward, and the system performance is dictated by the same chemical and physical laws governing other microwave-assisted systems. From the previous discussion, it is clear that microwave acid digestion can be easily adapted for closed-vessel digestions; hence, its application has been limited to digestions in closed Teﬂon-lined vessels made of non-metallic microwave-transparent materials operating with a maximum upper safe pressure of around 60 –100 bar. In response to these limitations, and focusing on the fact that rapid heating of solvents and samples within a polymer vessel can lead to signiﬁcant advantages over high-pressure steel-jacketed Teﬂon bombs (which are thermally heated), Matusiewicz [101] developed a focusedmicrowave-heated bomb that would exceed the operational capabilities of existing microwave digestion systems and permit the construction of an integrated microwave source/bomb combination. The combined advantage of having a high-pressure Teﬂon bomb [59] incorporating microwave heating [31] has produced a focused high-pressure, high-temperature microwave-heated digestion system [101] capable of being water or ﬂuid cooled in situ. Another vessel conﬁguration integrates the microwave chamber around the vessel. These systems consist of one or several microwave transparent vessels (Teﬂon, quartz), which can either be opened [102] or sealed [103], enclosed in an acidresistant stainless steel chamber. The steel chamber acts as both the pressure vessel and microwave chamber. Modern systems can handle acid decompositions at temperatures up to 3208C and pressures of 130 –200 bar. Very recently, a novel microwave-assisted high-temperature UV digestion for accelerated decomposition of dissolved organic compounds or slurries was developed [104]. The technique is based on a closed, pressurized, microwave

209

H. Matusiewicz

digestion device wherein UV irradiation is generated by immersed electrodeless Cd discharge lamps (228 nm) operated by the microwave ﬁeld in the oven cavity. The immersion system enables maximum reaction temperatures up to 250– 2808C, resulting in a tremendous increase of mineralization efﬁciency. Today, there are a number of microwave-digestion bombs available: Parr bombs (Parr Instrument Company, USA), Berghof all-PTFE digestion vessels (Berghof GmbH, Germany), and TFM digestion vessels (Bohlender, Germany). 6.5.3

Flow systems

Discrete vessel systems, whether at elevated or atmospheric pressure, require a large amount of handling. Processes such as assembling, closing, opening, and positioning the vessel in the ordinary oven or microwave ﬁled are laborious and time consuming. Continuous ﬂow-through thermal digestion, UV digestion, and microwave digestion systems were designed to overcome some of the limitations by replacing the vessels with ﬂow-through tubing (coil). Samples are digested by pumping them through a coil containing a digestion matrix while being heated (by thermal, UV, or microwave). The continuous ﬂow of a carrier stream through these systems washes the system, removing the need for tedious vessel clean-up procedures. These systems can handle reactions that produce sudden increases in temperature and pressure, or unstable samples. Many different designs of ﬂow digestion systems have been published, but very few meet the prerequisites for high-performance sample decomposition. 6.5.3.1 Conventional heating (thermal) Many of the disadvantages of sample digestion can be overcome by automating sample preparation in an enclosed system through the use of ﬂow technology. A well-established digestion system, based on the ﬂow-stream principle, was developed by Technicon [105]. A slowly rotating, very wide glass tube with a spiral-type cavity is heated externally. Liquid sample, together with the decomposition reagent, is continuously pumped in one direction. By rotation of the tube, the decomposition mixture moves through the heated tube within a few minutes. At the other end of the tube, the ﬁnished sample is continuously pumped away and is ready for further analytical steps. A disadvantage of this system is that only liquid samples can be used, and the “memory effect” is large. The sample throughput is very high, however, for comparatively small apparatus cost. Another system was presented by Gluodenis and Tyson [106]. Here, the PTFE tubing is loosely embedded in a resistively heated oven. By using PTFE tubing, the maximum digestion temperature is restricted to ca. 2108C. The limited mechanical strength of the material merely allows maximum working pressures of up to 35 bar. Therefore, the usual working pressure is about 10–20 bar. The potential of the system was illustrated by the digestion of cocoa

210

Wet digestion methods

powder slurried in 10% HNO3 injected into the manifold and digested under stopped-ﬂow, medium-pressure conditions. In a series of papers [107 –109], Berndt described development of a new high-temperature/high-pressure ﬂow system for the continuous digestion of biological and environmental samples. It was shown [107] that temperatures up to 2608C and pressures up to 300 bar can be reached in a ﬂow system when an electrically heated Teﬂon lined HPLC tube is used as the digestion capillary. The required back-pressure was obtained by using a restrictor capillary with an inner diameter of 50 mm and a length of about 10 cm. Digested biological samples (blood, liver, leaves) were collected at the outlet of the ﬂow system. In subsequent papers [108,109], an electrically heated Pt/Ir capillary served as a digestion tube at temperatures of 320– 3608C and pressures of about 300 bar, and withstands concentrated acids. Due to the totally glass-free environment, samples having high silicate content can be digested by the addition of hydroﬂuoric acid. 6.5.3.2 UV on-line decomposition UV digestion is a clean sample preparation method, as it does not require the use of large amounts of oxidants. Furthermore, UV digestion is effective and can be readily incorporated into ﬂow injection manifolds. The sample ﬂows, in the presence of H2O2, H2S2O8 or HNO3, through a tube (PTFE, quartz) coiled around a ﬁxed UV lamp(s). A short review of such ﬂow systems has appeared recently [49]. Analyzers of this kind are produced by SKALAR Analytical, Holland, for example. Fernandes et al. [110] developed a manifold based on a two-stage on-line UV/thermal induced digestion procedure for oxidation purposes. The UV digestion apparatus consisted of a 4 m long PTFE tube tightly wounded directly around the UV source (15 W) to form a spiral reactor. The thermal digestion apparatus consisted of a 2 m long PTFE tube coiled in a helix and submerged in a thermostatic bath at 908C. Flow systems are becoming more popular in analysis, because of their ease of automation, speed, small volume of sample, and elegance, and thus provide for a promising future. 6.5.3.3 Microwave heating (microwave-assisted pressurized ﬂow-through digestion) Many different designs of microwave-assisted ﬂow digestion systems have been published [31,39,111], which open up new possibilities, primarily in fully automated sample preparation for elemental analysis. The earliest work reported in this ﬁeld was by Burguera et al. [112] who applied a ﬂow injection system for on-line decomposition of samples and determined metals (Cu, Fe, Zn) by ﬂame AAS. The methodology involved the synchronous merging of reagent and sample followed by decomposition of serum, blood, or plasma in a Pyrex coil located inside the microwave oven. This

211

H. Matusiewicz

approach permits essentially continuous sample digestion and drastically reduces sample processing time, and is suitable for those samples that require mild digestion conditions (especially liquids). According to the location of the digestion unit in the system, there are two types of manifolds described in the literature to date: before and after the injection unit. In the former arrangement, the sample is introduced into the microwave oven in a continuous ﬂow [113] or a stopped ﬂow mode [114]; after digestion, a discrete aliquot is delivered to the detector. In the second arrangement, the injected sample ﬂows to the microwave oven unit together with the reagent(s) to be digested, and is then cooled and degassed prior to its delivery to the detector [115]. In both cases, the measurements can be performed partially or totally off-line or on-line. Solid samples call for more sophisticated ﬂow systems because they need to digested in the presence of highly concentrated acids, which rapidly destroy organic matrices. A ﬁrst attempt aimed at simplifying manipulation of the digest was reported in 1988 [116]. Lyophilized, ﬁnely ground and weighed samples of liver and kidney were placed in test tubes together with mineral acids and the contents shaken before exposing them to microwave radiation to avoid violent reaction with abundant foam formation. The tubes were loaded into a covered Pyrex jar inside a domestic microwave oven operated for a speciﬁed time at a given power. Carbonell et al. [113] initiated the determination of metallic elements in solid samples using the slurry approach coupled with microwave oven digestion in a ﬂow injection system for F-AAS determination of lead. Various natural samples (artichoke, chocolate, sewage sludge, tomato leaves), real and certiﬁed, were slurried in a mixture of HNO3 and H2O2 using magnetic stirring, followed by continuous pumping around an open recirculating system, part of which (120 cm PTFE tubing) was located in a domestic microwave oven. A microwave-heated, ﬂow-through digestion container (coiled Teﬂon tubing) was designed for a commercial (Prolabo A300) focused microwave system (instead of microwave oven) and applied to the on-line preparation of biological samples, including milk, blood, and urine [117]. For an extensive oxidation of organic sample constituents with nitric acid, temperatures of more than 2008C are necessary. The PTFE tubes used, however, cannot withstand the vapor pressure of the digestion mixture at 2008C or more. Thus, new alternatives had to be found to overcome this limitation. One way to increase the pressure resistance of the tubes is to wrap them with a plastic tape of high mechanical strength. Results from a digestion system (CEM SpectroPrep system) equipped with such tubes have been published [118]. A CEM SpectroPrep system was used at moderate powers to perform on-line digestion of slurried samples of biological tissues (0.5% m/v) and marine sediment (1% m/v). The pressure thresholds of this system are near 25 bar. To achieve the desired temperatures of approximately 2508C, however, it is necessary to be able to increase the pressure in the system up to 35 bar or

212

Wet digestion methods

so. A recently developed device enables the application of such high temperatures (2508C) by means of a new pressure equilibrium system (with a pressure of 40 bar) [119]. The pressure equilibrium system keeps the pressure inside and outside the digestion tube (PTFE or PFA) equal, even for extremely fast oxidation reactions. Advantages of this high-performance ﬂow digestion device are high sample throughput (up to 60 samples per hour), fast and complete digestion of liquids, emulsions and slurries, and computer-controlled fully automated sample decomposition. The systems ability to handle only up to 1% m/v slurries and lower slurry concentrations for biological materials restricts the type of sample that can be analyzed, unless the most sensitive elemental detection devices are used, such as ICP-MS. Therefore, Mason et al. [120] modiﬁed the SpectroPrep oven and developed a wide bore continuous ﬂow microwave digestion system for the determination of trace metals (Cd, Cr, Mn, Ni, Pb) following aqua regia extraction. This device differs from existing commercially available devices as it uses a double pumping action to replace the back-pressure regulator traditionally used to achieve internal pressurization. The described system demonstrated an ability to cope with real soil samples ground to a larger particle size (250 mm) and slurried without the use of surfactants. Perhaps, the current fascination for using microwave heating for on-line digestion has led to the introduction of commercial instruments based on this hybrid technique. CEM developed the SpectroPrep continuous-ﬂow automated microwave-digestion system. Similarly, Questron Technologies Corporation is marketing the QLAB AUTOPREP DISCRETE FLOW SYSTEM. Perkin-Elmer offers an on-line ﬂow injection microwave-digestion system, as does SGT Middelburg BV (The Netherlands, FLOWAVE), a fully automatic continuous ﬂow sample preparation and digestion system based on microwave technology. The advantages of microwave-enhanced ﬂow systems basically include a signiﬁcant reduction in sample preparation time, the ability to accomplish reactions that would normally be too dangerous in a closed vessel because of sudden increases in temperature and pressure, and the capability to handle transient or readily decomposed samples or intermediates. However, ﬂowthrough systems are a problem area because all samples must be homogeneous and small enough to pass through the tube, and the majority of samples requires some form of processing before they can be put into the tube. 6.5.4

Vapor-phase acid digestion (gas-phase reactions)

An alternative approach to acid digestion of the sample matrix that prevents introduction of impurities exploits gas-phase reactions. In the past four decades, several novel approaches to sample digestion procedures have been considered using inorganic acid vapor produced in one vessel to attack and dissolve material in another. A review by Matusiewicz [121] summarized analytical methods based on vapor-phase attack in promoting the dissolution

213

H. Matusiewicz

and decomposition of inorganic and organic materials prior to determination of their trace element content. This approach is currently used (in open, semiclosed, and closed systems) whenever applicable because digestion using gasphase reagents is preferred to the solution. The combination of hydroﬂuoric acid and nitric acid vapor as a digestion agent has proven effective in the preparation of samples for spectrographic determination of trace impurities in open system. Zilbershtein et al. [122] used this approach to dissolve silicon and to concentrate impurities on a PTFE sheet. The residue and PTFE sheet were transferred to a graphite electrode that subsequently served as one electrode of the dc arc for spectrographic trace analysis. However, dissolution with acids in open systems is unsuitable for attaining a sufﬁciently low determination when the analyte impurities in the reaction mixture are often far greater than those in the test component. With respect to semi-closed systems, a PTFE apparatus generating HF vapor has been speciﬁcally designed to minimize contamination during traceelement determination of ultrapure silicon, quartz, and glass [123]. The sample is placed in a PTFE beaker mounted on a perforated PTFE plate that is kept above the level of liquid HF in the chamber. The apparatus has the advantage of a closed system by preventing air-borne particulates from entering the vessel but is continuously purged by a positive pressure of HF vapor during operation. Thomas and Smythe [124] describe a simple all-glass apparatus for vaporphase oxidation of up to 90% of plant material with nitric acid. Addition of perchloric acid ensured fast and complete oxidation, and the presence of HNO3 during the ﬁnal HClO4 oxidation step eliminated any danger of explosion. Klitenick et al. [125] used the same technique, with a simpliﬁed pressurized PTFE digestion vessel, for determination of zinc in brain tissue. Some materials may not be fully dissolved by acid digestion at atmospheric pressure. A more vigorous treatment involves bomb digestion in pressure vessels designed to incorporate the techniques of a closed pressure vessel and vapor-phase digestion in a single unit. This has the advantage of being easier to construct than the apparatus described in previous papers [122 –125], and it requires considerably smaller volumes of acids. Heating can be accomplished in an ordinary oven (with conductive heating) or using a microwave ﬁeld. A predecessor of this concept of closed-vessel vapor-phase sample digestion was introduced by Woolley [126]. He described a low-temperature (up to 1108C) and high-temperature (up to 2508C) version of the apparatus. Each device consists of an airtight PTFE vessel containing two concentric chambers: an inner chamber that holds the sample cup and an outer chamber. Both vessels were designed for the digestion of high-purity glass using relatively impure solvent acids: a 50:50 mixture of concentrated HNO3 and HF. A completely closed PTFE bomb or autoclave [127] has been developed with a temperature gradient for digestion of more difﬁcult compounds, such as siliceous material. Marinescu [128] presented an interesting development in which the conventional single-sample pressure digestion bomb was converted for multi-sample

214

Wet digestion methods

vapor-phase digestion. A multiplace holder for ﬁeld sampling was developed to ﬁt directly into the digestion bomb. This technique has been used for organic and inorganic solid, semisolid, and liquid samples. Kojima et al. [129] modiﬁed a sealed PTFE bomb in which the dissolution of highly pure silica with HNO3, HCl, and HF acid vapor was possible using a PTFE vial placed in a PTFE outer vessel. A possible disadvantage of this system is that the vial has to be replaced regularly when used under pressure. This could make the method very costly. A laboratory-made high-pressure digestion bomb with a PTFE microsampling device was developed by Matusiewicz [130]. This simple and inexpensive apparatus was found to be convenient for treating a small number of samples and can be easily made by modifying available PTFE bombs [131]. It should be noted that PTFE microsampling devices can be used for both vapor-phase digestion and discrete nebulization techniques in atomic spectrometry. Vaporphase digestion in a closed system (bomb) of high-purity materials for spectrographic determination of trace elements is a convenient and useful technique [132]. The method uses graphite electrodes with an enlarged cavity and excludes the use of a collector. Contact of the probes and the analytical impurities with laboratory glassware and the atmosphere during the preconcentration process is also avoided. The blank signal is determined only by the purity of the electrode used for spectral analysis. A technique [133] has been developed that employs the vapor-phase acid generated in the quartz vessel of a commercial high-pressure, high-temperature digestion apparatus (High Pressure Asher HPA, Anton Paar, Graz, Austria). Small biological samples (50–165 mg) were digested in a mini-quartz sample holder (3.1 ml volume). When biological standard reference materials were digested at 2308C and 122 bar, the residual carbon content (RCC) in the digested samples was less than 1.8%. Despite methodologies previously proposed for closed systems with conventional heating being successful, very few attempts to employ microwave power for vapor-phase digestions have been described. An early trial with a low-pressure microwave arrangement was unsatisfactory [134], although recently an interesting variant of the digestion vessel design has been proposed for dissolution and decomposition of samples [135]. The method developed was an extension of the acid vapor-phase thermal pressure decomposition of biological materials reported previously by Matusiewicz [136]. Microwaveassisted vapor-phase acid digestion employing a special PTFE microsampling cup, suitable for 250 mg subsamples of marine biological and sediment reference materials were digested with HNO3 and HNO3 –HF, respectively, at a maximum pressure of ca. 14 bar [135]. Very recently, several papers [137–141] discussed the further application and evaluation of this pioneering concept of Matusiewicz et al. [135], employing either commercial pressurized microwave digestion systems and quartz sample containers [137], quartz inserts [138,139], TFM inner vessels [140] or focused microwave ovens operating at atmospheric pressure and PTFE microsampling cups [141].

215

H. Matusiewicz

To summarize this section, use of acid vapor-phase digestion and attack of some organic and inorganic matrices as a sample preparation method is a convenient and useful technique. Closed pressure systems are the technique of choice to avoid losses of elements by volatilization while still maintaining extremely low values for the blank (by application of isopiestic distillation of the reagents and technical grade acids). 6.5.5 Efﬁciency of wet digestion (decomposition and dissolution) procedures Quality control is becoming increasingly more signiﬁcant in analytical chemistry. However, it is presently applied primarily to measurement techniques and not to sample preparation. For quality control in sample digestion, it is necessary to measure and record certain parameters exactly to be able to subsequently trace the course of the digestion process. In spite of that, complete decomposition of the sample is required to achieve reproducible and accurate elemental results by instrumental analytical methods. This is particularly the case for all voltammetric and polarographic determinations [142–145]. Interferences caused by incomplete decomposed organic compounds also occur, to a certain degree, when using atomic spectrometric methods such as AAS [146,147], ICP-OES [148,149], and ICPMS [150,151]. As noted earlier, nitric acid is the most frequently utilized sample dissolution medium. Unfortunately, the carbon contained in organic materials is only partly converted to CO2 by HNO3 at temperatures of up to 2008C (maximum operating temperature of PTFE vessels) [26]. In these cases, extending the digestion time and increasing the quantity of nitric acid do not improve the extent of decomposition. In principle, temperature and digestion time ultimately determine the effectiveness of a digestion, with RCC serving as a useful measure of quantitative assessment [152–155]; in other words, the highest temperatures are required to achieve a decomposition as complete as possible [156,157]. It should be noted here that the usefulness of the decomposition technique should be judged not from a visual point of view, because it often happens that a clear, colorless solution, indistinguishable from water, still contains signiﬁcant amounts of carbon. In closed systems the pressure depends not only on the temperature but also on the type and quantity of the sample, the size of the vessel, and the nature and quantity of the decomposition reagent. This pressure is not responsible for the decomposition quality, but nevertheless it should be controlled automatically. Wu¨rfels et al. [157 –159] described the extremely strong impact from residual organic compounds on elemental determinations by means of inverse voltammetry and demonstrated that a temperature of 300 – 3208C is necessary for pressurized sample digestion with pure nitric acid to obtain a solution containing less than 0.1% carbon. Otherwise, trace elements cannot be determined with inverse voltammetry. This was conﬁrmed by

216

Wet digestion methods

Wasilewska et al. [160], who showed that for complete oxidation of organic compounds with nitric acid, the decomposition temperature should be raised to 3008C. The inﬂuence of the digestion equipment (either thermal or microwave) is negligible if the digestion time employed is long enough to reach the steadystate temperature. Sample digestion with nitric acid between 220 and 2508C (most commercial equipment is able to fulﬁll this prerequisite) leads to RCCs in the low percentage range. The mode of heating of the digestion vessels is more and more supplanted by microwave technology; therefore, microwave-assisted wet digestion is a frequently used sample preparation technique for trace element determinations in organic materials. Studies of the RCC as a measure of decomposition efﬁciency have been undertaken [134,161 –165]. Using gas chromatography, Stoeppler et al. [152] quantiﬁed the ashing ability of conventional pressurized decomposition. Differences between the carbon content in the original sample and the carbon converted to CO2 showed that the investigated biological and environmental samples were not completely ashed with nitric acid. Wu¨rfels and Jackwerth [166] determined the residual carbon in samples digested under pressure or evaporated with HNO3. In most cases, microwave digestion of biological material was incomplete. Subsequently, the undigested compounds were identiﬁed [156]. Parallel to Wu¨rfels and Jackwerth’s studies [166], the residual organic species in nitric acid digests of bovine liver were identiﬁed by Pratt et al. [167]. Kingston and Jassie [168] evaluated the dissolution of several biological and botanical samples wet digested with HNO3. Free amino acid concentrations of human urine samples were typically reduced by a factor of 105. This reﬂects the comparative efﬁciency of protein hydrolysis, and is not necessarily equivalent to the total carbon oxidation efﬁciency. Nakashima et al. [163] investigated the digestion efﬁciency of HNO3 –HClO4 mixtures. The total RCC in a number of digested marine biological reference material (NRCC TORT-1) solutions was determined and used as a relative measure of the efﬁciency of various digestion schemes. Two-stage microwave digestions (i.e. conventional digestion followed by cooling, venting of excess gas from the bomb, re-capping, and re-heating) were superior to single-stage digestions. However, even the two-stage procedures were not complete and 24% carbon remained. The determination of residual carbon in digests of biological material with simultaneous ICP-OES analysis was described by Hee and Boyle [164] and Krushevska et al. [165]. The oxidation efﬁciencies of different dry and wet ashing procedures for milk samples were compared by Krushevska et al. [169], who noted that the residual carbon concentrations obtained with mediumpressure microwave digestions varied between 5 and 15%. Oxidizing mixtures of H2O2 or H2SO4 with HNO3 applied in a medium pressure (11 bar) microwave system did not yield a digestion efﬁciency higher than that for pure nitric acid (total acid volume was kept constant). Thus, until now, no suitable closed lowor medium-pressure microwave-heated oxidation system has been available to completely decompose biological samples leaving no carbon residue. In spite of

217

H. Matusiewicz

that, the safest way to obtain total mineralization is to complete these decomposition techniques with the addition of perchloric acid followed by heating until white perchloric fumes appear. However, with the high pressure/ temperature focused-microwave-heated TFM-Teﬂon bomb device, organic material is totally oxidized with nitric acid in a single-step procedure [101,103] (the closed TFM-Teﬂon focused-microwave-heated bomb enables very high pressure and temperature to be reached). Continuous or stopped ﬂow on-line microwave-heated digestion appears very attractive because it can increase the sample throughput and extend the automation of sample handling. However, on-line microwave-heated digestion was a priori expected to be associated with the problem of incomplete digestion of organic matter, because complete decomposition can only be achieved under vigorous conditions requiring high temperature and pressure. Matusiewicz and Sturgeon [170] critically evaluated on-line and high-pressure/temperature closed-vessel techniques with regard to efﬁciency of digestion. The completeness of destruction of biological materials (standard and certiﬁed reference materials) was characterized in terms of their RCC in the solution following digestion. Pressurized decomposition in a TFM-Teﬂon vessel was the most effective procedure (organic material was totally oxidized with nitric acid in a single-step procedure), whereas urine and sewage plant efﬂuent were incompletely decomposed (between 56 and 72%) with on-line microwave-heated digestion using nitric acid, nitric acid and hydrogen peroxide, and peroxydisulphate oxidation. Very recently, the residual weight of a bottom anti-reﬂective coating (BARC) sample was successfully used as an indicator to evaluate the digestion kinetics [171]. The weight degradation rate was independent of the sample weight under various temperatures, but was strongly dependent on the digestion acid volume and the digestion temperature. Mathematical modeling for prediction of digestion efﬁciency for the BARC sample was achieved by employing digestion kinetics as the backbone. With empirical ﬁtting of a pre-exponential factor, a novel equation incorporating the temperature, acid digestion volume, digestion time, and sample weight was developed. As a result, appropriate digestion parameters could be logically evaluated using the resultant model to achieve the desired digestion efﬁciency. Hydrogen peroxide is a very popular oxidizing reagent as it is converted to water and oxygen during the oxidation of biological material [134,172 –174]. No acid corrosion of the digestion vessel PTFE walls, no formation of insoluble salts with an acid anion, and no change of the sample matrix by an acid are additional advantages. Because of its strong oxidation power, only small amounts of H2O2 need be used, so that concentrated sample solutions can be obtained. Furthermore, high purity H2O2 is available and low blank concentration values (and thus low detection limits) can be achieved. However, explosion can occur if too much H2O2 is present. In addition, experiments with HNO3 –H2O2 mixtures conducted by Matusiewicz et al. [134] showed that all versions of pressurized microwave digestion with HNO3 and H2O2 gave

218

Wet digestion methods

incomplete digestion. No signiﬁcant improvement in the efﬁciency was achieved with 50% H2O2. The extension of this observation to mediumpressure and high-temperature microwave heating provided veriﬁcation of this observation [175]. Nitric acid digestion with the addition of H2O2 did not enhance digestion efﬁciency in this study compared to use of only HNO3. Wet acid digestion of aquatic samples is one of the most efﬁcient processes; however, it is time consuming and there is risk of contamination by the reagents used. Thus, an alternative oxidizing reagent is desirable to completely and safely decompose organic carbon residues. The application of ozone seems to have advantages in this respect. It was found that ozone is very effective in destroying natural organic compounds [176 –178], and has the potential to be used as an additional decomposition and/or ﬁnishing reagent [179]. Improvement in the pressurized microwave-assisted digestion procedure was achieved by adding optimum concentrations of strong oxidizing agents such as ozone [175]. The digestion efﬁciency of an optimal nitric acid system was improved by 13% by addition of ozone, with the further advantage that the agent does not contribute to the blank. A single digestion procedure is often insufﬁcient for the complete decomposition of a complex matrix, leading some authors to recommend a combination of two or more techniques. Two examples will sufﬁce to illustrate the principle [104,143]. First, pressure digestion followed by UV photolysis. Thus, it has been shown that analysis of olive leaves for heavy metals by voltammetric methods leads to distorted results after “pressure digestion” alone. Reliable data can be obtained only by supplementing the digestion with UV irradiation to ensure adequate decomposition of the matrix [143]. Second, a novel microwave-assisted high-temperature UV digestion procedure was developed for the accelerated decomposition of interfering dissolved organic carbon prior to trace element determination in liquid samples. This new technique signiﬁcantly improved the performance of the process of UV digestion (oxidation) and is especially useful for ultratrace analysis due to its extremely low risk of contamination [104]. In order to investigate the completeness of dissolution of inorganic materials, the recovery (or incomplete recovery) and accuracy of major, minor, and trace element determinations are usually applied. If silicates are present, which is usually the major inorganic component of many matrices (i.e. soils, sediments, sludges, ceramics, and other similar samples), the use of HF to achieve complete dissolution is mandatory [180,181]. 6.5.6

Comparison of wet digestion techniques

A careful comparison of several digestion techniques is the only way of assuring accurate results, particularly when little experience is available with respect to the digestion of a speciﬁc matrix, or existing reports are contradictory. The analyst must choose the sample preparation technique carefully to ensure that

219

H. Matusiewicz

the system is optimal for the analyses at hand. However, there is still no universal sample preparation system. With respect to requirements speciﬁc to contamination or losses through volatilization or retention, convection-heated or microwave-assisted wet digestion, quartz-lined high-pressure wet digestion, UV digestion, and vapor-phase acid digestion seem to be the best choice. However, all of these techniques require considerable investment for apparatus. Digestion of samples in an open vessel presents a serious risk of signiﬁcant analyte loss, despite the use of a reﬂux condenser. As far as economic aspects are concerned (low procurement, short digestion time, high sample throughput), microwave-assisted wet digestion and especially microwaveassisted pressurized on-line digestion appear to rank high. According to completion of the digestion, complete degradation of many samples is achieved only through high-pressure, high-temperature Teﬂon- or quartz-lined pressure vessel digestion, or by combination of a closed wet digestion system with UV irradiation. Table 6.4 summarizes the advantages and disadvantages of the wet digestion techniques discussed in Section 6.5 with respect to losses of analytes, blank levels, contamination problems, sample size, digestion time, degree of digestion, and economic aspects. 6.5.7

Digestion systems (instrumentation, equipment, automation)

Presently, the instrumentation market offers many devices to make wet digestion more efﬁcient and easier to manage by means of possible automation, but this is achieved principally with microwave energy. Wet digestions in open vessels are undertaken with or without reﬂuxing. Because it is very critical to adhere very closely to the optimized time and temperature digestion parameters, mechanization of the digestion not only leads to higher sample throughput with less human intervention but also to the avoidance of errors. The simplest form of mechanization can be implemented through a time (programmable timer) and temperature (via an autotransformer) controlled heating block. There are many models of heating blocks on the market. A greater degree of mechanization would also incorporate control of reagent reﬂux during digestion. These procedures operate batch-wise. Continuous sample handling has some advantages over discontinuous handling; the former generally better matches analytical needs. The automated wet digestion device (VAO, Anton Paar, Austria) is such a continuously operating digestion system, an ideal instrument for laboratories requiring high throughput of similar samples with which all methods of wet chemical digestions can be performed [182]. With the help of a microprocessor, all important digestion parameters are controlled. Automation controls the time – temperature/pressure program for sample digestion, so that different sample materials can be processed under optimum conditions. The loading or charging of the high-pressure asher with sample

220

TABLE 6.4 Advantages and disadvantages of wet digestion methods Digestion technique

Open systems Conventional heating

Possible way of losses

Volatilization

Microwave heating

Volatilization

UV digestion

None

Closed systems Conventional heating Microwave heating

Source of blank

Acids, vessels, air Acids, vessels, air

Sample size (g)

Maximum

Digestion time

Degree of digestion

Economical aspects

Inexpensive, needs supervision Inexpensive, needs supervision Inexpensive, needs supervision

Organic

Inorganic

Temperature (8C)

Pressure (bar)

,5

,10

,400

Several hours

Incomplete

,5

,10

,400

,1 h

Incomplete

Several hours

High

,90

Liquid

Retention

Acids (low)

,0.5

,3

,320

,150

Several hours

High

Retention

Acids (low)

,0.5

,3

,300

,200

,1 h

High

Needs no supervision Expensive, needs no supervision continued

TABLE 6.4 (continuation) Digestion technique

Possible way of losses

Source of blank

Sample size (g) Organic

Inorganic

Temperature (8C)

Pressure (bar)

,0.1 (slurry)

,0.1 (slurry)

,320

.300

Flow systems Conventional heating

Incomplete digestion

Acids (low)

UV on-line digestion

Incomplete digestion

None

Microwave heating

Incomplete digestion

Acids (low)

,0.1 (slurry)

None

,0.1

Vapor-phase None acid digestion

Maximum

,90

Liquid

Digestion time

Degree of digestion

Economical aspects

Several minutes

High

Several minutes

High

Expensive, needs no upervision Inexpensive, needs no supervision Expensive, needs no supervision

,0.3 (slurry)

,250

,40

Several minutes

High

,0.1

,200

,20

,1 h

High

Needs no supervision

Wet digestion methods

material is achieved manually. A fully automated version of this high-pressure asher is not available. Berghof pressure digestion systems [183] serve for sample preparation of inorganic and organic matrices at high temperature (max. 200 – 2508C) and high pressure (max. 100 and 200 bar) in pure, isostatically pressed PTFE or quartz vessels. As noted already, three basic types of microwave-assisted digestion systems have evolved: atmospheric pressure, elevated pressure (closed vessel), and ﬂow-through, working in the two most common modes: multimode cavity and focused-type (waveguide). Reviews of commercially available microwaveassisted digestion systems and vessels (summary of the vessels, ovens, and oven systems) are given in Refs. [29–32,35,37,41,184 –186] together with speciﬁcations and features for elevated-pressure, atmospheric pressure, and ﬂow-through units. The reader should also consult Chapter 8 of this volume. The simplicity and efﬁcacy of microwave digestion easily lends itself to automation and robotics. Systems have been developed that are capable of weighing samples, adding acids, capping and uncapping vessels, accomplishing microwave-assisted digestion, diluting digestates, transferring vessels, and even cleaning and reusing the vessels. Once such a system is operational, the only things the analyst has to do is supply and place the representative sample(s) in locations recognized by the system and then initiate the controlling program (Table 18 in Ref. [35] primarily summarizes the application and functioning of these systems). 6.5.8

Safety of acid digestions (sample acid digestion safety)

The reagents, instruments, and operations employed in the digestion of materials are potentially hazardous, even when used as directed. The operator must always be properly protected with a laboratory coat, gloves, and safety glasses or, better still, face protection. Some concentrated fuming acids (HF, HNO3, HCl) are to be handled only in a well-ventilated hood. Oxidizing acids (HNO3, HClO4) are more hazardous than non-oxidizing acids (HCl, H3PO4, HF), being more prone to explosion, especially in the presence of reducing agents, such as organic matter. Perchloric acid is oxidizing only when it is concentrated and hot; it must never be brought into contact with organic matter unless diluted with nitric acid. Acid digestion must be conducted in a fume cupboard with efﬁcient scrubbers installed. The evaporation of perchloric acid is to be performed only in an appropriate stainless steel, stoneware, or polypropylene hood, with washing facilities to eliminate any perchlorate deposit. Great care should be taken when using “pressure digestion” methods. Pressure digestion vessels (bombs) contain the acid fumes and are useful for rapid, one-step digestions without losses. But, again, there are restrictions; in some reactions (especially spontaneous) potentially explosive gases are

223

H. Matusiewicz

produced that exceed the safety limits of the vessels. For instance, nitric acid and especially the spontaneous HNO3 and H2O2 digestion of organic matter in a closed vessel may result in explosion due to unintended pressure build-up within the vessel. These systems produce high-pressure spikes, which can be avoided by decreasing the sample weight or by applying a gradual temperature increase. Microwave-assisted sample digestion has its own safety requirements. As a result of the direct energy absorption and rapid heating, microwave techniques introduce unique safety considerations that are not encountered in other methods. Differences in conditions between traditional laboratory practices and microwave-implemented methods should be examined before microwave energy is applied to heat reagents or samples. An excellent summary of this aspect is extensively reviewed in Refs. [30,31].

6.6

CONCLUSIONS AND FUTURE TRENDS

The chief methods used for the digestion of organic and inorganic samples have been evaluated. A brief summary of applications of these techniques to various sample matrices is presented in Table 6.5. The variety of approaches currently available for the digestion of solid and liquid samples allows the most suitable method to be selected for each application, depending on both the matrix and type of analyte, and the subsequent steps to be developed in order to complete the analytical process. In spite of that, it is fair to point out that sample digestion must not be looked at as an isolated step, but one that needs to be integrated into the entire analytical process. Attention has been focused on digestion at elevated temperature and pressure. High-pressure digestion with its large digestion temperature range is the most universal digestion system at present, and is the technique of choice for the vast majority of both inorganic and organic materials. New ways to further increase the efﬁciency of sample preparation should continue with development of hyphenated digestion techniques. A novel, microwaveassisted, high-temperature UV digestion for accelerated decomposition of dissolved organic compounds or slurries was developed [104]. This new technique is ideal for extreme trace analysis due to the low blank values and low acid concentration. In addition, this digestion method can be used for the determination of non-metals by ion chromatography. Alternatively, within the limits of the Teﬂon-lined digestion vessels, improvement in the digestion efﬁciency can be achieved by adding optimum concentrations of strong oxidizing agents, such as ozone or oxygen, which appear to be efﬁcient digestion agents for the treatment of biological material. Again, this has the advantage that the agent does not contribute to the analysis blank. It should be mentioned that vapor-phase acid digestion offers an alternative solution to

224

TABLE 6.5 Summary of applications of total wet digestion procedures to the analysis of materials (determination of elements) Required acid(s)a

Digestion technique (modeb)

Reference

Water(s) Environmental samples Coal Coal ﬂy ash Dust Catalysts Waste materials Sewage sludge Waste water Biological samples Botanicals Plants Clinical Marine Forensic Food(s) Beverages Silicates Soils Sediments Glasses Geological samples Rocks

H2O2, HNO3

UV radiation

[49]

HNO3, HCl, HF Aqua regiac þ HFd Aqua regia þ HF Aqua regia

Open or closed system Open or closed system Open or closed system Open systems

[20,31,35,38] [20,31,35,38] [20,31,35,38] [20,21]

HNO3, HCl HNO3

Open or closed or ﬂow systems Flow systems

[18,31] [31,39]

HNO3 þ H2O2 þ HF HNO3 þ H2O2 þ HF HNO3 HNO3 HNO3 HNO3 HNO3, H2O2

Open or closed system Open or closed system Open or closed system Open or closed system Open or closed system Open or closed system Open or closed or ﬂow systems

[15,16,18,31] [15,16,18,31] [15,16,18,31] [15,16,18,31] [41] [31,41] [41]

Aqua regia þ HF Aqua regia þ HF HF

Open and/or closed systems Open and/or closed systems Open systems

[19–22,35] [19–22,35] [19–22]

Aqua regia þ HFe

Open or closed systems

[20–22,31,33] continued

Wet digestion methods

225

Material/matrix/sample

226

TABLE 6.5 6.4 (continuation) Required acid(s)a

Digestion technique (modeb)

Reference

Ores Minerals Petroleum products Fuels Oils Drugs and pharmaceuticals Metals Ferrous Non-ferrous Alloys Steels Chemicals Polymers Refractory compoundsg Ceramics Composites Nuclear materials

Aqua regia þ HF HF þ H2SO4, HCl

Open or closed systems Open systems

[20– 22,31,33] [20– 22,31,33]

HNO3 þ HCl HNO3 þ HCl HCl, HNO3

Open or closed system Open or closed system Open systems

[23,31] [23,31] [41]

HNO3 þ (HF or HNO3 or H2SO4) HCl or HNO3 or HF Aqua regia þ HF HCl þ HNO3, HClO4f HCl, HNO3, HF, H2SO4 HCl, HNO3, HF, H2SO4

Open systems Open systems Open systems Open systems Open or closed systems Open or closed systems

[41] [41] [41] [41] [20,23] [23,41]

HNO3, HCl, HF, H2SO4, H2O2 HNO3, HCl, HF, H2SO4, H2O2 HNO3 or HCl, H3PO4, HClO4

Open or closed systems Open or closed systems Open or closed systems

[20] [20] [21]

a

Concentrated acids are usually employed; H2O2 is 30%; in most cases alternative digestions are possible depending on requirements of analyst. b Conventional or microwave. c Unstable. d Use only Teﬂon vessels, the addition of HF is required to obtain quantitative recoveries for Cr. e Addition of H3BO3 to neutralize the HF by forming tetraﬂuoro-boric acid. f Danger of explosion. g Certain refractory materials are not decomposed; these must be solubilized by fusion.

H. Matusiewicz

Material/matrix/sample

Wet digestion methods

these problems: reduced concentration of acid in the digestate and the possibility of using a technical grade acid without any deterioration of the analytical blank. Another example where signiﬁcant improvement in digestion and dissolution was obtained is the use of a reactor that combines microwave and ultrasound energy [48]. It is expected that these two methods could open a new research ﬁeld “combined digestion techniques”. It can be said with certainty that the majority of all digestions will be performed in the future by means of microwave assistance. Progress has been made over the past several years in reducing systematic errors and improving detection limits with microwave digestion, as well as its automation. A noticeable trend toward pressurized closed-vessel systems permitting hightemperature decomposition compatible with trace analysis has occurred. While some researchers advocate high-pressure (.100 bar) digestion at 250 –3008C to destroy interferences in refractory compounds, manufacturers are working to devise sample vessels that can withstand these conditions. There has been a growing trend in recent years toward development of fully automated on-line analysis techniques. Microwave-assisted high-pressure ﬂow digestion system with PTFE or PFA tubes for digestion temperatures up to 2508C opens up new possibilities for fully automated sample preparation [119]. On the other hand, the development of new high-temperature/high-pressure ﬂow digestion systems that incorporate resistively heated capillaries for the continuous digestion of various samples coupled with atomic spectrometric instruments has arisen [107 –109]. It is predicted that ﬂow systems will become dominant for liquid samples and slurries and extend the analytical capabilities of instrumental methods by combining sample preparation with simultaneous analysis using only micrograms of sample and microliters of reagents. The ﬁnal goal of these studies should be the adaptation of standard batch digestion methods to on-line systems combining ﬂow-through digestion directly to analyzers. It is evident that wet digestion methods will remain a fertile area for development. New digestion techniques need to be designed that address the limitations of the instrumentation and maximize its potential. Development trends for conventional and microwave instruments will focus on sample throughput, enhanced vessel performance speciﬁcations, the use of new materials, further reﬁnement of in situ vessel control (direct temperature and pressure, incident and reﬂected microwave power), and computer-controlled sample digesters with automated capability. Finally, the development of automated methods for wet digestion of solid samples without human participation can only be achieved with the use of a robotic station. Nevertheless, a number of auxiliary energies and commercially available modules can facilitate and/or accelerate one of the most timeconsuming steps of the analytical process, i.e. to obtain the analyte(s) from a solid sample in the form of a solution.

227

H. Matusiewicz

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

17 18

19 20 21

22 23

24

25 26 27 28 29 30

228

A. Duﬂos, Hand. d. Pharm. Chem. Praxis, 2. Auﬂ., I. Max u. Comp., Breslau, (1938) 534. R. Fresenius and L.v. Babo, Justus Liebigs Ann. Chem., 49 (1844) 287. F.P. Danger and C. Flandin, C.R. Acad. Sci., 12 (1841) 1089. L. Carius, Justus Liebigs Ann. Chem., 116 (1860) 1; L. Carius, Justus Liebigs Ann. Chem., 136 (1865) 129. J. Kjeldahl, Z. Anal. Chem., 22 (1883) 366. A. Classen and O. Bauer, Ber. Dtsch. Chem. Ges., 16 (1884) 1061. A. Stcherbak, Arch. Med. Exp., 5 (1893) 309. D.D. Van Slyke, Anal. Chem., 26 (1954) 1706. D. Polley and V.L. Miller, Anal. Chem., 27 (1955) 1162. H.J.H. Fenton, J. Chem. Soc. (London), 65 (1894) 899. B. Sansoni, O. Sigmund, E. Bauer-Schreiber, W. Wiegand and L. Perrera, Angew. Chem., 73 (1961) 763. B. Sansoni and W. Kracke, Z. Anal. Chem., 243 (1968) 209. A.M. Ure, L.R.P. Butler, R.O. Scott and R. Jenkins, Pure Appl. Chem., 60, (1988) 1461. B. Griepink and G. Tolg, Pure Appl. Chem., 61 (1989) 1139. T.T. Gorsuch, The Destruction of Organic Matter. Pergamon Press, Oxford, 1970. B. Sansoni and V.K. Panday, Ashing in trace element analysis of biological material. In: S. Fachetti (Ed.), Analytical Techniques for Heavy Metals in Biological Fluids. Elsevier, Amsterdam, 1983, p. 91. K.S. Subramanian, Spectrochim. Acta, Part B, 51 (1996) 291. G.V. Iyengar, K.S. Subramanian and J.R.W. Woittiez, Sample decomposition. In: Element Analysis of Biological Samples. Principles and Practice CRC Press, Boca Raton, FL, 1998, p. 103. Z. Sˇulcek, P. Povondra and J. Dolezˇal, CRC Crit. Rev. Anal. Chem., 6 (1977) 255. Z. Sˇulcek and P. Povondra, Methods of Decomposition in Inorganic Analysis. CRC Press, Boca Raton, FL, 1989. P. Povondra and Z. Sˇulcek, Modern methods of decomposition of inorganic substances. In: J. Zy´ka (Ed.), Instrumentation in Analytical Chemistry. Ellis Horwood, New York, 1991, p. 188. T.T. Chao and R.F. Sanzolone, J. Geochem. Explor., 44 (1992) 65. R. Bock, A Handbook of Decomposition Methods in Analytical Chemistry, translated and revised by I.L. Marr. International Textbook Company, Glasgow, 1979. S. Bajo, Dissolution of matrices. In: Z.B. Alfassi and C.M. Wai (Eds.), Preconcentration Techniques for Trace Elements. CRC Press, Boca Raton, FL, 1992, p. 3. C. Vandecasteele and C.B. Block, Sample preparation. In: Modern Methods for Trace Element Determination. Wiley, Chichester, 1993, p. 9. M. Stoeppler (Ed.), Sampling and Sample Preparation. Springer, Berlin, 1997. M. Hoenig and A.-M. de Kersabiec, Spectrochim. Acta, Part B, 51 (1996) 1297. M. Hoenig, Talanta, 54 (2001) 1021. E. Krakovska´ and H.-M. Kuss, Rozklady v Analitickej Che´mii. VIENALA, Kosˇice, Slovakia, 2001, in Slovak. R.T. White, Jr, Open reﬂux vessels for microwave digestion: botanical, biological, and food samples for elemental analysis, In: H.M. Kingston and L.B. Jassie (Eds.),

Wet digestion methods

31

32 33 34 35 36 37

38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63

Introduction to Microwave Sample Preparation. Theory and Practice. ACS, Washington, DC, 1988. J.M. Mermet, Focused-microwave-assisted reactions: atmospheric-pressure acid digestion, on-line pretreatment and acid digestion, volatile species production, and extraction, In: H.M. “Skip” Kingston and S.J. Haswell (Eds.), MicrowaveEnhanced Chemistry. Fundamentals, Sample Preparation, and Applications. ACS, Washington, DC, 1997. H. Matusiewicz and R.E. Sturgeon, Prog. Anal. Spectrosc., 12 (1989) 21. H.-M. Kuss, Fresenius J. Anal. Chem., 343 (1992) 788. A. Zlotorzynski, Crit. Rev. Anal. Chem., 25 (1995) 43. F.E. Smith and E.A. Arsenault, Talanta, 43 (1996) 1207. M. Burguera and J.L. Burguera, Quim. Anal., 15 (1996) 112. H.M. “Skip” Kingston and P.J. Walter, The art and science of microwave sample preparations for trace and ultratrace elemental analysis. In: A. Montaser (Ed.), Inductively Coupled Plasma Mass Spectrometry. Wiley-VCH, New York, 1998, p. 33. K.J. Lamble and S.J. Hill, Analyst, 123 (1998) 103R. M. Burguera and J.L. Burguera, Anal. Chim. Acta, 366 (1998) 63. R. Chakraborty, A.K. Das, M.L. Cervera and M. de la Guardia, Fresenius J. Anal. Chem., 355 (1996) 99. R.A. Meyers (Ed.), Encyclopedia of Analytical Chemistry. Wiley, Chichester, 2000. E.C. Kuehner, R. Alvarez, P.J. Paulsen and T.J. Murphy, Anal. Chem., 44 (1972) 2051. R.M. Barnes, S.P. Quina´ia, J.A. No´brega and T. Blanco, Spectrochim. Acta, Part B, 53 (1998) 769. P.O. Bethge, Anal. Chim. Acta, 10 (1954) 317. R.A. Nadkarni, Anal. Chem., 56 (1984) 2233. A. Abu-Samra, J.S. Morris and S.R. Koirtyohann, Anal. Chem., 47 (1975) 1475. J.A. No´brega, L.C. Trevizan, G.C.L. Arau´jo and A.R.A. Nogueira, Spectrochim. Acta, Part B, 57 (2002) 1855. A. Lagha, S. Chemat, P.V. Bartels and F. Chemat, Analusis, 27 (1999) 452. J. Golimowski and K. Golimowska, Anal. Chim. Acta, 325 (1996) 111. M. Trapido, A. Hirvonen, Y. Veressinina, J. Hentunen and R. Munter, Ozone Sci. Eng., 19 (1997) 75. A. Mitscherlich, J. Prakt. Chem., 81 (1860) 108. G.L. Carius, Ber. Dtsch. Chem. Ges., 3 (1870) 697. C.L. Gordon, J. Res. Natl. Bur. Stand., 30 (1943) 107. E. Wichers, W.G. Schlecht and C.L. Gordon, J. Res. Natl. Bur. Stand., 33 (1944) 363. E. Wichers, W.G. Schlecht and C.L. Gordon, J. Res. Natl. Bur. Stand., 33 (1944) 451. S.E. Long and W.R. Kelly, Anal. Chem., 74 (2002) 1477. P. Jannasch, Z. Anorg. Allgem. Chem., 6 (1894) 72. E. May and J.J. Rowe, Anal. Chim. Acta, 33 (1965) 648. E. Jackwerth and S. Gomisˇcˇek, Pure Appl. Chem., 56 (1984) 479. J. Ito, Bull. Chem. Soc. Jpn, 35 (1962) 225. F.J. Langmyhr and S. Sveen, Anal. Chim. Acta, 32 (1965) 1. B. Bernas, Anal. Chem., 40 (1968) 1682. K. Okamoto and K. Fuwa, Anal. Chem., 56 (1984) 1758.

229

H. Matusiewicz 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78

79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 100 101

230

M. Ravey, B. Farberman, I. Hendel, S. Epstein and R. Shemer, Anal. Chem., 67 (1995) 2296. R. Uhrberg, Anal. Chem., 54 (1982) 1906. L. Kotz, G. Henze, G. Kaiser, S. Pahlke, M. Veber and G. To¨lg, Talanta, 26 (1979) 681. M. Stoeppler and F. Backhaus, Fresenius Z. Anal. Chem., 291 (1978) 116. L. Kotz, G. Kaiser, P. Tscho¨pel and G. To¨lg, Z. Anal. Chem., 260 (1972) 207. M. Tomljanovic and Z. Grobenski, At. Absorption Newslett., 14 (1975) 52. T. Uchida, I. Kojima and C. Iida, Anal. Chim. Acta, 116 (1980) 205. M. Stoeppler, K.P. Mu¨ller and F. Backhaus, Fresenius Z. Anal. Chem., 297 (1979) 107. M. Takenaka, S. Kozuka, M. Hayashi and H. Endo, Analyst, 122 (1997) 129. P.J. Lechler, M.O. Desilets and F.J. Cherne, Analyst, 113 (1988) 201. G. Knapp, ICP Information Newslett., 10 (1984) 91. G. Knapp, Fresenius Z. Anal. Chem., 317 (1984) 213. G. Knapp, Int. J. Environ. Anal. Chem., 22 (1985) 71. S. Strenger and A.V. Hirner, Fresenius J. Anal. Chem., 371 (2001) 831. P. Kettisch, B. Maichin, M. Zischka and G. Knapp, Trends in Sample Preparation 2002 Development and Application. (Conference, Abstract T-11), Seggau, Austria, 2002. M. Hale, M. Thompson and J. Lovell, Analyst, 110 (1985) 225. D.W. Eggimann and P.R. Betzer, Anal. Chem., 48 (1976) 886. W.J. Adrian, At. Absorption Newslett., 10 (1971) 96. R.W. Kuennen, K.A. Wolnik, F.L. Fricke and J.A. Caruso, Anal. Chem., 54 (1982) 2146. K. May and M. Stoeppler, Fresenius Z. Anal. Chem., 317 (1984) 248. H. Matusiewicz, Chem. Anal. (Warsaw), 28 (1983) 439. S.A. Matthes, R.F. Farrell and A.J. Mackie, Tech. Prog. Rep.—US Bur. Mines, 120 (1983) 9. F. Smith, B. Cousins, J. Bozic and W. Flora, Anal. Chim. Acta, 177 (1985) 243. O. Buresch, W. Ho¨nle, U. Haid and H.G.v. Schnering, Fresenius Z. Anal. Chem., 328 (1987) 82. J. Alvarado, L.E. Leo´n, F. Lo´pez and C. Lima, J. Anal. At. Spectrom., 3 (1988) 135. I. Kojima, T. Uchida and C. Iida, Anal. Sci., 4 (1988) 211. H.M. Kingston and L.B. Jassie, Anal. Chem., 58 (1986) 2534. L.Q. Xu and W.X. Shen, Fresenius Z. Anal. Chem., 332 (1988) 45. J.R.P. Nicholson, M.G. Savory, J. Savory and M.R. Wills, Clin. Chem., 35 (1989) 488. S. Baldwin, M. Deaker and W. Maher, Analyst, 119 (1994) 1701. H.J. Reid, S. Greenﬁeld, T.E. Edmonds and R.M. Kapdi, Analyst, 118 (1993) 1299. H.J. Reid, S. Greenﬁeld and T.E. Edmonds, Analyst, 118 (1993) 443. G. Heltai and K. Percsich, Talanta, 41 (1994) 1067. M. Pougnet, S. Michelson and B. Downing, J. Microwave Power Electromagn. Energy, 26 (1991) 140. M. Pougnet, B. Downing and S. Michelson, J. Microwave Power Electromagn. Energy, 28 (1993) 18. G. Le´ge`re and E.D. Salin, Appl. Spectrosc., 49 (1995) 14A. G. Le´ge`re and E.D. Salin, Anal. Chem., 70 (1998) 5029. H. Matusiewicz, Anal. Chem., 66 (1994) 751.

Wet digestion methods 102 103 104 105 106 107 108 109 110 111

112 113 114 115 116 117 118 119 120 121 122 123 124 125 126 127 128 129 130 131 132 133 134 135 136 137 138

W. Lautenschla¨ger, Apparatus for performing chemical and physical reactions, US Patent 5,382,414, January 17, 1995. H. Matusiewicz, Anal. Chem., 71 (1999) 3145. D. Florian and G. Knapp, Anal. Chem., 73 (2001) 1515. E. Allen, Automation in Analytical Chemistry, Technicon Symposia, 1966, p. 247. T.J. Gluodenis and J.F. Tyson, J. Anal. At. Spectrom., 7 (1992) 301. C. Gra¨ber and H. Berndt, J. Anal. At. Spectrom., 14 (1999) 683. S. Haiber and H. Berndt, Fresenius J. Anal. Chem., 368 (2000) 52. P. Jacob and H. Berndt, J. Anal. At. Spectrom., 17 (2002) 1615. S.M.V. Fernandes, J.L.F.C. Lima and A.O.S.S. Rangel, Fresenius J. Anal. Chem., 366 (2000) 112. J.L. Burguera and M. Burguera, Flow injection systems for on-line sample dissolution/decomposition. In: A. Sanz-Medel (Ed.), Flow Analysis with Atomic Spectrometric Detectors. Elsevier, Amsterdam, 1999, pp. 135 –167, Ch. 5. M. Burguera, J.L. Burguera and O.M. Alarco´n, Anal. Chim. Acta, 179 (1986) 351. V. Carbonell, M. de la Guardia, A. Salvador, J.L. Burguera and M. Burguera, Anal. Chim. Acta, 238 (1990) 417. V. Karanassios, F.H. Li, B. Liu and E.D. Salin, J. Anal. At. Spectrom., 6 (1991) 457. S.J. Haswell and D. Barclay, Analyst, 117 (1992) 117. M. Burguera, J.L. Burguera and O.M. Alarco´n, Anal. Chim. Acta, 214 (1988) 421. L.J. Martines Stewart and R.M. Barnes, Analyst, 119 (1994) 1003. R.E. Sturgeon, S.N. Willie, B.A. Methven, J.W. Lam and H. Matusiewicz, J. Anal. At. Spectrom., 10 (1995) 981. U. Pichler, A. Haase, G. Knapp and M. Michaelis, Anal. Chem., 71 (1999) 4050. C.J. Mason, G. Coe, M. Edwards and P. Riby, Analyst, 125 (2000) 1875. H. Matusiewicz, Spectroscopy, 6 (1991) 38. Kh.I. Zilbershtein, M.M. Piriutko, T.P. Jevtushenko, I.L. Sacharnova and O.N. Nikitina, Zavod. Lab., 25 (1959) 1474. J.W. Mitchell and D.L. Nash, Anal. Chem., 46 (1974) 326. A.D. Thomas and L.E. Smythe, Talanta, 20 (1973) 469. M.A. Klitenick, C.J. Frederickson and W.I. Manton, Anal. Chem., 55 (1983) 921. J.F. Woolley, Analyst, 100 (1975) 896. Yu.A. Karpov and V.A. Orlova, Vysokochist. Veshchestva, 2 (1990) 40. D.M. Marinescu, Analusis, 13 (1985) 469. I. Kojima, F. Jinno, Y. Noda and C. Iida, Anal. Chim. Acta, 245 (1991) 35. H. Matusiewicz, Chem. Anal. (Warsaw), 33 (1988) 173. H. Matusiewicz, Chem. Anal. (Warsaw), 28 (1983) 439. V.G. Pimenov, A.N. Pronchatov, G.A. Maksimov, V.N. Shishov, E.M. Shcheplyagin and S.G. Krasnova, Zh. Anal. Khim., 39 (1984) 1636. D. Amaarsiriwardena, A. Krushevska, M. Argentine and R.M. Barnes, Analyst, 119 (1994) 1017. H. Matusiewicz, R.E. Sturgeon and S.S. Berman, J. Anal. At. Spectrom., 4 (1989) 323. H. Matusiewicz, R.E. Sturgeon and S.S. Berman, J. Anal. At. Spectrom., 6 (1991) 283. H. Matusiewicz, J. Anal. At. Spectrom., 4 (1989) 265. D. Amarasiriwardena, A. Krushevska and R.M. Barnes, Appl. Spectrosc., 52 (1998) 900. ´ va´ri, M. Garcia Tapia and Gy. Za´ray, Microchem. J., Zs. Cze´ge´ny, B. Berente, M. O 59 (1998) 100.

231

H. Matusiewicz 139 140 141 142 143 144 145 146 147 148 149 150 151 152 153 154 155 156 157 158 159 160 161 162 163 164 165 166 167 168 169 170 171 172 173 174 175 176

232

K. Eilola and P. Pera¨ma¨ki, Fresenius J. Anal. Chem., 369 (2001) 107. Y. Han, H.M. Kingston, R.C. Richter and C. Pirola, Anal. Chem., 73 (2001) 1106. G.C.L. Arau´jo, A.R.A. Nogueira and J.A. No´brega, Analyst, 125 (2000) 1861. M. Wu¨rfels, E. Jackwerth and M. Stoeppler, Fresenius Z. Anal. Chem., 329 (1987) 459. J. Hertz and R. Pani, Fresenius Z. Anal. Chem., 328 (1987) 487. P. Schramel and S. Hasse, Fresenius J. Anal. Chem., 346 (1993) 794. H.J. Reid, S. Greenﬁeld and T.E. Edmonds, Analyst, 120 (1995) 1543. J. Ne`ve, M. Hanocq, L. Molle and G. Lefebvre, Analyst, 107 (1982) 934. B. Welz, M. Melcher and J. Ne`ve, Anal. Chim. Acta, 165 (1984) 131. G. Knapp, B. Maichin and U. Baumgartner, At. Spectrosc., 19 (1998) 220. J. Machat, V. Otruba and V. Kanicky, J. Anal. At. Spectrom., 17 (2002) 1096. D. Ashley, At. Spectrosc., 13 (1992) 169. J. Begerov, M. Turfeld and L. Duneman, J. Anal. At. Spectrom., 12 (1997) 1095. M. Stoeppler, K.P. Mu¨ller and F. Backhaus, Fresenius Z. Anal. Chem., 297 (1979) 107. H. Matusiewicz, B. Golik and A. Suszka, Chem. Anal. (Warsaw), 44 (1999) 559. E.N.V.M. Carrilho, A.R.A. Nogueira, J.A. No´brega, G.B. de Souza and G.M. Cruz, Fresenius J. Anal. Chem., 371 (2001) 536. L.M. Costa, F.V. Silva, S.T. Gouveia, A.R.A. Nogueira and J.A. No´brega, Spectrochim. Acta, Part B, 56 (2001) 1981. M. Wu¨rfels, E. Jackwerth and M. Stoeppler, Anal. Chim. Acta, 226 (1989) 1. M. Wu¨rfels, E. Jackwerth and M. Stoeppler, Anal. Chim. Acta, 226 (1989) 17. M. Wu¨rfels, E. Jackwerth and M. Stoeppler, Fresenius J. Anal. Chem., 329 (1987) 459. M. Wu¨rfels, Mar. Chem., 28 (1989) 259. M. Wasilewska, W. Goessler, M. Zischka, B. Maichin and G. Knapp, J. Anal. At. Spectrom., 17 (2002) 1121. H. Matusiewicz, A. Suszka and A. Ciszewski, Acta Chim. Hung., 128 (1991) 849. A. Krushevska, R.M. Barnes, C.J. Amarasiriwaradena, H. Foner and L. Martines, J. Anal. At. Spectrom., 7 (1992) 851. S. Nakashima, R.E. Sturgeon, S.N. Willie and S.S. Berman, Analyst, 113 (1988) 159. S.S.Q. Hee and J.R. Boyle, Anal. Chem., 60 (1988) 1033. A. Krushevska, R.M. Barnes and C.J. Amarasiriwaradena, Analyst, 118 (1993) 1175. M. Wu¨rfels and E. Jackwerth, Fresenius Z. Anal. Chem., 322 (1985) 354. K.W. Pratt, H.M. Kingston, W.A. MacCrehan and W.F. Koch, Anal. Chem., 60 (1988) 2024. H.M. Kingston and L.B. Jassie, Anal. Chem., 58 (1986) 2534. A. Krushevska, R.M. Barnes, C.J. Amarasiriwaradena, H. Foner and L. Martines, J. Anal. At. Spectrom., 7 (1992) 845. H. Matusiewicz and R.E. Sturgeon, Fresenius J. Anal. Chem., 349 (1994) 428. F.-H. Ko and H.-L. Chen, J. Anal. At. Spectrom., 16 (2001) 1337. H. Matusiewicz and R.M. Barnes, Anal. Chem., 57 (1985) 406. R.S. Sah and R.O. Miller, Anal. Chem., 64 (1992) 230. E. Veschetti, D. Maresca, D. Cutilli, A. Santarsiero and M. Ottaviani, Microchem. J., 67 (2000) 171. H. Matusiewicz, Chem. Anal. (Warsaw), 46 (2001) 897. R.G. Clem and A.T. Hodgson, Anal. Chem., 50 (1978) 102.

Wet digestion methods 177 178 179 180 181 182 183 184

185 186

Zˇ. Filipovic´-Kovacˇevic´ and L. Sipos, Talanta, 45 (1998) 843. K. Sasaki and G.E. Pacey, Talanta, 50 (1999) 175. W. Jiang, S.J. Chalk and H.M. “Skip” Kingston, Analyst, 122 (1997) 211. P. Schramel, G. Lill and R. Seif, Fresenius Z. Anal. Chem., 326 (1987) 135. H. Matusiewicz, Mikrochim. Acta, 111 (1993) 71. K.W. Budna and G. Knapp, Fresenius Z. Anal. Chem., 294 (1979) 122. Stainless steel pressure vessels with Teﬂon inserts, Berghof Laborprodukte GmbH, Eningen, Germany. H. Matusiewicz, Development of high-pressure closed-vessel systems for microwave-assisted sample digestion in microwave-enhanced chemistry. In: H.M. “Skip” Kingston and S.J. Haswell (Eds.), Microwave-Enhanced Chemistry. Fundamentals, Sample Preparation, and Applications. ACS, Washington, DC, 1997, pp. 353– 369, Ch. 4. B. Erickson, Anal. Chem., 70 (1998) 467A. R.C. Richter, D. Link and H.M. “Skip” Kingston, Anal. Chem., 73 (2001) 31A.

233

This page intentionally left blank

Chapter 7

Dry ashing Michel Hoenig

7.1

GENERAL CONSIDERATIONS

Trace element determinations in most inorganic analytical laboratories are usually performed using atomic spectroscopic techniques. The market for this type of instrumentation was principally initially dedicated to the analysis of liquid samples. Only a few manufacturers provide equipment suitable for direct analysis of solid samples: X-ray ﬂuorescence spectrometry (XRF) and arc/spark optical emission spectrometry (OES). These two techniques were intensively utilized between the 1950s and 1970s; presently, they cover, with difﬁculty, environmental applications necessitating determinations of elements at trace or ultra-trace concentrations. These techniques remain in use in several industrial domains, however, owing to several practical aspects, i.e., analyses can often be performed directly on solid samples, overcoming the problems associated with dissolution procedures. In environmental monitoring and health diagnostics, the determination of trace elements has, since the 1970s, generally been tackled using atomic absorption and atomic emission techniques. Flame atomic absorption spectrometry (F-AAS) offers detection power ranging from the mg/l to the mg/l level, depending on the element considered. Electrothermal atomic absorption spectrometry (ET-AAS) is able to address, on average, the mg/l range. However, AAS techniques are limited to the determination of metallic elements only. During the 1970s, inductively coupled plasma optical emission spectrometry (ICP-OES) appeared, exhibiting a detection power lying between F-AAS and ET-AAS. Recently, during the 1980s, instrumentation resulting from the coupling of an ICP excitation source with a mass spectrometer (ICP-MS) became commercially available. The detection power of ICP-MS is very high, on the order of ng/l or lower. Both ICP-OES and ICP-MS permit the multielement determination of metals and non-metals. Moreover, in comparison with AAS, particularly ET-AAS, the analytical throughput using plasma-based techniques is considerably enhanced. Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

235

Michel Hoenig

Currently, substantial improvements to the utility of such instrumentation are continuously sought. However, in such situations where a dramatic lowering of detection limits is obtained, the risk of errors suddenly appearing due to sample handling is increased. Prior to commercial introduction of this modern instrumentation, these “new” errors were practically imperceptible to the determination of relatively high analyte concentrations that were measured with less sensitive techniques. The danger of contamination is now increasingly present: the choice of sample preparation procedure, the quality of its application and the need for an adequate laboratory environment have therefore become the most critical points deﬁning successful trace element determinations. Some basic notions that will afford a better understanding of the general philosophy regarding existing trace element analysis principles will be enumerated herein. First, it is necessary to understand that chemical analysis comprises a set of closely bound steps: for example, the choice of sample preparation procedure will depend on the measurement technique used and vice versa. It is then not sufﬁcient to simply intuitively apply a non-validated procedure (mineralization, dissolution, measurement technique, etc.) to a sample having an unknown composition. The set of analytical criteria has to be selected following a global consideration of the ﬁnal objective—reliable results in terms of accuracy and precision. The topic is quite vast and for this reason, in this chapter, trends relating to dry ashing methods, their principles, advantages and drawbacks, are discussed. It is hoped that the necessary ingredients needed to resolve usual cases will be treated here or in the references cited. It remains for the analyst to choose the appropriate preparation methodology in relation to the sample type, the available equipment and any imperatives of the analysis and, indeed, of the whole study. In most cases, preparation of solid samples involves several stages: drying (air, laboratory oven, etc.), homogenization (mixing, crushing, etc.), grinding (mills, mortars, etc.), followed by mineralization and dissolution of a subsample. The solution so obtained is ultimately diluted to volume. Ideally, the organic fraction of the sample has been decomposed and completely eliminated during these preparation steps and only dissolved inorganic compounds constitute the dissolved residue to be analysed. In the following sections, cases associated with the determination of total analyte content in the sample will be discussed. This generally means that quantitative dissolution of solid samples is required. To ensure this criterion is met for some difﬁcult types of solids, particularly those having a matrix containing silicate compounds (soils, sediments, plant material), the known procedures are often too labour- or time-consuming to be systematically applied in routine analyses. For environmental monitoring purposes, for example, they are often replaced by simpler and more easily applicable procedures. However, these substitutive methodologies rarely lead to an accurate determination of

236

Dry ashing

total analyte content, but they are generally sufﬁcient to satisfy the objectives of the study. Before the analysis, samples of organic or of a mixed nature are subject to two distinct steps, which often take place simultaneously: mineralization and dissolution. Samples of purely inorganic composition are simply dissolved. The composition of environmental and biological samples varies from purely inorganic (e.g. ﬂy ash) to purely organic (e.g. fats), but generally, they are an intermediate combination of these extremes. This implies that the total dissolution of samples usually cannot be achieved in a single step using a single reagent. In practice, the necessary number of steps and reagents is dictated by the matrix composition. Purely organic or mixed samples are usually brought into solution by some type of oxidation process combined with an acid dissolution of the resulting residue, as well as of the initial inorganic part of the matrix. In 1844, Fresenius and von Babo [1] published their method for the destruction of animal tissues prior to trace element determinations. In the intervening years, many procedures have been described for this purpose. However, despite numerous possible variations, almost all of the methods fall into one of two main classes, i.e. dry ashing and wet digestion. Dry ashing methods are especially appropriate for samples having a high organic matter content. The ﬁrst step of the method ensures the decomposition of organic matter by heating the sample to a relatively high temperature, with atmospheric oxygen serving as the oxidation agent. Chemical compounds (the so-called ashing aids) may sometimes be added to aid this process. The second step of a dry ashing method is the subsequent solubilization of the resultant ash using an appropriate acid or a mixture of acids. With wet digestions, the applied temperature is much lower, liquid conditions are maintained during the entire process and oxidizing agents in solution complete the oxidation. Generally, various combinations and proportions of strong acids with hydrogen peroxide ensure the decomposition of organic matter and the dissolution of the residue. Detailed aspects and the role of wet digestion procedures are discussed in Chapters 6 and 8. Depending on the sample type, the dissolution procedure generally involves several steps. Here, the terminology is precise: the term “mineralization” relates to samples having a totally or partly organic matrix only (animal and plant tissues, food samples, soils, etc.). Prior to the analysis, any organic compounds present must be decomposed and/or completely eliminated by the mineralization procedure. Using various reagents, the organic matter is decomposed into carbon dioxide, nitrogen oxides and water, thus liberating into solution all elements initially associated with it. After the mineralization procedure, the resulting sample residue should be essentially inorganic: it will be subject to a ﬁnal dissolution step similar to that used for a sample having an initially total inorganic composition (rocks, metals, etc.). For more complex samples (organic plus inorganic composition: soils, sludge, plant samples, etc.),

237

Michel Hoenig

chemical reagents and physical means are most often used to ensure these two roles (mineralization and dissolution) are simultaneously achieved. The objective of the sample preparation stage is usually to bring all available means into play in order to determine as readily as possible the elements of interest. First, these means have to convert the sample to a form that is compatible with the measurement technique utilized (generally a dissolution). Second, they should ensure the transformation and simpliﬁcation of the matrix (mineralization: wet digestion, dry ashing). Additionally, they may also perform the analyte separation or preconcentration (topics treated in subsequent sections). 7.2

WHY DRY ASHING?

Surprisingly, after the appearance of commercial advertising praising the universality and absolute necessity of wet digestion microwave heating devices for trace element analysis, several scientiﬁc papers have radically condemned dry ashing procedures, despite their long record of usefulness, e.g., “…from the general point of view, dry ashing decomposition is very problematic, especially in the determination of the trace content of heavy metals…” [2] or “…Dry ashing should be completely abandoned from the protocols of trace analysis. The experimental parameters are poorly reproducible and make dry ashing very error prone…” [3]. Moreover, “…as a result of risk of systematic errors related to dry ashing, the Measurement and Testing Programme (formerly BCR) of the European Commission has decided to withdraw mineralization procedures based on dry ashing from certiﬁcation of reference materials…” [4]. In contrast, many respected institutions, such as the AOAC International or the Nordic Committee on Food Analysis, as well as numerous other laboratories, have developed, veriﬁed, validated and successfully used classical dry ashing in practical analyses of a number of materials of biological origin. The same conclusions are perceivable in the very interesting and comprehensive study on the efﬁciency of dry ashing procedures for electrochemical methods published by Mader et al. [5]. Additionally, our own extensive experience in the ﬁeld of sample preparation shows that, when performed well, conventional dry ashing leads to complete removal of the organic matrix and to accurate analytical results for most usually determined elements. Better than the other known mineralization procedures, dry ashing methods ensure the quantitative decomposition and elimination of organic matter and an efﬁcient liberation of elements initially associated with it. Usually, these procedures are performed by calcination at atmospheric pressure in programmable mufﬂe furnaces. The commonly utilized temperature for this step is 4508C. In addition to conventionally heated mufﬂe furnaces generally employed for dry ashing purposes, the market now also provides microwave furnaces especially adapted to attain elevated temperatures. The

238

Dry ashing

unique advantage of the latter is the capacity to ensure application of very fast heating ramps. However, this interesting property is not applicable directly to usual dry ashing procedures, because heating ramps needed here have to be precisely slow. Additionally, a low temperature ashing (LTA) procedure in electronically excited oxygen plasma is available [6], very useful for sample preparation when volatile elements are to be determined. The instrumentation is, unfortunately, very expensive and not readily available at present. In addition, LTA is a particularly time-consuming procedure. In the usual high temperature ashing, fresh or dried (generally 103– 1058C) samples are weighed into suitable ashing vessels (vitreous silica, porcelain, platinum) and placed in the furnace. The temperature is then progressively elevated, following a convenient heating program, to attain 4508C, and then maintained for several hours. The resulting inorganic residue (ash) is dissolved using an appropriate acid. The resulting solution is transferred to a volumetric ﬂask, diluted to volume and analysed. Depending on the initial sample condition, results are expressed based on a fresh or dry weight basis. The application of dry ashing methods is simple and large series of samples may be treated at the same time. This is not their unique advantage—compared to wet digestions, dry ashing procedures present several other useful characteristics: †

†

†

The principal advantage of dry ashing procedures is the possibility of treating large sample amounts and dissolving the resulting ash in a small volume of acid (generally nitric or hydrochloric). This procedure permits preconcentration of trace elements in the ﬁnal solution, which is useful when very low analyte concentrations are to be determined. Such an advantage is not realizable with wet digestion methods. Additionally, heterogeneity is a typical property of many biological materials. The possibility of processing larger masses of sample, which, upon mineralization, provides a homogeneous solution, helps to minimize subsampling errors. The sample matrix is substantially simpliﬁed and the resulting ash is completely free of organic matter. This is a prerequisite for ensuring accuracy with some analytical techniques (e.g. ICP-MS, ICP-OES with ultrasonic nebulization, electrochemical methods) wherein analyte response may be inﬂuenced by the presence of residual carbon or some undigested organic molecules. The resulting solutions are of very acceptable aspect (clear, colourless and odourless), rarely the case when wet digestion methods are used and residual carbon content often attains elevated values. This is systematically observed not only for both conventionally heated and open microwave digestions (our experiments), but also for closed microwaveassisted digestions [7]. With high-pressure bombs, the residual carbon content is signiﬁcantly lower but it is never quantitatively eliminated as in the case of dry ashing (our experiments and Ref. [8]). Compared to wet digestion methods, reagent volumes and their handling are reduced in dry ashing procedures.

239

Michel Hoenig

†

The acidity of the ﬁnal solutions can efﬁciently be controlled: the acid is added directly to the ash and only a small fraction is consummated during its dissolution. With wet digestion procedures, added acids must also ensure the destruction of the organic matter and their effective amounts utilized during these chemical reactions vary quite signiﬁcantly. This results in unknown acid concentrations in the ﬁnal solutions to be analysed. This fact is at variance with well-known requirements for all methods based on atomic spectroscopy concerning the need for similarity of acidities between standards and samples. In some situations, this similarity is absolutely obligatory, e.g. for the determination of nickel by ICP-MS. The commonly used nickel cones in the ICP-MS interface always produce relatively high Ni backgrounds due to their ﬁnite dissolution by the aerosol being introduced. With variable acid concentrations, the background can vary signiﬁcantly from one sample to another, resulting in erratic Ni-results. Such unfavourable conditions may be avoided using dry ashing methods that ensure a practically constant acid concentration from sample to sample, allowing consistent ICP-MS determinations of nickel to be made.

Despite these several advantages, one must also accept several drawbacks of dry ashing procedures: the chemistry of the charring process is very complex, as one may have both oxidizing and reducing conditions varying throughout the sample and this may also occur as a function of time [9]. Moreover, during the combustion process, the actual temperature in the sample remains unknown; in some cases, it may be several hundred degrees above that of the furnace [10,11]. This may result in poor recoveries of some elements due to volatilization losses. A preliminary control of the methodology is then absolutely mandatory: the whole procedure must ﬁrst be validated using certiﬁed reference materials having composition similar to the samples to be analysed. However, the same remark applies to all other methodologies. 7.3

OXIDATION PROCESS AND DISSOLUTION OF THE RESIDUE

Even the experimentally well-established procedures based on conventional dry ashing largely lack an exact and objective process interpretation in physicochemical terms. Hence, a need exists for a greater elucidation of what can so far be considered as mostly empirically established procedures of classical dry ashing. Mader [12] studied how classical dry ashing actually proceeds in terms of dynamics and chemistry of organic matter degradation. Results not only revealed the strong exothermic character of decomposition during charring but also a possible variability, depending on the speciﬁc type of biological material. The former ﬁndings call for sufﬁcient moderation of the charring step (ramp heating) in order to prevent local overheating of the sample and subsequent risk of loss of a fraction of the analyte due to its mechanical removal in the form

240

Dry ashing

of solid particles of aerosol (smoke). Only under such controlled conditions, classical dry ashing can have the potential to yield accurate results. The term “dry oxidation” is generally applied to those procedures wherein organic matter is oxidized by reaction with gaseous oxygen, generally with the supply of energy in some form [13]. Included in this general term are methods in which the sample is heated to a relatively high temperature in an open vessel (conventional dry ashing), or in a stream of oxygen or air. In addition, related low-temperature techniques employing excited oxygen, bomb methods using oxygen under pressure and the classical oxygen ﬂask technique in which the sample is ignited in a closed system must also be included. All these methods involve two processes, although the relative signiﬁcance of each of them varies from one method to another, i.e., they ensure evaporation of the moisture and of volatile materials, including those produced by thermal cracking or partial oxidation, and ensure the progressive oxidation of the non-volatile residue until all organic matter is destroyed. Although these processes occur in all dry oxidations, it is not always possible to distinguish them as separate events. They are probably most easily separated in the conventional ashing procedure in which the organic material is heated in an open vessel with free access to air or oxygen. In usual analytical practice, the ﬁrst steps of such a procedure are usually conducted at a temperature much lower than that used to complete the oxidation. This is largely to prevent the ignition of the volatile and inﬂammable material produced by the process of destructive distillation and partial oxidation, as this would lead to an uncontrolled rise in the temperature resulting in an increased danger of analyte losses. The analysis of petroleum products presents an exception, because it is practically impossible to overcome their auto-ignition in a mufﬂe furnace, even using very low heating ramps. These samples are often purposely ignited in order to remove the bulk of the inﬂammable material before commencing the actual ashing procedure. Our experience has repeatedly shown that the most severe element losses are systematically observed during the heating ramp, not as a consequence of very high ﬁnal ashing temperature. An inadequate heating ramp may provoke the auto-ignition of the sample and the resulting rapid temperature increase results in volatilization losses. A preliminary low temperature treatment in the conventional process can be achieved in many ways: by heating gently over a ﬂame, on a heating plate, in a sand bath or with an overhead source of heat such as an infrared lamp. The most often utilized means of avoiding ignition problems is direct insertion of the sample into the mufﬂe furnace at low temperature, followed by heating with an appropriately slow ramp. Before the last stage of the process—progressive oxidation of the nonvolatile residue—the material remaining after the preliminary treatment is a more or less porous mass of charred organic matter containing variable amounts of inorganic material distributed throughout it. In reality, this picture is highly variable and it will depend not only on the type and the composition of

241

Michel Hoenig

samples analysed, but also on the action of possible reagents added that can change the initial chemistry of the process. Consequently, the kinetics of oxidation of such material will be dependent on the nature of the material itself, the inorganic substances it contains and its particle size and porosity. Tentative ﬁndings derived from such reactions with pure carbon or graphite can only be applied with caution to the complex chars existing in dry ashing of real samples. The temperatures generally recommended for dry ashing are, at about 5008C, low compared with those reported for the oxidation of graphite [14], but the chars produced are probably far more reactive due to unknown catalytic effects of the inorganic constituents present. The large amount of work published on the oxidation of material such as coal, coke and charcoal is probably more relevant to dry ashing discussed here, but the increased complexity makes the results more difﬁcult to interpret. In dry ashing procedures used for the analysis of environmental or biological samples (animal and plant tissues, food samples, blood, milk, etc.), the ﬁnal temperature is maintained for several hours. If the oxidation is achieved under optimal conditions, it leads to white or light grey coloured ashes, easily soluble in acids. Sometimes, depending on the sample type, the oxidation of organic matter is not completely achieved; in this case, the ash exhibits darker spots (dark grey to black) attributable to insufﬁciently oxidized carbon. Because this phenomenon is always responsible for a difﬁcult subsequent dissolution (often resulting in incomplete recoveries for several elements), such a residue must be re-treated using a few drops of nitric acid and brieﬂy recalcinated at the usual ashing temperature. After this treatment, ashes generally become clear and easily soluble. During the oxidation process, the analyte(s) will behave in one or more of a number of ways. Ideally, they will quantitatively remain in the residue (ash) arising from the oxidation, and in a form in which they can be readily recovered, generally by a simple dissolution of the ash in an appropriate acid. Fortunately, for the usefulness of the method, this is the case for most analytes and samples. In some cases, a part (or the total) of the analyte may be converted to a volatile form that may escape from the vessel (i.e. volatilization losses) or may be combined with the vessel surface or with some components of the inorganic residue remaining after oxidation (i.e. retention losses). In practical trace element analysis, the most often reported volatilization losses pertain mainly to mercury, arsenic and selenium. The analysis of mercury may be considered as a particular case and it will not be treated here: its high volatility implies the application of a preparation procedure exclusively based on wet digestion methods. Alternatively, with the help of a particular atomic absorption technique preceded by the combustion of the sample in a stream of oxygen and the subsequent amalgamation of mercury followed by its thermal release, this element can also be determined directly in the solid sample, thereby avoiding any mineralization problems (AMA-254, Altec, Czech Republic or DMA-80, Milestone, USA).

242

Dry ashing

On the other hand, retention losses result in poor recoveries of one or more analytes using the normal procedure for solubilization of the ash. They are generally observed for a particular quality of ashing vessel or in the presence of silicates or other insoluble compounds in the sample matrix. 7.3.1

Particular case of plant matrices

During intercomparison studies involving analyses of plant matrices, signiﬁcant discrepancies amongst results are often observed between laboratories using simple mineralization procedures and those that apply procedures that include a hydroﬂuoric acid attack followed by evaporation to dryness. In the former, the values obtained are systematically lower because complete digestion is not achieved. In the often utilized wet digestion procedures, mixtures of various acids with hydrogen peroxide may also lead to poor recoveries due to the presence of silicate compounds in the sample or to (co)precipitation phenomena [15]. As a consequence, Al, Fe, Cu and Mn, in particular, are not completely recovered, depending upon the speciﬁc plant matrix, probably related to the binding of analytes with the insoluble residue [16–18]. Mader et al. [19] and Quevauviller et al. [20] have noted similar problems with agricultural matrices such as composts, animal meats or brewers yeast. These statements, among others, are supported by an interesting study performed some years ago by Ledent et al. [21] wherein a variety of plants were charred at 4508C and the ash dissolved with hydrochloric acid. After ﬁltration, washing and drying, insoluble solid residues were analysed directly by optical emission-arc spectrography. Expressed as the sample dry weight, the Si content of the plants studied ranged between 0.3 and 10.3%, the highest being exhibited by a rice sample. The insoluble residue was also the highest for this sample (11.3%). Consequently, this means that 1% of other elements is retained in this insoluble residue. For several plants studied, analyses of insoluble residues have revealed that a signiﬁcant fraction of elements (major, minor, trace) is retained, varying between 0.06 and 95%, depending on the type of sample and the element studied. The most affected elements were Al (14–95% retained), Fe (7 –35%) and B (0.8– 23%). Consequently, we often use the recovery of aluminium as a marker for the procedure; if the Al recovery is incomplete, it may be concluded that the dissolution step was not performed under good conditions and that many other elements may be affected in a similar fashion. The same work [21] has shown that, in some cases, Na, an element generally considered to be easily solubilized, may also not be recovered in solution, often with more than 20% lost. Retained percentages of Ca, Mg, Cu and Mn were lower in all cases, ranging between 0.1 and 4%. For trace elements, the limited sensitivity of the technique used did not provide results

243

Michel Hoenig

with good precision, but it was clear that the retention of some elements in the ash, such as Pb, Ni, Cr, V and Ti, may occur. A further detailed study focussing on retention phenomena relating to dry ashing of plant samples has been performed by Hoenig and Vanderstappen [22]. Here also, losses of trace elements induced by incorporation into the insoluble residue were by far the most important: after a simple dissolution of ash by nitric acid losses could reach 20% for Pb, 15% for Cd, 10% for Cu, 8% for Zn and 5% for Mn. Such problems highlight the paramount need for a more elaborate dissolution procedure for plant ashes. For this reason, we have repeatedly claimed the absolute necessity of utilizing an HF-assisted dissolution of the ash if the objective is the determination of total element content. This problem, typically associated with plant samples, is similar when applying a wet digestion procedure: if an insoluble residue remains, an additional HF step, followed by evaporation to dryness, must also be performed. Another example highlighting this problem was reported by Dams et al. [23], who demonstrated that ICP-MS interferences from residual silicon resulted in up to 30% positive bias in intensities from 63Cu, 65Cu and 55Mn derived from soil and sewage sludge digests, due to spectral interferences from 28 27 Si Alþ, 28Si35Clþ and 28Si37Clþ. Addition of an HF step to the procedure followed by evaporation to dryness removed the interfering Si and permitted accurate analysis to be obtained. Such a procedure is not always easy to achieve with most commercially available microwave heating (closed) devices and is, in any case, much more difﬁcult to apply than with a dry ashing procedure. Finally, plants are often considered as purely organic samples with some trace elements present. It is clear that the aforementioned problem is comparable to that encountered in soil, sediment, sludge and rock samples where silicon is typically the primary matrix element. In this case, however, all analysts are aware of the absolute necessity of dissolving the entire sample if the total analyte content has to be determined. 7.4 7.4.1

METHODOLOGY Heating devices

Dry ashing methods can be applied to mineralization of organic materials, biological tissues, plant and food samples, sludge, etc. Well mastered, they ensure total destruction of the organic matter; the associated elements are generally transformed to carbonate or oxide forms. At present, they are generally performed using fully programmable (ramp and holding times/temperatures) mufﬂe furnaces equipped with an efﬁcient temperature control and reproducible thermal programs. Required intermediate evaporations to dryness are usually achieved on sand baths or on hot plates.

244

Dry ashing

7.4.2

Ashing vessels

It is mandatory to select an ashing temperature that ensures quantitative decomposition of the organic matter without partial or total loss of analytes by volatilization or by their incorporation into a residue that is insoluble in usual reagents. The latter may result from formation of refractory oxides, from combinations with other sample constituents present, as well as from reactions with the walls of the crucible. As noted earlier, one of the causes of losses during dry ashing procedures is the reaction of the analyte with some of the solid matter present in the system. In order for a reaction of this nature to constitute a problem, it is ﬁrst necessary that it occurs to a signiﬁcant extent and, second, that the product of the reaction be insoluble in the reagents generally used for dissolving the resultant ash. The solid matter available for such a reaction is generally the material of the ashing vessel and the residue from the sample itself. It is obvious that their nature will have a considerable effect on the extent of the losses. The most commonly used ashing vessels are made of silica or porcelain, with platinum as a third alternative. Vitreous silica is a glass consisting almost entirely of SiO2 (^99.8%), with some Na, Al, Fe, Mg and Ti oxide impurities, whereas the glaze on porcelain ware is a more complex material containing Al, K, Ca and Na oxides (up to 30%) in addition to silica. For both vitreous silica and porcelain, the obvious reaction is between the oxide of the analyte and the ashing vessel to produce a complex silicate, resulting in a loss. Studies with radiotracers performed by Gorsuch [24] have shown that retention of metals by reaction vessels made of vitreous silica may be very signiﬁcant during dry ashing. The most affected element is silver (47%), followed by gold (17%), copper (4%), lead and cobalt (0%). This type of reaction clearly occurs, but it is dependent on many factors. Some oxides react much more readily than others and, even if silicates are formed, some will be stable to subsequent acid attack while others will readily be decomposed and so cannot be considered to cause losses. These reactions will, of course, be exacerbated if the ashing vessel is made of silica or porcelain, exhibits a marked weakening of the silicate structure, or a worn surface because of extensive use. Because the extent of such reactions remains unknown, the alternative practice of using essentially inert platinum crucibles is much more reliable. This metal is virtually unaffected by any of the usual acids, including hydroﬂuoric. Platinum is attacked only by concentrated phosphoric acid at very high temperatures and it dissolves readily in mixtures of hydrochloric and nitric acids (or of nitric acid with other chlorides). Consequently, platinum vessels cannot be used for aqua regia digestion procedures. Of course, the initial cost for platinum is signiﬁcantly higher than for other types of ashing vessel, but its lifetime is practically unlimited. Moreover, the total cost of a modern programmable mufﬂe furnace equipped with a set of

245

Michel Hoenig

platinum crucibles is, in any case, much less than the cost of commercial microwave-assisted wet digestion systems. 7.4.3

Inﬂuence of the sample composition

An additional retention problem encountered with dry ashing procedures is that posed by the sequestering action of some materials produced during ignition. The binding of iron by condensed phosphates produced by the action of heat on simple phosphates [13] or retention of several elements on silicate compounds present in the sample [25] is the best-known example. Munter et al. [26] reported Al, Fe and Cr losses as a result of their retention on the ash residue, and Cu through its retention on the silica crucible, after ashing at 450– 5008C. Similarly, Dahlquist and Knoll [27] found losses of Fe, Ni, V, Al and Cr due to their retention on insoluble silica residues, although these were not observed when nitric acid was used as an ashing aid. Koirtyohann and Hopkins [28] demonstrated that the losses of Cd, Cr, Fe and Zn from animal tissue samples ashed at temperatures below 6008C were also attributable to retention on the insoluble residue. The often contradictory observations described in all these, and other studies, illustrate the complexity of the retention problems and the need for an adequate dissolution step. On the other hand, with the exception of arsenic and selenium, these considerations indicate that several losses reported as being due to volatilization are, in reality, due to retention problems. Dry ashing temperatures commonly used for trace element analysis range between 450 and 5008C. They are generally high enough to ensure a complete oxidation of the organic matrix while avoiding the dangers of volatilization losses for most analytes (except Hg, As and Se). Losses are further minimized if the ashing temperature is attained using a slow ramp (8 h from ambient temperature) that prevents any local hot spots or self-ignition of the sample from developing. Volatilization losses can also result from overshoot of the maximum temperature set, hence the importance of efﬁcient temperature control on the heating device. 7.4.4

Operating modes for environmental samples

The objectives of dry ashing methodology are very simple and may be summarized as follows: an efﬁcient decomposition of organic matter without losses of elements of interest, and a subsequent easy dissolution of the residue. For most samples and analytes studied, the application of a conventional dry ashing method satisﬁes these requirements and numerous alternatives are then not necessary. From the literature cited, and from our own experience, it may be stated that a unique procedure also permits good recoveries for most major (Na, K, Ca, Mg, P), minor (Al, Fe, Mn) and trace (B, Be, Ba, Cd, Co, Cr,

246

Dry ashing

Cu, Ni, Mo, Pb, Sb, Tl, V, Zn) elements in environmental, biological and food samples. Of course, the above-mentioned problem associated with arsenic and selenium remains. Dry ashing procedures are apparently slow, sometimes requiring up to 24 h per sample, but they have the advantage of being readily adapted to a large batch processing and do not require constant operator attention. The operating modes for two of the most often utilized dry ashing procedures are presented below: Materials: platinum crucibles, programmable mufﬂe furnace, sand bath or hot plate, demineralized water, concentrated nitric and hydroﬂuoric acids of adequate purity, appropriate micropipettes, calibrated ﬂasks, storage bottles. Procedure 1: dry ashing method for mineralization of animal tissues, milk, fruit juices, blood, faeces, urine, etc. (and all samples of organic nature without silicates in the matrix). If well applied, this procedure is safe and gives very accurate and reproducible results for major, minor and trace elements (As and Se excepted) [29]. If the % moisture is required, its determination may be performed using a separate vessel: after weighing of the raw sample, it is dried for 16 h at 1038C, cooled in a desiccator and weighed again. † † †

† †

Following the measurement technique used, weigh in a platinum crucible 0.2 –2.0 g dried sample (1058C) or up to 10 g fresh sample. Place the crucible into a cold mufﬂe furnace and progressively elevate the temperature to attain 4508C in 8 h. Maintain this temperature for 5 h, cool. If the resulting ash is not of white or light grey colour, add 0.5 ml demineralized water and 0.1 –0.2 ml nitric acid to the residue. Evaporate to dryness on a sand bath or a hot plate, place the crucible again into the mufﬂe furnace, elevate the temperature to attain 4508C in 2 h and maintain it for 1 h. After this treatment, ashes generally become clear and easily solubilized. Dissolve the cool residue with 1 ml nitric acid and 20 ml demineralized water and heat to a gentle boil. After cooling, quantitatively transfer the solution to a 50 or 100 ml calibrated ﬂask and dilute to volume with demineralized water.

Procedure 2: modiﬁed dry ashing method of the Comite´ Inter-Instituts d’Etude des Techniques Analytiques (CII) for mineralization of plant samples with removal of silica (may possibly be used for soils, sludges and sediments having high organic content). This reference dry ashing method dedicated to plant tissues was tested and largely validated during the long existence of the CII group [30–37]. If well performed, this dry ashing method is safe and gives very accurate and reproducible results for major, minor and trace elements in all plant and similar samples (As and Se excepted). If the % moisture is required, perform its determination using a separate crucible, as noted earlier.

247

Michel Hoenig

† † †

† † †

Following the measurement technique used, weigh in a platinum crucible 0.2 –2 g dried sample (1038C). Place the crucible into a cold mufﬂe furnace and progressively elevate the temperature to attain 4508C in 8 h. Maintain this temperature for 5 h, cool. Following the initial sample weighing, moisten the residue with 0.5 –2.0 ml demineralized water, add 1.0 – 3.0 ml concentrated nitric acid and 0.5 –2.0 ml concentrated hydroﬂuoric acid. Evaporate slowly to dryness on a sand bath or on a hot plate. Repeat the last step two times. Dissolve the cool residue with 1 ml nitric acid and 20 ml demineralized water and heat to a gentle boil. After cooling, quantitatively transfer the solution to a 50 or 100 ml calibrated ﬂask and dilute to volume with demineralized water.

As reported in a previous study [37], amongst all the procedures known for the mineralization of plant samples, only a method able to volatilize silicon allows one to avoid problems due to retention losses of trace elements by insoluble silica residues. Only methods that include hydroﬂuoric acid treatment will ensure the removal of silicon and the mobilization of analytes into solution. Solutions obtained using this dry ashing method permit all major, minor and trace elements usually studied (As and Se excepted) to be easily determined by all atomic spectrometric techniques (F-AAS, ET-AAS, ICP-OES, ICP-MS). With the objective being multielement routine analysis, it would be highly desirable to also determine arsenic and selenium in these same solutions. In addition, because of the very low As and Se contents in most plant samples, the effective preconcentration that can be achieved during dry ashing is also welcome. Unfortunately, this alternative initially seems to be unrealizable because of possible As and Se volatilization losses. Nevertheless, recent studies indicate that volatilization losses of arsenic and selenium during the ashing of plant material can be avoided [38]. 7.5

PARTICULAR CASES OF ARSENIC AND SELENIUM

It is likely that the major analytical concern has been with the determination of these two volatile elements occurring in low concentrations in environmental materials. As might be expected, virtually all the usual oxidation procedures have been applied to ensure their recovery, with a pronounced preference for wet oxidation methods. The number of references to methods for decomposition of organic samples in the literature is immense, practically impossible to summarize. Many comparisons of various methods have been conducted and, despite inevitable contradictions, some conclusions can be drawn. Sample preparation for the determination of arsenic and selenium is generally performed using wet digestions that are, from the outset, less likely

248

Dry ashing

to suffer from volatilization losses. Nevertheless, more than for other analytes, and independent of the preparation procedure adopted, there are other severe problems that can arise during the analysis of these elements. These are due to a number of unfavourable analytical factors, ranging from insufﬁcient instrumental sensitivity (ICP-OES with low nebulization efﬁciency, ET-AAS with energetically poor primary sources) and/or low analyte levels (milk, plant and food samples), to more pronounced spectral interferences (ET-AAS with a structured background at the As and Se analytical wavelengths, ICP-MS with the unique As isotope and possible 75As/40Ar35Cl interference) and unusually severe matrix effects (ET-AAS with strongly decreasing slopes of working curves and HG-AAS with changing hydride generation kinetics in the presence of some matrices). Such conditions sometimes necessitate such extreme dilution factors that they are incompatible with the initial analyte content needed to permit ﬁnal determinations (ET-AAS analysis of urine and milk). Precisely uncontrolled matrix effects generally lead to signal suppressions resulting in low recovered values. However, volatilization losses (preparation steps), preatomization losses (ET-AAS) or As losses in the condensation stage of ultrasonic nebulizers (USN-ICP-OES) also result in low recovered values. In such situations, the weak link in the analytical chain remains unknown, particularly for inexperienced analysts. This clearly indicates that not only preparation steps, but measurement steps also have to be carefully controlled. In other words, it can be said that the entire methodology used for arsenic and selenium remains one of the most arguable points encountered in environmental trace element analysis. Turning ﬁrst to a consideration of methods used for dry ashing, a number of recovery experiments have been reported, and quite signiﬁcant controversy amongst the results is to be seen. Despite the great volume of work devoted to investigating the recovery of arsenic during dry ashing, it is apparent that much remains to be done. Studies such as that of Hamilton et al. [39], in which a radioactive tracer is fully incorporated into the sample, need to be extended, with both animals and plants being used, and with the effect of the many possible ashing aids being investigated. One point appears clear, arsenic recoveries from plant samples are signiﬁcantly higher than those from samples of animal origin, such as blood, bone or kidney [24,39,40]. Similar recovery studies have also been performed with selenium; e.g., Reamer and Veillon [41] have proposed both wet and dry oxidation methods, which lead to complete selenium recoveries. Unfortunately, these claims were based on experiments with biological materials containing added selenium only and, consequently, were not representative of real forms of selenium likely to be present in the samples. The suitability of dry oxidation methods for recovery of arsenic is, at present, somewhat uncertain. Charring with magnesium nitrate as an ashing aid appears to be the most satisfactory of the many dry procedures described, but more rigorous testing is required, using a wide variety of different samples.

249

Michel Hoenig

In the discussion concerning the recovery of arsenic from biological material, the nearly 60-year-old work of Satterlee and Blodgett [42] is of considerable signiﬁcance. They found a thermolabile arsenic-containing fraction in blood and tissues that was lost on drying at temperatures as low as about 608C. If such substances are really of common occurrence, the use of practically all available dry and wet oxidation methods of sample decomposition is essentially forbidden for preparation of such materials. 7.5.1

Ashing aids

In its initial form, a dry ashing procedure cannot be considered appropriate for preparation of samples to be used for the determination of As and Se. However, many of the reported ashing methods describe the addition of some extra inorganic compounds to the sample to improve the efﬁciency of the procedure. These added materials are generally called ashing aids, and they serve one or both of two purposes, i.e., to facilitate the decomposition of the organic matter, or to improve the recovery of the element to be determined. The most common aid, used to purely hasten the oxidation of organic material, is nitric acid. It is generally added towards the end of the ashing process to decompose small amounts of remaining carbonaceous material (see Section 2.2.4.4). Because the ash from most biological materials contains up to several tens of per cent carbonates, nitrates are formed in situ after the addition of nitric acid. Additional ashing is then, in fact, melting with nitrates and should help to remove the most resistant degradation products present in the organic matrix. This step, leading to the production of a clean ash, has to be performed with care because when appreciable amounts of organic material are still present, it can cause the ignition of the residue when it is returned to the furnace, resulting in a possible loss of material. Some substances serve as auxiliary oxidants as well as serving other purposes. These are commonly the nitrates of light metals such as magnesium, calcium or aluminium, which decompose on heating to yield oxides of nitrogen. Of all the ashing aids noted, magnesium nitrate is probably the most widely used. According to Gorsuch [13], these auxiliary oxidants also fulﬁl the important function of being an inactive diluent in the process. As the organic matter in a sample is progressively decomposed, the analytes are brought into closer contact with the material of the vessel and other constituents of the residue. If a reaction with them is feasible, then the increased proximity will increase the chance of its occurrence. Under these circumstances, dilution of the ash with an inert material, such as magnesium oxide, should greatly reduce the possibility of undesirable solid state reactions, resulting in improvement of recoveries. The well-known utilization of relatively unstable magnesium nitrate as an ashing aid likely offers both the advantage of more rapid oxidation and of decreased retention losses.

250

Dry ashing

These oxidative-dilution agents improve recoveries without entering into any reaction with the sample itself. Another group of ashing aids achieves the same end by altering the chemical nature of some of the constituents. The best example of this is the use of sulphuric acid to convert volatile chlorides to involatile sulphates; this may prevent losses of Cd, Pb or Cu at ash temperatures up to 7508C [43]. Arsenic and Se determinations can, in some cases and under particular conditions, also beneﬁt from the advantages offered by a dry ashing procedure. The addition of ashing aids—generally MgO and/or MgNO3—can give rise to less volatile As or Se compounds during the ashing procedure. The successful use of ashing aids is, of course, strongly dependent on the initial form of the analyte. In any case, utilization of ashing aids is a particularly delicate step because some successful examples cannot lead to generalizations: for routine use, the procedure necessitates a serious and time-consuming validation for each type of sample analysed. In addition, the utilization of ashing aids signiﬁcantly increases the total dissolved solids content of solutions and enhances the dangers of contamination, limiting the use of this approach for ICP-MS analysis. General recommendations for the use of ashing aids may be found in the appropriate Analytical Methods Committee report [44]. 7.5.2

What to do?

Using radiotracers, Gorsuch [39] reported recoveries of As of about 90% for a cocoa sample following a dry ashing at 5508C (99% with Mg nitrate). Similarly, Hoenig et al. [38] have shown that a dry ashing procedure (4508C) applied without ashing aids to plants of terrestrial origin (leaves, grasses, etc.) provided consistent recoveries for As and Se but this was not the case for plants of aquatic origin (algae). Arsenic recoveries from blood samples treated without ashing aids at various temperatures are widely divergent and may vary between 0% (500 and 7008C ashing temperature) and 23% (4008C) [40] or between 57% (8508C) and 72% (4508C) [39]. This clearly shows that the form of the analyte initially present in the sample (organic, inorganic, oxidized, etc.), as well as the particularly experimental conditions used, plays, more than for other analytes, roles of paramount importance during the analysis of arsenic. It appears that As in terrestrial plants is present in an inorganic form (e.g. oxides, probably coming from dust emission) that is not volatilized during dry ashing. In plants of aquatic origin, it is perhaps represented by more volatile organic species formed after its assimilation directly from water. Despite these suppositions, the complete recovery of arsenic from terrestrial plants cannot be explained because arsenic oxides possess melting/boiling points that are lower than the applied ashing temperature of 4508C. Amongst other possibly more refractory compounds, only arsenic sulphide, having a boiling point higher than 7008C, might be considered, but

251

Michel Hoenig

in this case a reaction is difﬁcult to formulate. Because a plant matrix may contain up to 5% K and Ca and up to 0.5% Mg, Na and P, more probable is the possible participation of these elements or their compounds, which might act as “natural” ashing aids during the process. But, here again, an unwelcome question arises: why is arsenic lost during the dry ashing of aquatic plants and animal tissues, where the matrix is of similar composition to the matrix of terrestrial plants? For animal tissues, the only real difference is the absence of silicates (that might ensure possible retention of As and then prevent its volatilization) but in aquatic plants silicates are also present at signiﬁcant concentration levels (e.g. IAEA-0390 Algae). Currently, there is no consistent explanation for the good arsenic recoveries obtained during dry ashing of terrestrial plants. An explanation of the characteristics of selenium is equally difﬁcult. This element closely resembles sulphur in many ways, and forms an extensive number of organic compounds. For trace element analysis, as in the case of arsenic, wet oxidation procedures have found much wider acceptance than dry ashing methods, due to the readiness with which the analyte may be lost during ignition. Oelschla¨ger [45] described the use of a dry ashing procedure in the presence of magnesium nitrate after a preliminary treatment with nitric acid; it is possible that the use of this ashing aid is adequate to prevent undesired reducing conditions or to permit other chemical reactions. For example, the presence of undigested carbon in the sample may lead to losses of As and Se as volatile carbides. The role of magnesium nitrate is to further oxidize any incompletely digested components of the organic matrix during dry ashing treatment by elimination of reactive carbon and to favour a possible mechanism that prevents the production of such carbides [46,47]. It must be acknowledged that numerous laboratories currently successfully apply various alternatives of this methodology. In an unpublished study, we have tested an additional approach for possible application of dry ashing for the determination of As. In order to modify initial forms of As, or the form of the sample matrix, or both, various plant and animal tissue reference samples were pretreated using a wet digestion (nitric acid plus hydrogen peroxide under reﬂux), followed by evaporation to dryness before the application of a classical dry ashing procedure. This approach should result in an original procedure that might combine the advantages of both wet and dry methods utilized in laboratory practice. The preliminary wet oxidation treatment modiﬁes the matrix and removes a large fraction of the organic matter initially present. Such pretreatment avoids the often reported dry ashing losses due to self-ignition of the sample during the ashing ramp, but probably also provides a matrix modiﬁcation that results in a more efﬁcient retention of As in the residue. With this relatively time-consuming preparation methodology, As recoveries increased up to about 60% (in comparison with the <30% obtained with classical dry ashing procedure). Unfortunately, complete

252

Dry ashing

As and Se recoveries have never been achieved, but this approach deserves to be further studied. All considerations summarized here indicate clearly that, with our present knowledge, preparation steps for As and Se determinations should be undertaken exclusively using wet digestion procedures that remain, despite their several disadvantages, more reliable in terms of minimization of volatilization losses of these analytes. 7.6

CONCLUSIONS

Presently, the instrumentation market offers devices primarily for making wet digestions more attractive and easier to manage, mainly by means of automation. This relates principally to microwave systems, where the energy transfer from reactive mixture to the sample is claimed to be better than in the case of conventional means of heating. Concerning sample mineralization steps for trace element analysis, the scientiﬁc and commercial literature has systematically claimed the superiority of microwave-assisted wet digestion methods for the last decade. It is, consequently, understandable why their use is so predominant in analytical laboratories. Despite high purchase prices and operating costs, the considerable sales of microwave heating devices are the best testimony to this general trend. However, as often observed, the claimed superiority of microwaves techniques is not signiﬁcantly evident in the digestion efﬁciency of typical environmental samples. Nevertheless, in some other cases, microwave heating exhibits particularly good performance: the best-known example concerns the mineralization of plastic materials where long carbonaceous chains must be destroyed. Compared to microwave systems, it must be admitted that the use of very high pressure acid digestion bombs heated by conventional means (e.g. Parr 4746, heated in an ordinary laboratory oven, operating at temperatures up to 2858C with working pressures of about 350 bar) may lead to better mineralization efﬁciency for some difﬁcult samples (e.g. those with a high content of fats such as milk, butter, etc.). Moreover, the purchase cost of such a device having a sufﬁcient number of bombs is signiﬁcantly lower than a modern microwave system. In addition, the lack of a generalized methodology for wet digestions, with or without microwave assistance, clearly indicates that a consensus has not been reached in this domain. On the other hand, due to their simplicity of application, dry ashing procedures in open systems have been clearly established for a long time and are well validated. Unfortunately, all the advantages afforded by this routinely used methodology are not often reported in the literature. The recent disinterest in dry ashing procedures is surely due to often reported dangers of volatilization losses, in addition to a generally weak interest in “old” and apparently not sufﬁciently “elegant” methodologies. The relatively poor advertising of modern dry ashing systems

253

Michel Hoenig

by manufacturers is also responsible for this situation. The danger of volatilization loss is, of course, more likely to occur than with wet digestion methods. Nevertheless, if the ashing process is well controlled, such losses may plague only three popular elements of environmental interest, viz., As, Se and Hg. For all other major, minor and trace elements, a dry ashing procedure remains a unique alternative, ensuring the best simpliﬁcation of the matrix through the total removal of organic matter, a prerequisite for several analytical techniques, including ICP-MS, ICP-OES with ultrasonic nebulization or electrochemical methods.

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12

13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28

254

R. Fresenius and L. von Babo, Ann. Chim. Pharm., 49 (1844) 287. J. Pikhart, Chem. Listy, 82 (1988) 881. P.B. Stockwell and G. Knapp, Int. Labmate, 14 (1989) 47. Ph. Quevauviller and E.A. Maier, Report EUR, European Commission, 1600 EN, 1994. P. Mader, J. Sza´kova´ and E. Curdova´, Talanta, 43 (1998) 521. C.E. Gleit and W.D. Holland, Anal. Chem., 34 (1962) 1454. L.M. Costa, F.V. Silva, S.T. Gouveia, A.R.A. Noguieira and J.A. Nobrega, Spectrochim. Acta B, 56 (2001) 1981. S. Strenger and A.V. Hirner, Fresenius J. Anal. Chem., 371 (2001) 831. A. Bock, A Handbook of Decomposition Methods in Analytical Chemistry. International Textbook Company Ltd., London, 1979, pp. 124 –130. W. Oelschla¨ger, Z. Anal. Chem., 246 (1969) 376. B. Bopel, Z. Anal. Chem., 266 (1973) 257. P. Mader, in: J. Cibulka (Ed.), Movement of Lead, Cadmium and Mercury in Agricultural Production and in Biosphere. State Agricultural Publishing House SZN, Prague, 1986, pp. 117– 156. T.T. Gorsuch, The Destruction of Organic Matter. Pergamon Press, Oxford, 1970. K. Kinoshita, Carbon—Electrochemical and Physicochemical Properties. Wiley, New York, 1988. E.J.M. Temminghoff and I. Novozamsky, Analyst, 117 (1992) 23. C.Y.L. Huang and E.E. Schulte, Commun. Soil. Sci. Plant Anal., 16 (1985) 943. Y.P. Kalra, D.G. Maynard and F.G. Radford, Can. J. For. Res., 19 (1989) 981. J.L. Havlin and P.N. Soltanpour, Commun. Soil. Sci. Plant Anal., 11 (1980) 969. P. Mader, J. Szakova and J. Kucera, Biol. Trace Elem. Res., 43–45 (1994) 633. Ph. Quevauviller, J.L. Imbert and M. Olle´, Microchim. Acta, 111 (1993) 147. G. Ledent, R. De Borger and S. Vanhentenrijk, Analusis, 12 (1984) 393. M. Hoenig and R. Vanderstappen, Analusis, 6 (1978) 312. R.F.J. Dams, J. Goossens and L. Moens, Microchim. Acta, 119 (1995) 277. T.T. Gorsuch, Analyst, 84 (1959) 135. M. Hoenig, Talanta, 54 (2001) 1021. R.C. Munter, R.A. Grande and P.C. Ahn, ICP Information Newslett., 5 (1979) 368. R.L. Dahlquist and J.W. Knoll, Appl. Spectrosc., 32 (1978) 1. S.R. Koirtyohann and C.A. Hopkins, Analyst, 101 (1976) 870.

Dry ashing 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47

M. Pinta, Spectrome´trie d’Absorption Atomique: Applications a` l’Analyse Chimique (Part II: Analytical Applications). Masson, Paris, 1979, in French. M. Pinta et le CII, Analusis, 3 (1975) 345 in French. M. Pinta, Oleagineux, 28 (1973) 87 in French. R.C. Daniel, G. Theiller and the CII members, Fresenius J. Anal. Chem., 345 (1993) 230. M. Pinta and the CII members, Geostand. Newsl., 1 (1977) 25. M. Pinta and the CII members, Geostand. Newsl., 1 (1977) 115. R.C. Daniel and the CII members, in: P. Martin-Prevel (Ed.), Proc. 6th Int. Coll. for the Optimisation of Plant Nutrition, Montpellier (F), 1984, Vol. 3, p. 837. R.C. Daniel, P. Lischer, D. Ruf and G. Theiller, Recherche Agronomique en Suisse, 31 (1991) 117. M. Hoenig, H. Baeten, S. Van Hentenrijk, E. Vassileva and Ph. Quevauviller, Anal. Chim. Acta, 358 (1998) 85. E. Vassileva, H. Docˇekalova´, H. Baeten, S. Vanhentenrijk and M. Hoenig, Talanta, 54 (2001) 187. E.L. Hamilton, M.J. Minski and J.J. Cleary, Analyst, 92 (1967) 257. J. Pijck, J. Hoste and J. Gillis, Int. Symp. Microchem., Birmingham, 1958. D.C. Reamer and C. Veillon, Anal. Chem., 53 (1981) 1192. H.S. Satterlee and G. Blodgett, Ind. Eng. Chem., 16 (1944) 400. M. Feinberg and C. Ducauze, Anal. Chem., 52 (1980) 207. Analytical Methods Committee, Analyst, 85 (1960) 643. W. Oelschla¨ger, Landw. Forsch., 18 (1965) 79. M. Stoeppler, K.P. Muller and Z. Backhaus, Z. Anal. Chem., 297 (1979) 107. D.L. Styris, L.J. Rell and D.A. Redﬁeld, Anal. Chem., 63 (1991) 503.

255

This page intentionally left blank

Chapter 8

Microwave-based extractionq Edward E. King and David Barclay

8.1

INTRODUCTION

This chapter explores microwave-based extraction, which includes dissolution, destruction, and leaching of analytes for the preparation of various sample media for elemental chemical analysis. After a brief introduction exploring the history and theoretical overview of microwave technology, various system types will be described with a list of application notes giving a readers reference guide to microwave sample extraction. 8.2

BRIEF HISTORY OF INDUSTRIAL MICROWAVE DEVICES

Commercial microwave extraction devices have been available since the late 1970s and are now commonplace in today’s modern laboratory. Microwave technology has beneﬁted greatly from past experimentalists, whose pioneering efforts have turned this highly complicated and technical discipline from an art form into a science. The turning point in the practical application of microwave energy came from researchers at the University of Birmingham during the Second World War. Randall and Booth designed the magnetron, a reliable device for generating ﬁxed frequency microwaves. Further investigation showed that microwaves could increase the internal temperature of foods much quicker than a conventional oven. Percy LeBaron Spencer of the Raytheon Company accidentally discovered that microwave energy could cook food when a candy bar in his pocket melted while he was experimenting with radar waves. This work ultimately led to the introduction of the ﬁrst commercial microwave oven for home use as a safe, cost-effective heating method [1]. q The authors dedicate this chapter to the memory of Russ Saville of the Savillex Corporation. Russ was well known for his contribution to Teﬂon vessel and vessel related products. He was instrumental in the growth and success of microwave pressure vessel technology over the past 15 years, which led to the successful adaptation of this technology to the modern analytical laboratory. Russ was a wonderful person and a great colleague—he will be missed.

Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

257

E.E. King and D. Barclay

With the advancement of commercial microwave ovens came the ﬁrst microwave system for the laboratory. Initially, home microwave ovens were modiﬁed for speciﬁc chemical heating applications. These systems were used to rapidly heat open vessel reactions for accelerated microwave extraction. The observations made during these test reactions showed that microwave energy has special properties for heating laboratory substances. These systems were inexpensive and not designed to withstand a rigorous laboratory environment; however, they provided a basic understanding of the effectiveness of this technique for rapid heating. In the 1980s, companies began to manufacture industrial microwave ovens speciﬁcally designed for use in laboratories. These systems incorporated Teﬂonw-lined, stainless steel cavities with safety systems, such as interlocking multi-layered doors, that could handle a laboratory environment and programmable control systems that could apply microwave energy dependent upon the actual chemistries of the sample. The application of microwave power was tailored to heat multiple samples evenly. These safety and control features distinguished the laboratory microwave systems from consumer microwave products. The next breakthrough occurred with the use of closed vessel reactors in the microwave. The beneﬁts of closed vessel, high-pressure reactors were well known as devices that easily raised the reaction temperature by providing a pressurized reaction environment. The challenge was to develop a closed reactor system, or vessel, incorporating materials that were compatible with both the microwave and chemical environments, as well as being safe for the user. Microwave vessel technology has improved greatly over the last 20 years. At the dawn of its existence, the microwave pressure vessel was limited to a maximum pressure of 100 psi (6.8 bar) and a maximum temperature of 1208C. Above these conditions, the vessel leaked or failed to hold and maintain a constant pressure. The advancement of modern thermoplastics and a better understanding of the distribution of the forces within and around the pressure vessel have advanced the limits of microwave extraction vessels to 1500 psi (100 bar) and temperatures as high as 3008C. These higher pressure and temperature conditions enable extractions of samples that were not possible in atmospheric or lower condition systems. Today, with modern industrial microwave systems and advanced vessel technology, laboratory chemists have safe, effective tools to aid in rapid sample extraction methods. 8.3

MICROWAVE THEORY

Microwaves are a powerful, reliable energy source that may be adapted to many applications. The following section will give a brief description of microwave theory and microwave energy transfer and how it is applied to the heating of samples for microwave extraction.

258

Microwave based extraction

Fig. 8.1. Electromagnetic spectrum.

A microwave is a form of electromagnetic energy that falls at the lower frequency end of the electromagnetic spectrum, and is deﬁned in the 300– 300,000 MHz frequency range (Fig. 8.1). Only four microwave frequencies are available commercially for use in industrial, scientiﬁc, or medical applications: 900; 2450; 5800; and 22,125 MHz. For home and laboratory systems, 2450 MHz is the most commonly used frequency. The microwaves at this frequency have a penetration depth appropriate (, 25 mm) to interact with laboratory samples. Also, due to mass production of consumer microwave ovens, cost-effective magnetron sources are available that can deliver adequate power for laboratory systems [2]. Microwave energy consists of an electric ﬁeld and a magnetic ﬁeld, though only the electric ﬁeld transfers energy to heat a substance (Fig. 8.2). Magnetic ﬁeld interactions are not a signiﬁcant factor in microwave extraction heating. Microwaves move at the speed of light (300,000 km/s). The quantum energy of microwaves (0.037 kcal/mol) is very low relative to the typical energy required to cleave molecular bonds (80–120 kcal/mol); thus, microwaves will not affect the structure of a molecule. The extraction or destruction of chemical bonds with microwaves is achieved entirely through kinetic energy transfer. Traditionally, laboratory samples were heated through conductive heat transfer using an external heat source (Fig. 8.3A). Heat is driven into the substance, passing ﬁrst through the walls of the vessel in order to reach the sample.

259

E.E. King and D. Barclay

Fig. 8.2. Electromagnetic radiation.

Heating of the sample is accomplished by transferring energy from the heating source through the container to the sample. In contrast, microwave heating is accomplished by directly coupling the microwave energy with the molecules that are present in the reaction mixture (Fig. 8.3B). Energy is

Fig. 8.3. Schematic of sample heating by (A) conventional heating and (B) microwave heating.

260

Microwave based extraction

transferred directly to the sample species leading to extremely rapid energy transfer. The result is an instantaneous localized superheating of anything that will respond to either dipole rotation or ionic conduction, the two fundamental mechanisms for transferring energy from microwaves to the substance being heated. Microwave heating also offers simplistic reaction control. It can be described as “instant on–instant off”. When the microwave energy is turned off, latent heat is all that remains. Dipole rotation and ionic conduction are the two fundamental mechanisms for transferring energy from the electric ﬁeld of the microwave to the substance being heated. Dipole rotation is an interaction in which polar molecules try to align themselves with the rapidly changing electric ﬁeld of the microwave. The rotational motion of the molecule as it tries to orient itself with the ﬁeld results in a transfer of energy. Ionic conduction happens when there are free ions or ionic species present in the substance being heated. The electric ﬁeld generates ionic motion as the molecules try to orient themselves to the rapidly changing ﬁeld [3]. The coupling ability of these mechanisms is related to the polarity and dipole moment of the molecules and their ability to align with the electric ﬁeld. There are a number of factors that will ultimately determine the coupling efﬁciency; however, any polar or ionic conductive species that are present will encounter these mechanisms of energy transfer. As the molecule’s dipole responds to the changing electric ﬁeld, its inability to move in synchronization with the oscillating ﬁeld results in an energy transfer. How this energy loss is optimized with the moving ﬁeld will determine to what extent a speciﬁc molecule will heat in a microwave ﬁeld. Mathematically, this is expressed as follows: e00 =e0 ¼ tan d

ð8:1Þ

00

where e is referred to as the dielectric loss factor or complex permittivity and is a measure of the efﬁciency with which the speciﬁc energy can be converted into heat. The dielectric constant or relative permittivity, e0 , is the measure of the ability of the molecule to store electrical charges [4]. The ratio of these parameters determines tan d, deﬁned as the ability of a substance to convert electromagnetic energy into heat at a given frequency and temperature. The tangent delta or loss tangent is the dissipation factor of the sample or how efﬁciently microwave energy is converted into thermal energy. Figure 8.4 shows how both the e00 and e0 for water vary with frequency [5]. The dotted line highlights the value for 2.45 GHz. This data was recorded using a sophisticated microwave measurement device known as a network analyzer. With this instrumentation, the dielectric loss, dielectric constant, and loss tangent values can be determined for substances at a particular frequency and temperature [6]. Microwaves are well suited as a laboratory heating source. The chemicals involved in microwave extraction are both highly polar and exceptionally ionic, providing both types of microwave heating mechanisms. In a well-distributed

261

E.E. King and D. Barclay

Fig. 8.4. Dielectric properties, e00 and e0 , of water as a function of frequency.

microwave system, microwaves will transfer directly to the most highly absorptive species in the system. In microwave extraction, the highly absorptive species are usually the sample and the extraction acid. The superheating of reaction components is essential to the kinetic process of sample extraction. Species absorb microwaves directly at a rate of over 2 billion times per second (2 £ 109 at 2450 MHz). Therefore, energy is transferred in 1029 s. In order to dissipate this absorbed energy, the highly energized molecule must collide with another molecule. Since collisions occur in the 1026 s time frame, the molecules may absorb energy faster than they can dissipate it and superheating occurs. This process of superheating creates instantaneous temperatures ðTI Þ substantially higher than the measured bulk temperatures ðTB Þ: Since temperature, by deﬁnition, is an equilibrium measurement, accurate understanding of temperature is hidden when the rate of dissociation is compared with reactions at the same temperature under non-microwave conditions. In conductive heating, TB and TI are equal because of the lower rate of heating. Therefore, at the same measured bulk temperature, microwave heating will have a higher dissolution rate due to the high instantaneous temperatures. The instantaneous temperature is much higher than the bulk temperature and explains the acceleration of reaction rates. Thousands of documented reaction chemistries are now being performed under microwave conditions and demonstrating decreased reaction times well beyond the normal temperature conditions of the reaction. It is believed that reaction kinetics for microwave reactions are identical and the basic

262

Microwave based extraction

mechanism is same as in conductive heating systems. The only difference between thermal and microwave heating is the rates of the applied energy and the extent to which the number of molecules colliding have sufﬁcient energy to move the reaction towards completion. The rate of effective kinetic collisions to breakdown and extract molecules is enhanced by the substantially higher instantaneous temperatures, which result from the rapid energy transfer of the microwaves. This applies to microwave heating at both elevated pressures and at atmospheric pressure. Microwave heating is generally associated with closed vessel pressurized reactions; however, atmospheric microwave reactions will proceed at a higher rate than those performed via conductive heating due to elevated instantaneous temperatures.

8.4

MICROWAVE LABORATORY EQUIPMENT

Over 20 years of development in microwave laboratory equipment has led to a signiﬁcant increase in the sophistication of these devices. To better study the equipment, an isometric drawing of a modern microwave system, with the internal components displayed, is shown in Fig. 8.5, followed by a brief description of each component.

Fig. 8.5. Schematic of a microwave cavity, magnetron, waveguide, isolator, reaction vessel, and the radiation pathway of forward and reﬂective microwaves.

263

E.E. King and D. Barclay

8.4.1

Magnetron

A magnetron tube is used to generate microwave energy (Fig. 8.6). The magnetron is a cylindrical diode with an anode and a cathode. A magnetic ﬁeld encompasses the diode and is aligned with the cathode. The anode is comprised of many small resonant cavities across which several thousand volts are placed. The theory of magnetron operation is based on the motion of electrons under the combined inﬂuence of electric and magnetic ﬁelds. For the tube to operate, electrons must ﬂow from the cathode to the anode. The force exerted by the electric ﬁeld on an electron is proportional to the strength of the ﬁeld. Electrons tend to move from a point of negative potential toward a positive potential. Figure 8.7A shows the uniform and direct movement of the electrons in an electric ﬁeld with no magnetic ﬁeld present. The force exerted on an

Fig. 8.6. Schematic of a magnetron tube.

264

Microwave based extraction

Fig. 8.7. Electron motion in a magnetron tube.

electron in a magnetic ﬁeld is at right angles to both the ﬁeld itself, and to the path of the electron. The direction of the force is such that the electron proceeds to the anode in a curve rather than a direct path. In Fig. 8.7B, two permanent magnets are added above and below the tube structure. In Fig. 8.7C, assume the upper magnet is a north pole and you are the device viewing from that position. The lower, south pole magnet, is located underneath the page, so that

265

E.E. King and D. Barclay

the magnetic ﬁeld appears to be coming right through the page. Just as electrons ﬂowing through a conductor cause a magnetic ﬁeld to build around that conductor, so an electron moving through space tends to build up a magnetic ﬁeld around itself. On one side of the electron’s path, this self-induced magnetic ﬁeld adds to the permanent magnetic ﬁeld surrounding it. On the other side of its path, it has the opposite effect of subtracting from the permanent magnetic ﬁeld. The magnetic ﬁeld on the right side is therefore weakened, and the electron’s trajectory bends in that direction, resulting in a circular motion of travel to the anode. The process begins with a low voltage being applied to the ﬁlament, which causes it to heat up. Molecular activity within the cathode is the result of the rise in temperature, to the extent that it begins to “boil-off” or emit electrons. Electrons leaving the surface of a heated ﬁlament wire might be compared to molecules that leave the surface of boiling water in the form of steam. They ﬂoat, or hover, just off the surface of the cathode waiting for some momentum. Electrons, being negative charges, are strongly repelled by other negative charges. So this ﬂoating cloud of electrons would be repelled from a negatively charged cathode. The distance and velocity of their travel would increase with the intensity of the applied negative charge. Momentum is thus provided by a voltage source of several thousand DC volts. The electrons blast off from the cathode like tiny rockets and try to accelerate straight toward the positive anode. As the electrons move towards the anode, they encounter the powerful magnetic ﬁeld of two permanent magnets. The effect of the magnetic ﬁeld tends to deﬂect the speeding electrons away from the anode, as shown in Fig. 8.7D. Instead of traveling in a straight path to the anode, they curve to a path at almost right angles to the previous direction, resulting in an expanding circular orbit around the cathode, which eventually reaches the anode. The whirling cloud of electrons, inﬂuenced by the high voltage and strong magnetic ﬁeld, form a rotation pattern that resembles the spokes in a spinning wheel. The interaction of this rotating space-charged wheel with the conﬁguration of the surface of the anode produces an alternating current ﬂow in the resonant cavities of the anode. As the “spoke” of electrons approaches an anode vane it induces a positive charge in that segment. As the electrons pass, the positive charge diminishes in the ﬁrst segment while another positive charge is being induced in the next segment. Current is induced because the physical structure of the anode forms the equivalent of a series of high-Q resonant inductive –capacitive (LC) circuits. The energy resonates at the speciﬁc frequency of the magnetron. The combination of the internal pattern of the magnetron with the acceleration of electrons under the inﬂuence of the magnetic ﬁeld through the anode produces waves of energy in the microwave region. This resulting energy radiates 3608 from the magnetron anode antenna [7,8].

266

Microwave based extraction

8.4.2

Power application

Twenty years ago, magnetrons were fed with cycled power at a duty cycle up to 10 s. This meant for a 50% power input with a 10 s duty cycle, the microwaves were turned on 5 s at 100% power, then turned off for 5 s. The control of the power ﬂow to the magnetron has improved greatly since then. The microwave system discussed in this section utilized a turntable to move the samples continuously through the microwave energy ﬁeld in the cavity. A single rotation of the sample typically takes 7–10 s. Since the distribution of microwave energy throughout the cavity is not perfect, the longer the duty cycle, the more uneven the heating of the moving sample within the cavity. Great strides have been made to eliminate this issue. Progressively, the duty cycle of microwave power supplies has been shortened from 10 s in 1980 to 1 s in 1990 to modern systems with duty cycles of 17 ms. The shortened duty cycle has virtually eliminated microwave uniformity issues and enabled ﬁner control of chemical extractions. 8.4.3

Waveguide

From the magnetron, energy is transferred to the microwave cavity via the waveguide. The waveguide is a metallic rectangular enclosure that collects and transfers the microwave energy through constructive interference of the waves and launches the energy into the microwave cavity with minimal loss. 8.4.4

Microwave cavity

Microwave energy is launched from the waveguide into the microwave cavity. Here, standing wave patterns are established between the walls of the cavity by repeated reﬂections of the microwaves. Constructive and destructive interference creates patterns of energy or modes throughout the cavity, as shown in Fig. 8.8. The interaction of microwaves within the cavity continues until the waves are totally dissipated. When the samples and sample components are placed in the cavity, the samples repeatedly interact with the microwave reﬂections and are heated. The exact distribution of energy within the cavity is affected by the materials, samples, vessels, and detection systems placed in the cavity. Altogether, this is a very difﬁcult, dynamic mathematical problem and beyond the purpose of this chapter. 8.4.5

Reﬂected energy

Lacking any signiﬁcant absorbing material within the microwave cavity, microwave energy can make its way back to the magnetron and become reabsorbed, which results in overheating the magnetron. Magnetrons are cooled and are equipped with thermal fuses to prevent extreme damage from

267

E.E. King and D. Barclay

Fig. 8.8. Schematic of a microwave pattern within a cavity with a turntable, reaction vessels, and temperature and pressure sensors.

overheating. Some modern laboratory microwave systems are equipped with an isolator or circulator. As shown in Fig. 8.9, this device redirects reﬂected energy from the cavity to a cooled microwave absorber and protects the magnetron from overheating. 8.4.6

Mode stirrer and turntables

To promote even heating within the cavity, two separate devices are used for the distribution of microwave energy in the cavity. As the mode stirrer rotates,

Fig. 8.9. Schematic of a microwave energy isolator (circulator) with a magnetron, waveguide, and the radiation pathway of forward and reﬂective microwaves.

268

Microwave based extraction

reﬂections from the metallic stirrer blades change the pathway of the microwaves, or “stir” the modes or energy packets within the cavity. This prevents high energy or “hot” spots from forming. Likewise, turntables are used to rotate the sample containers within the cavity. It is important for a microwave system to have at least one of the devices. 8.4.7

Microwave compatible materials

Like any form of electromagnetic radiation, microwaves interact with materials by reﬂection, transmission, or absorption of the energy. Metallic materials are reﬂective and used for construction of the microwave cavity and structural materials. Vessel materials, whose purpose is to contain and support the sample, are transparent or, at worst, slightly absorptive. All materials have some microwave interaction, and generally, the microwave will be attracted to the most highly absorbing species within the cavity. For example, if a glass support structure, not in contact with the sample, is placed in close proximity to a sample, the glass will barely heat. If you remove the sample from the cavity, the glass becomes the most highly absorbing species in the cavity to the point where the glass can absorb enough energy to melt. Table 8.1 shows some typical materials and how they absorb microwave energy. The second column shows how the listed materials absorb energy compared to water [9]. TABLE 8.1 Microwave dissipation factor of different materials at 258C at 3.0 GHz Material

Tangent d ( £ 104)

Referenced to water

Ethylene glycol Ethanol Methanol Water Fused quartz Ceramic F66 Phosphate glass Borosilicate glass Corning glass No. 0080 Polycarbonate Nylon 66 Polyvinyl chloride Polyethylene Polystyrene Teﬂon PFA

10,000.0 8740.0 6790.0 1460.0 0.6 5.5 11.0 46.0 126.0 57.0 128.0 55.0 3.1 3.3 1.5

6.8 6.0 4.7 1.0 1/2500 1/270 1/130 1/30 1/12 1/26 1/11 1/27 1/470 1/440 1/970

269

E.E. King and D. Barclay

Fig. 8.10. First and latest generation: CEM PFA vessel (on left) and the CEM XP-1500 Plus vessel.

8.5

VESSELS

Microwave vessel design, begun with sealed glass or quartz Carius tubes, has evolved into a complex mixture of advanced molded materials speciﬁcally engineered to handle the stresses of high temperature and pressure associated with closed vessel microwave extractions. Early pressure vessels (Fig. 8.10), constructed of molded PFA Teﬂon, were sealed with a torquing device and incorporated a ﬂexing disc, which allowed excess pressure to be vented before damage to the vessel components occurred. Many of these vessels are still used today, but their pressure and temperature limitations have led to the development of vessels with increasingly higher pressure and temperature capability. Modern 21st century microwave pressure vessel design includes open architecture for rapid cooling and maximum strength to weight ratio, ease of assembly without complex threading, rotor modularity, and the use of composite strength components to assist the inert liner material in resisting the stresses of pressure. The challenge of producing vessels of this strength, that are inert to microwave energy, is a difﬁcult one, bearing in mind that metals, due to their reﬂective properties, are almost completely ruled out as a material source.

270

Microwave based extraction

8.5.1

Materials

Materials of vessel construction fall into one broad class subdivided into three structural areas. First, all materials must be largely, if not completely, transparent to microwave energy. This property allows energy to be coupled with the sample directly, while preserving the structural components of the vessel to withstand the pressure and temperature required to achieve the desired extraction. However, the area or inside layers in contact with the sample and reagent mixture must also be inert and clean, so that they will not contribute a signiﬁcant background contamination to the analysis performed. Generally, this restricts the internal surfaces of the vessel to the following materials: Teﬂon (PFA or TFM), glass, or quartz. Preference for the materials is a trade-off between material cost, mechanical properties, and contamination issues. Any or all of these materials are currently available and they are all appropriate choices for a range of analyses. However, the actual vessel material chosen is usually determined by the conditions required for the extraction, along with the level of the blank obtainable from the material. In general, their uses can be broadly classiﬁed in the following manner: Glass: Used for extractions with organic solvents where elemental analysis is not performed. It is a very economical material, but does absorb microwave energy slightly. Attainment of a very low blank value with glass as a vessel liner material is difﬁcult for elemental analysis, and for this reason, most elemental determinations are performed in vessels whose internal linings are constructed from quartz or Teﬂon. Quartz: Popular in certain areas of the world, quartz can be an excellent material for vessel liner construction. Microwave transparent, it has the disadvantage of contributing a small, albeit somewhat predictable, contamination to each sample. It also prohibits (as does glass) the use of hydroﬂuoric acid as a reagent, since this will digest the quartz vessel. Teﬂon (PFA): This material is widely used for low-to-medium temperature digestion/extractions. It could be said to be the material of choice for these applications. It is fully inert to all reagents, but should not be used at temperatures above 2108C, where it begins to soften. Structurally, as with all Teﬂon-type materials, it requires the addition of strength components around it to allow vessels to withstand pressure and some temperatures, but its moldability means that it can be obtained very clean, with a minimal wastage of material. Part cost is then very economical for the end user, and when utilized within the appropriate pressure and temperature range, the lifetime of vessel liners made from this material can be extensive. Teﬂon (TFM): This formulation of Teﬂon has more structural strength than PFA and better temperature resistance. For extractions occurring over 2108C, it is the material of choice, effectively inert to all reagents. However, it is difﬁcult to mold because of these properties, and thus, parts are often more expensive. Depending on the thickness of the liner itself, vessels utilizing this

271

E.E. King and D. Barclay

material can withstand temperatures over 3008C, though they are commonly employed at pressure and temperature combinations of 2408C and 800 psi. 8.5.2

Structural components

The strength components of pressurized extraction vessel design have changed radically over the last 5 years. The liner and cover containing the pressurized reaction should be composed of an inert, clean material and must be supported with materials which are light, strong, transparent to microwave energy, inert to spillages of the reagents used, and yet, easy to assemble and economically viable. One commonly utilized material, however, should be approached with caution. It can be demonstrated that at elevated temperatures, polyetheretherketone (PEEK) can absorb microwave energy to a signiﬁcant level. This material, at room temperature, offers an economical and strong material for this type of application, but at elevated temperatures, the absorption of energy can cause a runaway melting and failure of the structural components of the vessel. It is our belief that this material has no place in pressurized microwave extraction vessel design and should be avoided for reasons of safety. Other appropriate, inert structural materials available include, composite formulations of polypropylene, Teﬂon itself (when an appropriate thickness), and various ceramic and glassy carbon materials, as well as a range of advanced multi-layer materials which have been custom designed for the purpose by various manufacturers. 8.5.3

Safety

Pressurized microwave extractions require strict adherence to safety rules for several obvious reasons. Appropriate cutouts in the cavity and safety features must be incorporated to ensure that the delivery of microwave energy is made in a safe, controlled manner, and that energy is contained and sealed inside the microwave cavity alone. To accomplish these ends, commercial equipment includes sensors to ensure that energy is applied only under these circumstances. Due to the volatile and gas-producing nature of some reactions performed inside the equipment, allied with the corrosive and hazardous nature of the reagents employed, sensors should be present that also accomplish the following functions: 1. Vessel event detection: Temperature or pressure has either risen or fallen too fast. Feedback data from pressure and temperature sensors can be utilized to detect events occurring inside the vessels that are outside their normal operating parameters. Rapid rises or falls in pressure or temperature can indicate a loss of control by the system over the extraction being performed. Sensors and feedback cutoffs designed to detect these events can provide a

272

Microwave based extraction

2.

3.

4.

5.

safe cutoff for energy and ensure that excessive pressures and temperatures, outside the design parameters of the vessels are not attained. Vessel release or relief sensors: Acoustic sensors or gas sensors can be used to detect the release of material, gaseous or liquid, from pressurized vessels during the extraction. Such sensors should be designed for maximum safety to ensure that the program is halted if they are tripped, since the release of hot corrosive reagent from the vessel followed by subsequent heating by further application of energy can lead to additional damage of vessel components, especially if they are constructed from materials which absorb energy when hot. Door protection: This is a critical area in safety. Since reactions are performed inside vessels, which commonly have an internal volume of about 100 ml and can contain pressures in excess of 800 psi, failure of a vessel can cause the pressurization of the cavity itself. The difﬁculty of containing this pressurization of the cavity in a safe manner without rendering the equipment useless for further reactions has led to the widespread use of pressure relieving doors. Now, all major manufacturers utilize a door sealing mechanism that allows the door to relieve the pressure wave resulting from an over pressurization of the cavity and reseal without damage to the instrument and without releasing vessel contents or material outside the instrument. Solvent sensors: Utilizing solvents to perform solvent extraction work without the use of a speciﬁc solvent sensor should be avoided due to the possibility of a vessel event releasing solvent into the cavity. Explosive mixtures can be formed and vary from solvent to solvent. For this reason, microwave energy should not be applied when solvent has escaped the vessel and is free in the cavity. Upon detection of any such release, the application of microwave energy should immediately be stopped. Vessel style and safety: All serious manufacturers will offer one or all of the following styles of vessel: closed, vent and reseal, and open. Materials of construction may vary, but slightly different safety considerations apply to each style.

8.5.4

Closed vessels

These vessels remain completely sealed throughout the process of the extraction. They allow the highest pressures and temperatures to be utilized and for safety reasons should be equipped with one or more mechanisms by which rapid or excess pressure can be relieved without catastrophic failure of the strength components of the vessel. Such mechanisms may include safety discs made of inert Teﬂon membranes or metallic discs. The reproducibility of the relief point is important in such mechanisms and employment of any disc or membrane whose characteristics or performance changes over time from one run to another should be avoided. Use of these mechanisms for pressure relief

273

E.E. King and D. Barclay

allow the integrity of the vessel components to be maintained for extended use and any over pressurization results only in the failure of one small, disposable component of the vessel. 8.5.5

Vent and reseal vessels

Vessels designed to relieve excess pressure in this manner fall into two types, low-pressure venting and high-pressure venting. The ﬁrst type is designed to relieve gaseous by-products at a set, relatively low pressure, allowing the temperature inside the vessel to continue to rise and the reactions to go to completion. This mechanism allows larger organic or gas-producing samples to be digested, which would otherwise be limited by pressure in the vessel. Care must be taken to ensure that the vent point or release point of these types of vessels is below the point at which any aerosol can be formed by the sample, as this would lead to leakage or analyte loss. The other type of vent and reseal technology that is employed sets the vent pressure of the vessel at an extremely high level and the vent and reseal is used as an alternative to a safety membrane without the advantage of a replaceable, reproducible part. This minimizes the apparent cost, but unfortunately, venting at this level across the sealed surface of the vessel can damage the surface and signiﬁcantly decrease the lifetime of major vessel components. Such mechanisms commonly rely on some sort of spring technology, and all springs wear and decrease in strength over time. This ensures that reproducibility of the vent point is not constant over time, and thus, the performance of the whole microwave system could be irreproducible over any extended period of time. 8.5.6

Open vessels

Open vessel technology relies on extractions performed at or below the atmospheric pressure boiling point of the reagents employed. Any pressure or elevated temperature hazards are avoided, but whole classes of dissolution cannot be performed within these constraints. However, sophisticated equipment is now available which takes advantage of the open nature of this system in order to automate reagent addition. Fume handling is the safety challenge for open systems and the incorporation of some type of fume removal blower, scrubber, or other handling mechanism should be a consideration when using such systems. While lacking the advantages of elevation of reaction temperature which are the mainstay of closed systems, open vessel systems still have the beneﬁts of a microwave heating source and commercial equipment is available with automated reagent and fume handling. In addition, open vessel accessories are usually available for closed systems and open vessel stages are sometimes incorporated into closed vessel dissolution procedures. However, with the improved pressure capabilities of modern closed

274

Microwave based extraction

vessel technology, predigests, or open vessel procedures are not now as necessary except for specialized applications or evaporation methodologies. 8.6

CONTROL SYSTEMS

Microwave extraction, with the inherent advantages of efﬁcient energy transfer and instantaneous on/off control can utilize a range of control parameters or styles. These control mechanisms effectively adjust the input wattage or power and cut the energy on and off in response to feedback provided by various sensor mechanisms associated with the extractions being performed. These systems are the mechanism by which this technology has achieved the wide range of applications it is currently used for, and without them, the usefulness of the system would be severely restricted. As with all technology and control systems, it is an evolving area, and signiﬁcant improvements have been made over time. Modern instrumentation makes available the full historical range of control styles, so that even with increasing sophistication, tried and trusted methods can still be utilized, while new method development can take advantage of the more sophisticated control styles. 8.6.1

Power/time

This was the ﬁrst, simplest and most obvious style of control and was utilized with the earliest equipment available for the laboratory. A very basic control, it essentially consists of an experienced operator who, by trial and error, determines the power level and time for which a reaction should be heated. Further moderation of the program is possible only by use of the “stop” key. This style has actually made somewhat of a resurgence recently with advances in vessel technology, which allow vessels to self-moderate the pressure during the reaction. Figure 8.11 shows a power/time digestion of a spiked lead wipe using self-moderating pressure vessels designed to vent at a speciﬁc pressure. Pressure and temperature were measured, but not used for control purposes. 8.6.2

Pressure

This is the ﬁrst of the two directly measurable physical parameters inside the vessel to be monitored and applied to feedback control. Pressure sensors are relatively inexpensive and the vessel can be constructed to allow remote mounting of the sensor with the instrument conﬁgured to moderate the input energy based on the measured internal pressure of the reaction vessel. Moderation of the input energy can be as simple as feedback limit control, where energy input is cutoff at a selected pressure limit and turned back on when the pressure falls below that limit, or as complex as ramping control, where the instrument applies energy to maintain a single ramp, or series of

275

E.E. King and D. Barclay

Fig. 8.11. Temperature, pressure, and power vs. time for digestion of spiked lead wipes (power/time control).

ramps, and holds to a step series of pressures. The complexity of the control is normally in direct relation to the propensity of the sample type to evolve large amounts of gas in a given temperature range or time period in the digestion. Considerable experience can be necessary in order to accurately assess the pressure control program that should be safely run for a given sample. Unfortunately, this type of control is often mistakenly thought of as an extra safety measure. The pervading impression that pressure is the parameter that has the possibility for causing damage can lead the inexperienced user to a method of control that is exceptionally difﬁcult to reproduce from sample to sample. It can be seen as working against the desired reaction and result. The drawbacks of pressure control alone, as a means of reproducibly achieving a desired extraction, include the following: the relationship between pressure and time during the digestion is not linear, and within the vessel, the actual pressure achieved at the completion of a digestion that evolves gas is dependent upon the amount of sample. Under these conditions or restrictions, reproducibility among a batch of digestions is virtually impossible. Even sophisticated controls, which would allow pressure monitoring of every vessel in a single

276

Microwave based extraction

batch, cannot generate reproducible dissolutions, since there is only one source of energy to be moderated. A choice must be made about which pressure value should control the reactions. In addition, the pressure achieved at a given temperature during the time of the digestion depends on a combination of the vapor pressure of the reagents and the gas pressure evolved by gaseous products of the sample dissolution process. Samples containing large organic polymeric material or complex organic molecules will break down to produce large amounts of CO2, as the sample matrix is destroyed. This often occurs within a relatively narrow temperature range, and if the bulk of the resulting pressure is made up of this component, then power moderation alone based on pressure is insufﬁcient to achieve control of the process. Over the whole program, the bulk of time is spent “under control”, but the bulk of the work of the digest is done in a narrow window of time where pressure is evolved rapidly by the complex reactions occurring. During this rapid pressure rise, power moderation causes energy to be on too much too early, and then off too much too late: in effect, control of the process itself is lost during the critical period of the process. All that can be achieved is a more sophisticated version of limit control applicable to multiple vessels, but having little bearing on the physical parameter temperature that sets the quality of the reaction or process performed. Figure 8.12 shows the results of a multistage pressure ramp on a 0.5 g oil sample digested in a vessel

Fig. 8.12. Multistep pressure control digestion of 0.5 g waste oil sample.

277

E.E. King and D. Barclay

with a pressure maximum rating for control of 200 psi. Temperature was measured, and the variations and ﬁnal temperature achieved demonstrate that unless the ﬁnal pressure is known to have been above that required to generate a high enough temperature for a good destruction, pressure control can result in wide variations in the temperatures of digestions. Far from being a safe parameter to control with, it has the tendency to produce intrabatch variations in the digestion temperature, and hence, the quality of the digest. 8.6.3

Temperature

Temperature is the control style of choice for microwave-based extraction. The quality of the dissolution is directly related to the temperature achieved during the process. Certain classes of compounds digest at certain critical temperatures, and driving the process reproducibly to the desired temperature and controlling it at that temperature for an appropriate period of time allows true reproducible control of a batch of sample dissolutions [10]. Since the temperature of the reaction sets the quality of the dissolution, ramping control, where the energy is varied to achieve a speciﬁed temperature ramp proﬁle in combination with an appropriate hold time at the ﬁnal temperature, allows a reproducible heating proﬁle, and thus, extraction to be achieved both within and between batches of samples. Variations in the heating proﬁle caused by differences in the samples are minimized and full control for most classes of sample is achievable. Unlike pressure control, where a temperature above the boiling point often must be achieved before pressure begins to rise, temperature response is under control throughout the whole process. As the critical temperatures are achieved in the process and the gaseous by-products are evolved, the temperature proﬁle is requested, maintained, and controlled while the pressure is simply allowed to generate within the vessel as the by-product of the process that it often can be, especially for samples classiﬁed as organic. The addition of overlapping pressure limit control maintains the reaction within the safety parameters of the vessel itself, should the reaction proceed to a point where the integrity of the parts may be compromised, but in effect, ramp-to-temperature control provides the maximum control possible for this type of equipment. In order to effectively control this process, a temperature sensor with feedback capability must possess the following attributes: it must be rapid response, truly displaying the internal temperature of at least one vessel in any batch; it must be capable of continuous temperature output faster than the bulk digest can respond; and, it must be inert to microwave energy and posses no lag or bias in relation to the energy input. Suitable sensors include ﬁber optic technology, based on interferometery, phosphorescence decay, or gas bulb thermometry. Infrared thermometry has the advantage of being remote and non-invasive, but typically, it only sees the outside of one vessel at a time, and the lag associated with heat conduction to the outside of the vessel to allow a

278

Microwave based extraction

view means that the response is too slow for reaction control. Infrared technology has, however, been proven useful by a number of manufacturers as a secondary safety mechanism and is used in some cases to ensure that all reactions within a batch are behaving in a similar manner. Proprietary technology exists to minimize the lag effect of IR by continual recalibration to a true internal temperature system, but this still requires a controlling measurement of one vessel to be present. Care should also be taken to avoid metallic sensor technologies such as resistance temperature detectors (RTDs) and thermocouples. While cost effective and simple, the metallic length of such devices makes it certain that they will interact with the microwave ﬁeld and undesired effects, such as self-heating of the probe and antenna effects increasing the ﬁeld density at the measured vessel, can ruin the reproducibility, and hence, the point of a control system within a batch of dissolutions or extractions. Figure 8.13 shows a similar oil sample to Fig. 8.12 at 0.5 g digested under ramp-to-temperature conditions in a vessel with a larger pressure control maximum. Here, the vessel’s pressure limit is sufﬁcient for ramp-to-temperature to achieve a nice smooth ramp and hold at an elevated temperature (2008C). The quality of the digestion is much improved and the pressure rise as the digestion of this organic material proceeds can be seen following the classic sigmoid shape. One can say that the destruction is complete when the pressure fails to continue to rise. That does not mean the whole sample is digested,

Fig. 8.13. Single-step ramp-to-temperature control of 0.5 g oil sample.

279

E.E. King and D. Barclay

just that all the gas that can be generated at the hold temperature has been generated when the pressure rise stops. Ramp-to-temperature offers a reproducible control parameter to ensure energy input is the same during the program for all samples; provided the vessels within a batch are the same, or have the same thermal properties, and the samples have similar absorbing capabilities, all samples should achieve similar ﬁnal temperatures and produce very similar dissolutions. 8.6.4

Power optimization feedback

Power optimization feedback operates in tandem with the most up-to-date temperature systems, generating a degree of control over the batch process of microwave extraction that has not previously been possible. In this system, instead of a simple on/off control at a set power level in response to sensor feedback, the power level of the microwave device is adjusted in response to the measured “on time” of the device actively during the process. This maximizes the on time of the microwave by a hunt-and-seek mechanism, in which the instrument itself ﬁnds the power level or percentage that will just meet the programmed parameters. Thus, instead of the operator having to make a decision on percentage power input, and the instrument merely applying, or not applying, that power level to follow a programmed temperature ramp, the on-board computer will adjust the power level to maintain a percentage on time for the power, at the appropriate level of greater than 90%. Figure 8.14 shows a representative nitric acid dissolution under ramp-totemperature control of a 0.5 g plant material sample. The power ﬂuctuations can be seen as the lower line on the graph, with pressure and temperature plotted above. As can be seen, the input power starts high, then decreases steadily as the reaction temperature rises in order to maximize the “on time” of the microwaves. After the reagents (nitric acid) reach their boiling point and hot vapor condenses on the upper walls of the liner, the area for heat loss dramatically increases and more energy is required to maintain a constant temperature rise for the bulk of the liquid containing the sample. Thus, above around 1208C the power increases to meet this increased demand. Once the hold temperature 2008C is achieved, the power decreases automatically, as the energy demand is reduced with no further temperature increase required. Effectively, the instrument can be seen to be optimizing the power required in order to maximize the on time of the energy source and allow the ﬁnest control possible at the programmed method set point. Here, you can see the dependence of the earlier control style on operator experience to estimate the power setting for the microwave is no longer required, as the instrument’s sophisticated optimization process hunts and seeks the optimal power to be applied based on the current power usage. At least two current manufacturers offer some kind of power optimization capability, rather than the more traditional on/off control at a set power level.

280

Microwave based extraction

Fig. 8.14. Power optimization feedback digestion of citrus leaves.

8.7

METHODOLOGY

In this section, we will deal with a number of examples of microwave extractions. Broadly divided into categories, an example of a representative extraction of each type will be presented along with graphical representation of the resulting extraction. Analytical results or elemental analyses will not be presented, but the data shown is generated by actual experimentation. For the analyst performing an extraction by microwave instrumentation, the initial questions one must ask oneself are the same as those asked for any other dissolution procedure: 1. How much sample do I need to digest? The answer to this question depends upon a multitude of factors associated with the detection limits of the analytical equipment to be used along with the homogeneity of the sample itself. These criteria, along with other considerations associated with sample size, are dealt with elsewhere in this volume. General guidelines on sample size for microwave open pressure extractions are included in the examples in the following section. The information and examples can be used as a representative tool for ﬁnding a starting point for the extraction program for most types of samples. 2. Which reagents should I use? This again depends on the circumstances associated with the sample itself, that is the type of digest or extraction

281

E.E. King and D. Barclay

required in order to make the analytes of interest available for analysis. Generally, however, if full digestion of silicate material is required, and even plant material for example contains some windblown silicate, then HF must be used. If silicates are present and HF is not an option, then a leach involving nitric acid is appropriate. Hydrochloric acid has specialized applications in the digestion of precious metals and is used in concert with nitric where the solubility of some elements needs to be improved. Sulfuric acid is rarely used except for predigests on large (. 2 g) organic samples where a char is required and other specialized applications, such as certain carbon catalyst materials. Hydrogen peroxide has limited use in microwave pressure dissolutions; the increased oxidation potential of nitric acid at 2008C or above means that the addition of peroxide is not required for good carbon destruction of organic matrices. Also, the vigor with which peroxide can react with certain samples and the large volumes of gaseous products can pose a safety hazard in a sealed system. In open vessel work, however, peroxide can be an excellent means with which to ﬁnish, or “clear up” an organic digestion. Perchloric acid is an excellent oxidant and is extensively used in certain areas of the world. Once the reagent of choice for hot plate digestions of soil and sedimentary material, its use has dropped in the developed world as concerns about its tendency to form explosive peroxides, if the sample dries, have become more prevalent. In the United States, a special fume hood is required if perchloric is to be used, and the expense of this type of hood has also contributed to a reduction in the use of this reagent. 3. Which heating program should I use? Again, the answer depends on the results required. Generally, however, use a program and temperature that will achieve the dissolution required without stressing the pressure vessels unnecessarily. Most manufacturers will provide applications advice, or at least a starting point for the procedure to be used, upon the purchase of the equipment and some provide free ongoing applications assistance for the lifetime of the instrument. So, if in doubt, call the manufacturer and ask, let them know the sample type, the analytes of interest, and other pertinent information such as any volume or reagent constraints, and in all likelihood, they will be able to provide methodology for an extraction. Included in the following section are examples of each of several broad classes of extractions. These cover about 90% of all sample matrices encountered and are useful starting points for the process. 8.7.1

Pressurized closed vessel extractions

The methodology for pressurized closed vessel technology has evolved over time, as discussed above, through power/time and pressure control, to state-ofthe-art full feedback internal temperature control. The representative extractions outlined below are performed with the latest available technology,

282

Microwave based extraction

including ramp-to-temperature control, and the methodology, as demonstrated, can be performed and repeated on any manufacturers’ instrumentation equipped with appropriate control and sensor technology. No great (or indeed any) attention will be given to theoretical considerations of partial gas pressures or molar fractions of compounds, since real world samples by their nature are unknown—if we knew what was in them, we would not be analyzing them! Such experimentation is useful for vessel and equipment design, but we believe real world operators prefer useful representative examples of reagent combinations and temperatures at which acceptable extractions can be performed for common sample types. It is this information, which we will try to convey here. It is, however, worth noting that there has been extensive discussion as to the mechanism by which microwave pressure vessels achieve such elevated temperatures for reagents, while the reagent vapor pressure remains consistently lower than expected. An excellent treatment of this discussion can be found in Haswell and Kingston along with other theoretical aspects of this area of science [11]. But unfortunately, real world samples are largely unknown and of limited availability. They are typically only 1–2 g in total, so the opportunity for extended study is limited, and hence, as a general rule use enough reagent to ensure it is not totally consumed before the extraction is complete and program a temperature proﬁle which will achieve the required extraction and ensure that the pressure limit of the vessel will not be exceeded. In general though, pressurized closed vessel reactions involve the following components: an appropriate sample size (usually % 1 g, bearing in mind a representative vessel volume of about 100 ml), a reagent combination capable of producing the required extraction at a given temperature, usually 5–10 ml, a ramp-to- and hold-at-temperature capable of producing the extraction with the reagents used, and a ﬁnal hold temperature usually in the range of 160 –2108C. This view may seem somewhat simplistic, but experience over many years tells us what reagents and temperatures will give what kind of extraction on a given sample type. The information below gives a starting point or representative procedure for each of the major sample classes that are encountered. Figure 8.15 shows a representative closed vessel batch digestion/ extraction system with pop in pressure and temperature sensors and high pressure digestion vessels can be seen inside. 8.7.2

Atmospheric open vessel extractions

Open vessel extractions have a lower degree of complexity, simply because the range of choices is more limited than closed vessel pressurized work. Here, the temperatures are essentially limited by the boiling points of the reagents and pressure is not a variable in the procedure. Basically, choose your reagents and follow a program to replenish, exchange or add additional or different reagents over a time period with the resulting extraction limited to the oxidation

283

E.E. King and D. Barclay

Fig. 8.15. Picture of CEM MARSe 5 system with a turntable of 12 vessels including pressure and temperature control systems.

potential of the reagents at or below their atmospheric boiling points. Use of high-temperature reagents, such as sulfuric acid, does allow small additions of, for example, nitric acid to be made at points where the bulk temperature of the reaction is well in excess of the boiling point of nitric acid. Thus, the hightemperature reagent can be seen to act as a thermal diluent for the extraction, allowing nitric oxidation to be performed at elevated temperatures; however, since the low-temperature reagents boil off rapidly, it can have a high reagent use and associated background. Use of sulfuric acid can lead to viscosity transport problems with nebulizers in subsequent analytical procedures, so great care must be taken in matrix matching. Recently, open vessel technology has evolved to perform the following functions: dissolution at atmospheric pressure with automated reagent addition and ramping temperature control; and, reagent exchange and automated evaporation to dryness with subsequent re-solvation of extractant if required. The attraction of this technology lies in the automated reagent handling and the speed and ﬂexibility with which the extractions can be manipulated, with additional reagent added as required or modiﬁcations to the reaction mixture made during the process. Automated reagent handling with individual pumps for each reagent can be an attractive feature in today’s laboratory with its emphasis on GLP and workplace safety.

284

Microwave based extraction

Fig. 8.16. Picture of a CEM STARe6 open vessel microwave system.

Figure 8.16 shows a representative open vessel digestion system with vapor containment and reagent addition features. 8.8

SAMPLE TYPES

This section deals with representative extractions of a range of sample types commonly encountered in the analytical workplace subsequently requiring elemental analysis. 8.8.1

Inorganic

Inorganic samples for the purposes of this chapter are classiﬁed as those samples requiring extraction or dissolution, which do not evolve signiﬁcant amounts of gas during the process. These samples develop pressure during the temperature program, most of which is vapor pressure from the reagent combinations used. Aqua regia, of course, will produce gaseous pressure as well as vapor pressure alone. This is not wholly the case, but these classes of samples usually require higher temperatures and develop signiﬁcantly lower pressures than samples composed primarily of organic carbon containing material. This is because extractions performed where the bulk of the sample is

285

E.E. King and D. Barclay

converted to gaseous material have an additive pressure above the vapor pressure of the reagent alone at any given temperature. 8.8.2

Leaches and other partial extractions

Soils sediments, sludges, oils, and many samples requiring an acid extraction without total sample matrix decomposition. Samples where the analytes of interest can be solvated without a total destruction. Incomplete dissolutions are extensively used for inorganic materials such as sediments and minerals in situations where an assessment of the biological hazard presented by the material is to be made, or where some part of the sample is prohibitively difﬁcult to solvate, or where elements of interest are not thought to be present in signiﬁcant amounts in the undissolved fraction. These extractions are performed at elevated temperatures, but the reagent combinations used are combined with the temperature proﬁle to give a resulting elemental extraction from which meaningful comparisons can be made between samples extracted in the same way. In fact, under certain circumstances, it could be construed as misleading: for example, providing a total dissolution where an assessment of toxicity for an element present may rely on that element being extracted by stomach acids after ingestion (i.e., if it takes 5 ml of HF and 2408C to break it down, how much will be extracted during a 10 h passage through an animal intestine?) When developing these procedures, the total concentration of all elements is traded off, in information terms, for the meaningfulness of an incomplete dissolution and the comparisons that can be made between samples where the same extraction reagents and temperatures are performed. For this type of extraction, however, it is possible to “overcook” the samples and care must be taken to perform the extraction the same way on each sample class. An example here is the US EPA procedure for “leachable extractables in soils, sediments, and sludges.” This methodology was developed so that comparisons can be made between samples without a total digest. The variability in soils and sediments, particularly from place to place, means that each sample may require a different set of reagents and conditions in order to achieve complete dissolution. This is patently impossible from any reasonable standpoint, and hence, each sample is subjected to a narrow range of applicable reagents and conditions that have been determined to provide meaningful analytical results, which can be compared between samples and batches. Thus, assessment of hazards and other results of elemental concentrations can be made and compared from area to area, laboratory to laboratory. While such procedures often take time to develop and technology often passes the written procedures by while they are in the publishing stage, there is no other viable alternative to collecting meaningful data for biohazard information for comparison purposes.

286

Microwave based extraction

Fig. 8.17. Temperature/pressure for digestion of oily soil by US EPA Method 3051.

US EPA Method 3051: Microwave-assisted acid digestion of sediments, sludges, soils, and oils. Oils, of course, are classiﬁed here as organic and can generate high pressures, so this example will deal with the digestion, or extraction, of an oily soil from a sample collected from a garage forecourt (Fig. 8.17) [12]. 8.8.3

Complete dissolutions

8.8.3.1 Soils, sediments, sludges, and materials containing silicates and titanium dioxide Due to the variable nature of sample matrices, care should be taken when attempting a complete dissolution of inorganic material. Slight variations in sample composition can alter the reagent mixture required and the temperature necessary to achieve a complete solvation of the sample. A more appropriate question might be—what will it take to extract all of each of the analytes of interest? However, protocols exist for certain sample types, which, if followed, should yield complete dissolutions. Other inorganic materials or

287

E.E. King and D. Barclay

Fig. 8.18. Digestion of soil sample by US EPA Method 3052.

complex inorganic components require several steps and the examination of residue followed by subsequent continued treatment to achieve complete dissolution. Procedures such as US EPA Method 3052 can be followed, but complete dissolution of the entire sample is often not possible even with this procedure and a residue often remains to be ﬁltered [13]. Any alumina present in the sample, for example, will not be digested with the reagent combination protocols allowable with this method. Hydroﬂuoric acid which has been used can be neutralized after the heating program by the addition of 4% boric acid per the individual manufacturer’s recommendation. The boric sample mixture is then heated in a second stage to above 1208C and the residual hydroﬂuoric acid is neutralized, making further sample handling easier. Care must be taken with these digestions that insoluble ﬂuorides are not precipitated out, such as CaF, and the boric acid stage can assist in solvating such precipitates, as well as neutralizing the residual HF. Basically, chemists should always make sure that any residual solid after an HF digestion is indigestible, rather than precipitated ﬂuorides, and performing a boric acid step can conﬁrm this before ﬁltering (Fig. 8.18). 8.8.4

High-temperature extractions

8.8.4.1 Alumina, alpha and gamma, certain ceramics, carbides, and other inorganic chemicals and compounds Simple non-complex matrices, such as alumina, can be completely dissolved and the trace elements solvated with the following procedures. Modern vessel

288

Microwave based extraction

technology allows the dissolution of certain compounds at elevated temperatures to occur with reagents that would previously have been impossible to get hot enough to do the job in any convenient time frame. One example is the digestion of alumina, which traditionally is done at high temperature by a mixture of sulfuric and phosphoric acids, but can now be achieved with HCl alone. While both are high temperature extractions, the HCl leaves a more manageable matrix for subsequent analysis. In Fig. 8.19, the temperature is taken to 2708C and held for 30 min in order to achieve the dissolution required when using sulfuric and phosphoric acid. As can be seen, the vapor pressure of these high boiling point reagents is quite low, less than 50 psi, throughout the procedure. The graph alongside shows a high temperature and pressure digest of the same sample with HCl alone. Here, the sample is taken to 2408C for 60 min, but the resulting reagent matrix is much more manageable than the sulfuric –phosphoric mixture for the analytical instrument step (e.g., ICP) which will follow.

Fig. 8.19. Comparison of two different procedures for the digestion of alumina.

289

E.E. King and D. Barclay

8.8.5

Complex sequential extractions

8.8.5.1 Geological ores, slags, ceramics, and a range of other minerals and rocks One of the advantages of microwave pressurized extractions is that dissolution of materials, which would normally require fusion, is possible in these systems. An example of this is an extraction such as chromite ore. The matrix problems associated with fusion on this sample mean that although the procedure involves several steps, with care being taken with reagent proportion, the resulting solution is cleaner and more manageable analytically than the results of a fusion. These types of dissolutions often require special attention in regard to the volumes and concentrations, as well as the order in which the reagents are added to the sample. So, if the chemist is unsuccessful, he or she should contact the manufacturer’s application assistance staff. Their experience with these types of dissolutions can often mean the difference between success and total failure in methodology. Figure 8.20 below shows the sequence of steps

Fig. 8.20. Sequence of stages for the digestion of chromite ore.

290

Microwave based extraction

necessary to achieve a complete dissolution of chromite ore. The composition of this sample; (50%Cr2O3, 20%Fe203, 15%Al203, 10%MgO, 5%SiO2, and 1%CaO) demonstrates nicely that various parts of the sample matrix must be attacked in sequence. Here, three stages are used. In the ﬁrst, the alumina component is digested with sulfuric/phosphoric acids, as above by heating to 2708C for 30 min. Then, the rest of the matrix is broken down by adding a further 5 ml nitric acid and 2 ml hydroﬂuoric acid to the residue and heating to 2608C for 15 min. Finally, the HF is neutralized and any CaF2 precipitated is redissolved with the addition of 30 ml boric acid (4%) and heating to 1608C for 10 min. The resulting dissolution is clear, greenish in color, and free of particulates. 8.8.6

Organic

8.8.6.1 Plant material, oils, greases, plastics, polymers, some chemicals, solvents, and sewage sludges, etc Organic materials, here classiﬁed as those containing high percentages of carbon polymeric material, present a range of challenges to the extraction. Limitations on sample size are often set here by the pressure capabilities of the vessel technology rather than the dissolving power of the reagents or their amounts. A whole range of carbon backboned polymeric materials will breakdown almost completely to gaseous products leaving a small elemental fraction of the original sample dissolved in the reagents used. However, a large residual pressure will be present, and during the procedure, pressures for 0.5 g of oil, for example, can reach in excess of 600 psi in a 100 ml vessel when nitric acid is used at a temperature of 2008C. Often, compounds and different polymeric backbones will digest at slightly different temperatures, and once going, these dissolution reactions can become self-sustaining exothermic reactions—where the reaction proceeds without further energy input from the microwave and a large pressure spike is seen over a narrow temperature range. The general rule of thumb to follow for these types of samples is: if you can get them to 2008C in nitric acid, an adequate carbon destruction will result, which will allow the vast majority of analytes to be determined with routine analytical equipment. This type of sample should dilute water clear and colorless after treatment at 2008C, and provided extremely low detection limits are not required a higher temperature is not necessary. Some ring structures are extremely stable at high temperatures and may not be broken down at 2008C. If complete destruction of these structures is required, for solubility or sample detection limit reasons, then the ﬁnal hold temperatures of the procedure should be raised to 2208C to ensure completion. It should be remembered that an adjustment of this kind to the ﬁnal temperature will be accompanied by an increase in ﬁnal pressure. Again, care should be taken to ensure the pressure limit of the vessel is not achieved, so that the procedure reaches the programmed temperature as planned. As described earlier, the

291

E.E. King and D. Barclay

pressure limit of the vessel will set the effective sample size, and provided the vessel can safely contain sufﬁcient pressure to allow the sample to reach 2008C, no problems should occur. If the sample is too large for the pressure limit of the vessel and the setpoint temperature is not achieved due to pressure constraints, an alternative to purchasing higher pressure vessels is to cool the vessels after heating, vent the gas produced, and then, rerun the program. Here, the majority of gas produced is vented off after the ﬁrst run with the reaction (e.g., 80% complete). The residual destruction makes less gas and the second run can proceed to the set point without problem. 8.8.7

Carbohydrates

8.8.7.1 Sugars, syrups, and beverages containing sugars Carbohydrate compounds or compounds containing large amounts of carbohydrates in relation to other organic compounds should be digested with care. Reaction begins relatively early in the process at a lower temperature than most other commonly encountered complex organic compounds, such as fats and oils, and goes to completion over a narrow temperature range. Effectively, the bulk of the digestion, and hence the bulk of the gaseous by-products of the digestion, are formed in a narrow temperature range around 1408C. In order to maximize the control possible over such samples, ramp-to-temperature control should be used with the selected rate of temperature rise relatively low, especially over the critical temperature range where the bulk of the digestion is occurring. Equipment manufacturers are known to use household sugar samples as one of a series of tests to verify the safety of their vessel and instrument designs due to the large rate of pressure rise at the critical range. Such care should be applied to all samples containing high percentages of carbohydrate such as syrups, sugars, and ﬂours. In Fig. 8.21, we can see the difference between running this sample in a low-pressure vessel with and without a predigest. Initially, the vessel’s pressure limit is reached early, and the temperature stays very low and never achieves a value high enough to give a good dissolution. After the predigest, the gas to be formed in the heating program is of a small enough amount to allow the temperature to be higher before the pressure limit is reached. An alternative is to use a higher pressure vessel, but this will not slow down the rapid exothermic reaction as the digest accelerates, and certainly for unfamiliar samples of this nature, a predigest is always a good idea. Another tool available is the addition of a small amount (0 –5 ml) of water to the digestion mixture. Often this will slow down or retard any extremely vigorous reactions that may occur. Some reactions involved in the digestion process are temperature speciﬁc, and once the critical temperature is achieved, they occur rapidly with self-generation of heat. Water can act as a moderator for these reactions, and conveniently, it can often improve the solubility of some of the extractants [14].

292

Microwave based extraction

Fig. 8.21. Comparison of wine digestion with and without predigest.

8.8.8

Proteins

Vegetable or animal derived proteins or samples such as meats, dry or wet, often contain high protein values. These compounds tend to break down and evolve gas, and thus, generate higher pressures at a higher temperature than carbohydrates, but the same methodology applies. Moderation of the ramp speed of temperature rise will increase the level of control and ensure that the pressure rise does not occur too rapidly. Sample size, as with all largely organic samples, should be carefully considered and selected in relation to the run or the pressure limit set by the manufacturer for the pressure vessel in use. There is no point in adding 1 g of dried meat to 10 ml of nitric acid and programming a 200 psi rated vessel to go to 2008C with ramping control. In the critical range of pressure evolution, the sample vessel will hit the pressure limit and the energy input will be moderated in response to this limit. The temperature of the reaction will be well below the 2008C target and a nasty yellow incomplete

293

E.E. King and D. Barclay

digestion will result as the instrument safety limits effectively halt the process before complete dissolution of all of the sample. If the pressure limit control is approached at too high a rate and the reaction is self-sustaining, then the pressure may continue to rise past the operating limit without any energy input from the device, and vessel failure may result. This will effectively shut down the program and all of at least one sample will be lost completely. All other samples will remain inside the unit, under pressure, hot and at some partial completion point in the process. Extreme care must be taken with any samples that will be restarted, since the exact pressure values of the noncontrol vessels are not measured and the rate of cooling of each can be unpredictable. Hence, no halted procedure should be continued without cooling the samples down and venting off any gas that has been evolved before resealing and beginning again. It is critical that all samples within a batch are at the same temperature and pressure conditions when the batch heating cycle is begun. 8.8.9

Fats, oils, and waxes

Fats, oils, and waxes are often the most difﬁcult compounds to digest effectively in organic samples and require higher temperatures in order to achieve the same quality of digestion, or residual carbon value, as other organic compounds. Often, they have the added disadvantage of being semi or somewhat liquid at temperatures below the point of digestion, but above room temperature. They are non-miscible with aqueous solvents, and they essentially ﬂoat around on top of the acid until the temperature is achieved at which they begin to digest. The surface area available for attack by acid is low up to the point at which the digestion is proceeding rapidly. Care must be taken here with the carbonization of these samples, as the digestion is partially complete. Small amounts of carbonaceous material can be thrown onto the sides of the vessel above the liquid level, dry out and interact with the microwaves to provide a source of ignition. If the partial pressure of combustible vapors is within an appropriate range, small ﬁres inside the vessel can result, causing damage to vessel components. Often, there is insufﬁcient oxygen inside the sealed vessels for such ﬁres to burn for more than a few milliseconds, but the heat of the ignition can melt or damage seal surfaces or Teﬂon liner components. Certain vessels, which are designed to vent their contents at extremely high pressures (above 500 psi) as a safety feature, can aerosol the contents of the vessels and effectively throw partially digested organic material out of the vessel onto surrounding parts. This material can further carbonize and provide a source for ignition that has a plentiful supply of oxygen and actual ﬁres can result. For this reason, we believe that all vessels which vent their contents at elevated pressures as a safety feature due to pressure rising too rapidly, should cause the microwave program to be halted such that restart can only be accomplished by a trained operator examining the

294

Microwave based extraction

Fig. 8.22. Conostan oil nitric acid digestion, sample size pressure – temperature comparison.

vessel set and electing to continue. In our estimation, it is unsafe for instrumentation to be designed so that even though the contents of a vessel are vented in excess of 800 psi into the cavity, due to an overpressure situation, the instrument continues to complete the programmed heating cycle. The instrument should stop the heating cycle at this point, having detected the event with a safety sensor. Figure 8.22 shows the inﬂuence of sample size on this type of digestion. Care must be taken, as can be seen, that the gas formation and exothermic reaction does not cause the instrument to lose control of the process (i.e., the halting of microwave energy does not stop the reaction). At this point, as you can see for the larger sample size, control has been lost and any further energy applied could result in the venting of the vessel by its safety mechanism. 8.8.9.1 Complex organic extractions Most real world samples of an organic nature are complex mixtures of all of the above or other types of carbon polymer structures. Plant materials, meats, tissues, mixtures of chemicals, solvents, greases, and oils are encountered, and the relative amounts of each fraction are not generally well documented for

295

E.E. King and D. Barclay

every sample. For this reason, calculations of possible partial pressures of gaseous materials formed and predictions of critical temperatures to be achieved is not possible, and it is somewhat a waste of time to bother trying. A few basic rules and a few target temperatures will allow most samples to be effectively digested without the use of an expert system, provided a general knowledge of the organic contents of the sample is known. It is not necessary to understand the exact make up of the sample, but just whether it is mostly of an organic nature or mostly inorganic. If it contains signiﬁcant organic content, be it carbohydrate, oil, fat, or protein, then it should be treated as organic. Assumptions should then be made for sample size based on the amount of pressure that could be generated if it was completely digested to gaseous products, and a temperature program selected which will achieve a temperature sufﬁcient for a good carbon destruction on the most complex types of materials—commonly 200 –2108C. Basically, if you have not seen the sample before, treat it as organic, use a low sample size, ramp the temperature slowly to 2008C, and hold it there for a period long enough for you to see that further pressure is not being evolved. If it is organic and you can get it to 2108C, it will often look like water upon dilution and no yellow color will be visible. 8.8.9.2 Optimizing the analysis Here are some basic rules for approaching organic or reactive samples. Assumptions are made regarding vessel volume and the vessels being completely sealed, along with the relative rate and pressures achieved for these materials, but they provide an excellent set of guidelines for approaching these samples. If the sample has the appearance of containing small molecule solvents, gasoline fractions, or anything like these materials, add the acid and leave it sitting for 30 min to predigest. The more gas you can get out of the sample before the vessel is sealed, the less material there is to generate pressure after the vessel is sealed. For a vessel volume around 100 ml with a control pressure limit over 700 psi, start with a maximum sample size of 0.5 g. If after trying a representative sample it is discovered that 0.5 g generates a rate of pressure rise which is not excessive and a ﬁnal pressure at the selected temperature that is well below the pressure control limit of the vessel, increase the sample size. But, most organic samples will remain within a pressure limit of 700 psi at 2008C at a sample size of 0.5 g. If the vessel is rated to 200 psi, start with fewer samples. Sample size will set the temperature that can be achieved, because too much sample will cause the pressure limit to be reached and the program will run at the pressure limit with the temperature well below that required for the extraction. Most organic samples will give an acceptable digest for analysis if 2008C can be achieved and held for 10–15 min. Set the ramp time carefully, especially if the sample is unknown. We suggest a 15 min ramp time to achieve 2008C.

296

Microwave based extraction

This rate of temperature rise will allow good control of reaction generating pressure as it occurs. Some samples, which are more difﬁcult to digest completely, may require higher temperatures. It should be remembered that additional pressure will be generated from the reagent as a result of the higher vapor pressure necessary to achieve these temperatures in addition to any extra gas that may be generated from an increased destruction of the carbon containing material. 8.9 8.9.1

ADVANCED APPLICATIONS Clean chemistry

Much advantage has been taken of the sealed nature of the pressurized digestion vessels in order to improve the overall “cleanliness” of the digestion process. The level of the analytical blank for the digestion process has been shown to be reduced by the use of this technology, both as a result of the environmental control possible with sealed vessels, as well as the reduced time taken for the dissolution process itself. Basically, the advantage of this approach is that rather than provide an entire clean-room for the process, good control of the environment and contamination level can be obtained by the provision of a clean area for sample storage and vessel assembly in combination with a clean area for vessel disassembly and subsequent sample transfer. The sealed nature of the vessels minimizes the possibility of invasion of contaminants during the process itself. This can prove exceptionally cost effective over the provision of entire clean-room facilities, and provided appropriate steps are taken to keep the materials of vessel construction clean to the level appropriate for the analysis, it is a major advantage for the use of this technology [15]. 8.9.2

Concentration/evaporation

There are many available accessories and options for vessels of various sizes, and pressure and temperature limits available from a range of manufacturers. However, recently, the use of evaporation technology has become extremely popular with microwave extraction technology. One of the inherent features of heating by microwave energy, as discussed in the earlier sections, is the fact that the microwaves move through microwave transparent vessel materials and are absorbed by the sample/reagent mixture itself. This facility is taken advantage of in evaporation technology, in that, if the samples after dissolution are irradiated while connected to some type of vacuum system, the residual digestion can be evaporated down to dryness rapidly. When dryness is achieved, there is no reagent left to absorb the energy and the temperature drops rapidly. This drop can be used by the instrumentation to automatically sense the moment of dryness and to stop the application of energy. Thus, the

297

E.E. King and D. Barclay

Fig. 8.23. Evaporation of 10 ml of oil digestion in CEM MicroVap accessory.

equipment is capable of rapid, automated evaporation to dryness with no possibility for overheating the samples. Figure 8.23 shows the evaporation of 10 ml of digest resulting from an oil sample’s treatment with concentrated nitric acid. The residual solution is evaporated under control, with auto cutoff in the same vessel liner in which the original dissolution was performed, without transfer. The instrument control is labeled as such, and the simultaneous temperatures of three of the other ﬁve vessels are shown to demonstrate the uniform rate of drying across a batch of samples. 8.10

CONCLUSIONS

In the last 20 years, the ﬁeld of microwave extraction has expanded dramatically, with tens of thousands of instruments now in routine use in analytical laboratories around the world. Steady improvements in vessel and instrument technology over this period, allied with improved microprocessor systems have meant that safety, control and optimization procedures for this methodology have also reached a peak of achievement. In summation, we feel that this technology is now mature and accepted, and current developments in this area are now demonstrably focused on ease of operation, usability, and safety. Vessel technology, however, continues to push the limits of the construction materials for pressure, temperature, and size. Barring any radical improvement in high temperature inert plastics (the invention of a super high

298

Microwave based extraction

temperature alternative to ﬂuoropolymers), future systems will strive to extract more samples in the same size cavities or smaller more convenient systems. It will be interesting to view this discipline in another 20 years and reﬂect on state-of-the-art technology for sample extraction at that time.

Acknowledgements We express our thanks and appreciation to the many professionals who have helped to complete this chapter. The authors would like to give a special thanks to J.C. Gallaway of the Microtech Corporation for his detailed contributions on how microwaves are generated from magnetron tubes. Also, a special thanks to CEM Corporation’s Marketing Communications department, especially Michelle Horn for her editorial skills and Steve Smith for his preparation of the artwork and drawings. Their assistance in the completion of this chapter has been invaluable. REFERENCES 1 2

3 4 5 6

7 8 9 10

11

12

13

B.L. Hayes, Microwave Synthesis: Chemistry at the Speed of Light. CEM-Publishing, Matthews, NC, 2002. E.D. Neas and M.J. Collins, in: L.B. Jasssie and H.M. Kingston (Eds.), Introduction to Microwave Sample Preparation: Theory and Practice. American Chemical Society, Washington, DC, 1988, p. 2. D.A. Copson, Microwave Heating. AVI, Westport, CT, 1975. F.E. Terman, Radio Engineers Handbook. McGraw-Hill, New York, 1943. U. Kaatze, Radiat. Phys. Chem., 45 (1995) 549. D.M.P. Mingos and D.R. Baghurst, in: S.J. Haswell and H.M. Kingston (Eds.), Microwave-Enhanced Chemistry: Fundamentals, Sample Preparation, and Application. American Chemical Society, Washington, DC, 1997, pp. 3– 53. J.C. Gallaway, The Complete Microwave Oven Handbook, Operation, Troubleshooting, Maintenance, and Repair. Prentice Hall, New York, 1989. J.C. Gallaway, The Magnetron Tube, Structure and Operation, http://www.gallawa. com/microtech/magnetron.html, 1989. A.R. Von Hippel, Dielectric Materials and Applications. John Wiley, New York, 1954, p. 301. L.B. Jassie and H.M. Kingston, in: L.B. Jasssie and H.M. Kingston (Eds.), Introduction to Microwave Sample Preparation: Theory and Practice. American Chemical Society, Washington, DC, 1988, p. 6. S.J. Haswell and H.M. Kingston (Eds.), Microwave-Enhanced Chemistry: Fundamentals, Sample Preparation, and Application. American Chemical Society, Washington, DC, 1997. US-EPA Method 3051: Microwave assisted acid digestion of sediments, sludges, and soils, and oils. In: Test Methods for Evaluating Solid Waste. U.S. Environmental Protection Agency, Washington, DC, 1986. US-EPA Method 3052: Microwave digestion of silceous and organically based matricies. In: Test Methods for Evaluating Solid Waste. U.S. Environmental Protection Agency, Washington, DC, 1996.

299

E.E. King and D. Barclay 14

15

300

H.M. Kingston, et al., In: S.J. Haswell, H.M. Kingston (Eds.), Microwave-Enhanced Chemistry: Fundamentals, Sample Preparation, and Application. American Chemical Society, Washington, DC, 1997, pp. 223–349. E.M. Shelly and F.T. DiStefano, Clean room and microwave digestion techniques: improvement in detection limits for aluminum determination by GF-AAS, Appl. Spectrosc., 42(7) (1988) 7.

Chapter 9

Fusion and ﬂuxes Fernand Claisse

9.1

INTRODUCTION

Fusion in borates is well known as a sample preparation technique for X-ray ﬂuorescence (XRF) analysis, but it is also used for the preparation of solutions for other spectrochemical methods and in wet chemistry. The present chapter is particularly dedicated to the making and use of fusion beads for XRF analysis, but since the procedure to make solutions is similar and simpler, it is described at the same time. Fusion beads are sometimes disregarded for the determination of trace elements by XRF analysis. Considering the capabilities of modern XRF equipment and the recent improvements in the theory of fusion and its technology, this should no longer be the case. The main reasons for reluctance in using fusion beads are: – – –

the lower ﬂuorescence intensities due to the dilution of sample; the higher background due to scattered radiation from the light elements of the diluent; occasionally, the difﬁculties in undertaking fusions. The objectives of this chapter are:

– – –

to describe the fusion procedure; to introduce recent developments that ensure more successful fusions; to suggest means to overcome the reluctance in using fusion beads.

9.2 9.2.1

FUSION IN LITHIUM BORATES General

Fusion is a chemical procedure in which a sample is transformed into a glass bead that has the required qualities for high accuracy in XRF analysis, i.e., be homogeneous near the atomic level, ﬂat, polished and inﬁnitely thick with regard to X-Ray penetration. The procedure is so simple that fusion was Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

301

F. Claisse

satisfactorily done manually for about 15 years before automatic instruments became available in the early 1970s [1 –3]. 9.2.1.1 The procedure Essentially, a fusion procedure consists of four operations: – – – –

mixing the sample with a ﬂux, and occasionally with an oxidant, in a crucible; heating the crucible at a temperature above which the ﬂux melts; agitating the crucible until the sample has dissolved in the molten ﬂux; pouring the molten glass into a hot mould so as to obtain a ﬂat glass bead of a given diameter after cooling, ready for measurement in the spectrometer.

Alternatively, the molten glass can be poured into a swirling warm acid solvent near ambient for dissolution. 9.2.1.2 The ﬂux The ﬂuxes mostly used are borates of lithium. They contain only light elements, lithium, boron and oxygen, that do not interfere with X-ray emission lines of other elements and do not absorb their radiations signiﬁcantly. The ﬂux composition may vary between lithium tetraborate Li2B4O7 (LiT) and lithium metaborate LiBO2 (LiM). It is obtained either by mixing together different proportions of those two stoichiometric compounds, or directly during manufacturing. Fluxes of several popular compositions and different purities are available commercially. Fused ﬂuxes are better than mixed ﬂuxes because they are not subject to segregation. High purity may or may not be important, but it should be borne in mind that any impurity in the ﬂux is observed as being an impurity in the sample, at a concentration equal to that in the ﬂux multiplied by the factor: ﬂux/sample. For example, in a fusion bead made from 1 g sample and 10 g ﬂux, a 2 ppm impurity in the ﬂux appears as 20 ppm in the sample. Water content must also be taken into account. Fluffy ﬂuxes can absorb water in a short time when exposed to air, and may already contain a few percents of water when received. This can affect the analytical accuracy. Water may be expelled almost explosively if heating is applied too rapidly, covering the surroundings with ﬁne powders of both ﬂux and sample. They cannot be recommended for accurate analyses. The weight of ﬂux needed for a fusion bead is of the order of 5 –8 g. For a solution, LiM is recommended for the ﬂux, on account of its better solubility for most oxides and the weight should not exceed 2 g. 9.2.1.3 The sample Oxides can only be dissolved in borate ﬂuxes; other materials must be oxidized separately, or at the beginning of fusion.

302

Fusion and ﬂuxes

Sulﬁde ores or concentrates can easily be oxidized using an additional step in the fusion procedure. A sample mixture is prepared in the crucible: typically, 0.4 –1 g of sample, 1–1.5 g LiNO3, 1 g of ﬂux and a few drops of nitric acid are suggested. The mixture is covered with the rest of the ﬂux (5 or 6 g); then heating should start slowly for about 2 min, and continued as outlined below for oxides. A procedure quite similar to that used for sulﬁdes has been developed for metals. However, some corrosion of the crucible may result; therefore, a wet chemical oxidation in the crucible prior to fusion, at near ambient temperature, is preferable. The ﬂux is then added and the fusion is made. Apart from usual acids, a reaction with hydrobromic acid followed by the removal of bromine by the addition of lithium carbonate can be tried. The weight of sample for a fusion bead may be as low as 100 mg (exceptional) and as high as 2 g or more; generally the ﬂux/sample weight ratio does not exceed 20, but it may be as low as 2, depending on the difﬁculty to fuse or the cooling procedure [3] (Section 9.2.1.8). For a solution, the sample weight may reach 1 g. The sample should be representative of the material to analyse, ground to less than about 100 mm, and well mixed with a fraction of the ﬂux, to avoid forming aggregates during fusion. 9.2.1.4 Crucibles and moulds Only precious metals can be used for fusion. Most crucibles and moulds are made of Pt –5%Au alloy. This material is supposed to be non-wetted by the molten glass. That is nearly true when new, but as the number of fusions increases, a non-wetting agent (NWA) (releasing agent) becomes more and more necessary to prevent glass residues in the crucible after pouring, and to release the glass bead from the mould after cooling. The recommended size is 30 ml for the crucibles, and 32 mm diameter for the moulds. Moulds larger than 35 mm lose their shape rapidly unless they are at least 1 mm thick. Deformations in the bottom of moulds will affect the analytical accuracy by as much as 1%. For maximal accuracy, the same mould should be used for standards (calibration) and samples (analysis). 9.2.1.5 The non-wetting agent Except on rare occasions, NWAs must be used. Only halides have that property, and only the heavier ones, bromine and iodine, are efﬁcient for this purpose. The nature of the cation does not appear important, except that lithium and ammonium are the only ones that do not contaminate the fusion. The author [5] interprets the behaviour of these elements as follows. Oxides are soluble in Li borates because the oxygen atoms do not disturb the boron– oxygen bonds that constitute the structure of borate glasses; the positive atoms contend with the interstitial positions in the lattice of the glass. Iodine and bromine are unable to be incorporated into molten Li borates because they are

303

F. Claisse

large negative atoms that cannot replace oxygen atoms; that would disturb the strong boron–oxygen bonds. They are soluble only when they are combined to oxygen as iodates, in which case they are small positive interstitial atoms, just like cations of all other soluble oxides. Consequently, most of those atoms stay at the surface of the melt while their cation penetrates inside. Without NWA, there is no well-deﬁned physical separation between the oxygen atoms of the protective layer on the platinum of the crucible (or mould), and the oxygen atoms in the molten glass; therefore some bonding between them may exist. An NWA that forms a thin layer of bromine or iodine between them prevents such bonding, because those atoms remain strongly bonded to their cations inside the melt and not to the oxygen on the platinum. NWAs are highly volatile, and their volatility increases with the atomic weight. Experiments indicate that about 50% of the original bromine and 90% of the iodine is lost after a 10-min fusion. Bromine from the NWA has been observed to be uniformly dissolved in the fusion disks, which suggests that some of it on the surface may enter the bead after oxygen from the atmosphere has formed a soluble bromate. Alternatively, when atmospheric oxygen reacts with the dissolved cation, Br is free to evaporate as Br2 molecules. The usual quantity of NWA to use in a fusion for production of fusion beads is of the order of 10–30 mg. Using more, may affect the analytical accuracy, because the halide ﬁlm is an efﬁcient absorber of both the incident and emergent X-rays. There is no objection to use of more than 30 mg in the case of the determination of trace elements, because the resulting error on low concentrations would be acceptable. For making solutions, an excess of NWA must be used to ensure that none of the usually small quantity of sample remains in the crucible; 100 mg is recommended. 9.2.1.6 Melting and homogenization All means of heating may be used, provided the atmosphere in the crucible is not reducing. Furnace heating is slow. Propane gas heating is fast, and is slightly oxidant. High frequency heating is also fast, but only one sample at a time can be processed. Electric heating of an individual crucible can be nearly as fast as propane heating. Continuous agitation for homogenization is preferable, but occasional manual shaking is also acceptable. The optimal fusion temperature is just enough to melt the ﬂux. Since the process is not the fusion of the sample but its dissolution in the molten ﬂux, a temperature not exceeding 950–1000C is the best to minimize the evaporation of the NWA, the ﬂux, or volatile elements such as sulfur (SO3) and ﬂuorine. The fusion temperature can even be lowered after melting, without risk of crystallization, but that is not necessary except for retaining ﬂuorine or chlorine. The quality of fusion beads is not affected signiﬁcantly by excessive temperature differences between fusions, but the analytical accuracy is affected by differences in the residual amount of NWA present as a surface ﬁlm.

304

Fusion and ﬂuxes

A typical fusion time is 5–6 min with continuous agitation of the crucible. For making solutions, 2 min heating is often sufﬁcient. 9.2.1.7 Pouring into moulds or beakers When the molten glass is completely homogeneous, it is poured into a hot mould in order to make fusion beads. The mould should be at a temperature not exceeding that of the crucible, preferably at 50 –1008C lower, and protected from drafts. For making solutions, the temperature of the acid solvent in the beaker must be just warm, and not exceed about 408C. The temperature of the crucible must generally be lowered before pouring the molten glass into the beaker. If the pouring temperature is too high, the drop of molten glass will pass completely through the solvent, hit the bottom of the beaker, damage it, or break it if made of glass. If the pouring temperature is too low, the drop of glass may separate into just a few particles that may take hours to dissolve. When the pouring temperature is just right, the drop of molten glass shatters into very ﬁne particles that usually dissolve in less than 5 min in the swirling warm solvent. The right range of temperatures depends on the solvent temperature, and must be determined experimentally. The ﬁnal solution is normally clear. When iodine is used as NWA, the solution has an orange-brown colour. 9.2.1.8 Cooling the fusion beads The mould should be level and held so as to cool as evenly as possible, with a minimum of contact with heat conductors. When the optimal ﬂux composition is used (Section 9.3.2), cooling is done in two steps: a draft-free air cooling for 1 – 2 min, followed by forced air cooling (fan) for another 2 min. This procedure might have to be modiﬁed when a different ﬂux is used. The ﬁnished bead can be picked up by means of a suction cup. It should be measured as is, without polishing. 9.2.1.9 Fusion instruments Despite the simplicity of a procedure consisting of melting, casting and cooling, the perfection of fusion beads depends on quite small things: imperfections in moulds, fusion mixture, unequal evaporation of NWA and instrumentation. Automatic ﬂuxers allow for reproducible operating conditions, but those conditions must strictly satisfy the requirements for making good beads. 9.3 9.3.1

THE KEY TO SUCCESSFUL FUSION BEADS The concept of “neutrality”

In science, neutrality appears to convey stability: atoms are made of positive and negative charges; simple chemical compounds contain anions and cations; magnetic materials tend to have magnetic domains of opposite polarities.

305

F. Claisse

Apparently, a similar concept of neutrality exists for fusion, which aids in understanding many observations. In turn, it can be used to choose the best conditions for successful fusions. The following three sections illustrate how the neutrality concept is utilized. 9.3.2

The optimal ﬂux and crystallization

In fusion, the neutrality concept may be expressed as: to be stable, a glass must be neutral with respect to the acidity of its components. In other words, the better a sample and a ﬂux match their acidic and basic components, the easier it is to produce a stable glass bead. In the early days of fusion with lithium borates, mention was made that a basic ﬂux should be used with an acidic sample, and vice versa, but since the basic ﬂux (LiM) yielded crystallized beads most of the time, most analysts were content with use of LiT for all types of samples. The original fusion technique [1] used sodium tetraborate because sodium was not detectable by X-rays; later, the adoption of LiT by a majority of practitioners seemed to be a logical choice. A detailed study of the effect of acidity on the quality of fusion beads was made by Claisse [6]. The maximum quantity of any oxide that can be dissolved successfully in a given quantity of ﬂux, called the solubility, was measured as a function of the fractions of LiT and LiM in the ﬂux. The term solubility meant no cloudiness, no undissolved particle, and no crystallization in the fusion bead. It was found that the solubility varies continuously with the composition of the ﬂux, between LiT and LiM (Fig. 9.1). All oxides MexOy with y=x # 1 (basic oxides), such as CaO, are more soluble in LiT (acidic ﬂux with y=x . 1); the solubility decreases as the ﬂux composition becomes more basic. By contrast, the solubility of all oxides with y=x . 1 (acidic oxides), such as SiO2 and Fe2O3, increases as the composition of the ﬂux gets closer to LiM

Fig. 9.1. Solubility of CaO, SiO2 and Fe2O3 in Li borate beads.

306

Fusion and ﬂuxes

(basic ﬂux with y=x ¼ 1). The difference between SiO2 and Fe2O3 above 80% LiM, the eutectic point of the LiT–LiM system, is due to the difference in size of the two cations. It was shown that the solubility increases as the size of atoms decreases [5]. In SiO2, crystallization is observed at low concentrations as in Fe2O3; it then decreases as the concentration of SiO2 increases, because the larger number of dissolved Si atoms impede the nucleation of crystals. The solubility curves for complex oxides are more or less similar to the sum of the curves of its components. An example is that of CaSO4 which is equivalent to CaO þ SO3 after dissolution in the ﬂux. Measurements made on acidic oxides that behave like Fe2O3 include: TiO2, Sb2O3, V2O5 and WO3. Measurements made on basic oxides that behave like CaO are MgO and BaO. The two divalent metallic oxides PbO and ZnO also behave like basic oxides. Measurements made on acidic oxides that behave like SiO2 are Al2O3, SO3 and P2O5. Acknowledgement is made of the 12 LiT:22 LiM mixture developed by Norrish and Thompson [7] for alumino-silicates, and the 20 LiT:80 LiM mixture of Bennett and Oliver [8], but in both cases one of the reasons evoked for success was the greater facility to fuse, on account of the lower melting point of the ﬂux near the eutectic point of the Li2O–B2O3 system. In practice, two ﬂux compositions are sufﬁcient only to process a wide variety of samples: LiT for essentially monovalent and divalent oxides, and one composition in the range 50–60% LiM with LiT as the rest, for all other oxides. 9.3.3

Cracking of fused beads

An additional observation by the author [5] that conﬁrms the tendency towards neutrality in lithium borate glasses is cracking. In fusion beads containing oxides of transition metals in a basic ﬂux (near LiM), the metal atoms should tend to be acidic, or take the maximum oxidation state; i.e., as a certain equilibrium between oxidation states exists, the maximum oxidation state is more favourable than the other. For example, between FeO and Fe2O3, the latter is favoured in a basic ﬂux, i.e., conﬁrmed at times by the colour of the disk. In an acidic ﬂux (near LiT), FeO is the favoured oxidation state. In such case, since a possible equilibrium between Fe and FeO exists, some atoms might be present as Fe. Close to the platinum surface, once in a while one such free Fe atom may permanently bond to the platinum. During fusion, the platinum crucible is gradually plated with Fe and becomes an iron crucible. The molten glass then adheres to the crucible, and after pouring into the mould, a residue remains in the crucible. The same situation arises in the mould: the glass sticks to it, and during cooling the mould shrinks while the glass does not, resulting in the glass bead cracking. NWA are rather inefﬁcient in such cases, because halides are acidic atoms in an acidic environment, which enhances the evaporation rate. This is

307

F. Claisse

conﬁrmed by the fact that all transition metals stick to the mould when fused in LiT, regardless of the presence of an NWA, and beads usually crack. Cracking is never observed with acidic oxides in the same situation. 9.3.4

Loss and retention of sulfur

Sulﬁdes do not dissolve in ﬂux because the negative sulfur atoms that should normally replace the negative oxygen atoms in the boron–oxygen lattice apparently are not accepted. However, a sulfate is soluble because it is the result of the combination of two oxides, a metal oxide and a sulfur oxide SO3. As a result, SO3 is an acidic oxide ðy=x ¼ 3Þ and is retained in a basic ﬂux (LiM). As described in the preceding section, SO3 becomes SO2 if the ﬂux is acidic, and, being a gas, it is lost. 9.4 9.4.1

APPLICATION TO TRACE ELEMENT ANALYSIS Maximizing X-ray intensities

Dilution of the sample by the ﬂux unfortunately decreases X-ray emission intensities of elements, but some compensation is possible. One way to maximize X-ray intensities is to increase the concentration of sample in the ﬂux by carefully choosing the most appropriate composition of the ﬂux. Dilution is often not as serious as may be imagined. An example is shown in Fig. 9.2. X-ray Ka line intensities of a mixture of 25% each of four oxides were calculated theoretically with a Fundamental Parameter program, Rhodium excitation;

Fig. 9.2. Calculated Ka line intensities of a four-oxide specimen as a function of the dilution in 50 LiT:50 LiM.

308

Fusion and ﬂuxes

ﬁrstly as a non-diluted sample, then after increasing dilution in 50 LiT:50 LiM. It is seen that the decrease in intensities is far less than in proportion to the dilution, and is not large if the diluent weight is not more than three times the sample weight, and if the atomic number of the element is above 18. Under such conditions, the loss is less than 20% for Fe2O3, still less for heavier elements, and only 50% for K2O. An additional means of increasing the intensities, surprisingly, is to add another diluent that increases the solubility of the sample. This possibility is suggested by the results of a few recent observations by the author [5], and is promising. A clear case is that of CaO. The maximum solubility of that oxide is 0.3 g per gram of LiT (Fig. 9.1). The solubility of Portland cement containing about 65% CaO is about 1.2 g per gram of LiT, which is equivalent to 0.8 g CaO per gram of LiT. The increase in solubility is explained by the presence of about 25% SiO2 (an acidic oxide) in the cement sample. A similar case is that of Na2O (a basic oxide) which increases the solubility of SiO2 in LiT. A third way to maintain intensities is to manage the cooling rate. It should be reminded that the term solubility used in this chapter is the solubility in the solid glass after given cooling conditions. The observed solubility depends on both the solubility in the molten state, and the sluggishness of the ﬁrst crystal to form during cooling. Thus, increasing the fusion temperature normally increases the solubility in the melt, and a high cooling rate may prevent nucleation from taking place. Eastell and Willis [4] described a fast cooling technique. 9.4.2

Minimizing background

Dilution with light elements (ﬂuxes) results in stronger scattered radiation and increases the background. The most efﬁcient way to achieve low background is to work with the most concentrated beads possible, i.e., the means are the same as for maximizing X-ray intensities. Measuring X-ray intensities under a particular set of conditions can also reduce background, but that subject is not within the framework of the present chapter. 9.5

FEATURES OF FUSION FOR TRACE ELEMENTS

As compared to pressed powders, fusion has some disadvantages, namely lower intensities and higher background, but that is compensated by several major advantages: –

–

Accuracy: a fusion bead specimen being perfectly homogeneous, the application of matrix correction algorithms or softwares yields high analytical accuracy, comparable to that of major elements. Trace elements can be determined accurately, without measuring the major elements if their approximate concentrations are known. An

309

F. Claisse TABLE 9.1 Quantitative determination of trace elements. Effect of approximation in concentrations of major elements. Relative standard deviations are from six determinations Oxide

Average concentration (ppm)

Na2O MgO Al2O3 SiO2 P2O5 CaO TiO2 Cr2O3 Mn3O4 Fe2O3

891 503,406 147,781 29,547 132 11,166 1317 170,268 1080 105,319

–

–

– –

Standard deviation on trace elements (ppm) 6

3 9 10

310

0.7 5.4 8.0 6.2 2.1 5.5 0.7 8.0 0.9 9.4

example is given in Table 9.1: the X-ray intensities of each element of a known standard (BCS 369) were measured; each intensity of the major elements only was multiplied by a random number between 0.88 and 1.12 to simulate a statistical error of about 8% on their concentrations; the process was repeated six times to simulate six different estimates of the concentrations of major elements. It is seen that the effect on the absolute error, in ppm, of the trace element concentration is always small, and the relative error is about 10-fold smaller than those of the major elements. Making standards for the analysis of trace elements is easy: a fusion bead with one or more pure oxides with accurately measured concentrations is made; then, a small piece of that bead is mixed with ﬂux and a more diluted bead is made with accurately known concentrations. Cost to make fusion beads with fusion instruments is somewhat higher than to make pressed pellets, but is lower when more than one pellet is required to ascertain the results. Time to make fusion beads with a multi-sample instrument is less than making pressed pellets manually. Background intensities are increased due to the scattered radiation. On the other hand, the background in glass specimens is more reliable because the specimen does not emit diffraction lines.

REFERENCES 1 2

Relative standard deviation (%)

F. Claisse, Norelco Rep., 4 (1957) 95. W.B. Stern, X-Ray Spectrom., 5 (1976) 56.

Fusion and ﬂuxes 3 4 5 6 7 8

M.T. Haukka and I.L. Thomas, X-Ray Spectrom., 9 (1977) 204. J. Eastell and J.P. Willis, X-Ray Spectrom., 19 (1990) 3. F. Claisse, unpublished notes. F. Claisse, J. Phys. IV France, 8 (1998) Pr5-379. K. Norrish and G.N. Thompson, X-Ray Spectrom., 19 (1990) 67. H. Bennett and G.J. Oliver, Analyst, 101 (1976) 803.

FURTHER READING V. Buhrke, R. Jenkins and D. Smith, A Practical Guide for the Preparation of Specimens for X-Ray Florescence and X-Ray Diffraction. John Wiley & Sons, 1998. H. Bennett and C.J. Oliver, XRF Analysis of Ceramics, Minerals, and Allied Materials. John Wiley & Sons, 1992.

311

This page intentionally left blank

Chapter 10

Supercritical ﬂuid extraction R. Alzaga, S. Dı´ez and J.M. Bayona

10.1

PROPERTIES OF SUPERCRITICAL FLUIDS

Supercritical ﬂuids (SFs) are gases at pressures and temperatures above their critical point. Accordingly, by increasing the pressure, SFs are not able to liquefy, irrespective of pressure or temperature, thus behaving as a singlephase system. However, for some applications, subcritical or near-critical ﬂuids may be advantageous in respect to SFs. SFs have intermediate properties between gases and liquids and they depend on pressure. At high pressures, they behave as a solvent-like ﬂuid but at pressures near the critical point they resemble a gas. Moreover, the ﬂuid density, and consequently its solubility, can be adjusted by modifying the pressure and temperature (Fig. 10.1). Consequently, a number of advantages accrue when they are used in extraction processes or in chromatography because of their lower viscosity, higher diffusion coefﬁcient and lower surface tension compared to liquids (Table 10.1). While the low viscosity leads to favorable dynamic properties of enhanced mass transfer, the compressibility allows a density change with pressure and, therefore, its solubility can be adjusted during extraction to increase the selectivity. Furthermore, the higher diffusion coefﬁcient enhances the transport properties and the low surface tension improves the penetration of the extracting ﬂuid into the matrix and enhances the extraction kinetics. Another important feature of SFs is the change of the solubility parameter, dSF, with pressure and temperature; in case of liquids, it remains constant because of their lack of compressibility, i.e., 1=2

dSF ¼ 1:25Pc ðrf =rl

10:1Þ

where Pc is the critical pressure of the solvent, rf is the reduced density of the SF and rl is the reduced density of the SF in the liquid state. Therefore, dSF is proportional to the density of the SF. This property is used in capillary supercritical ﬂuid chromatography (SFC) where an increase in density arising from increasing pressure is utilized to perform separations [1]. Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

313

R. Alzaga et al.

Fig. 10.1. Effect of pressure and temperature on the solubility of a model solute in an SF.

Temperature plays an important role in supercritical ﬂuid extraction (SFE). An increase in the extraction temperature at constant pressure leads to a decrease in density and, therefore, of solubility. However, in the case of analytes having a substantial vapor pressure, an increase in temperature leads to an increase in solubility. Temperature is also relevant to mitigate the matrix effects because an increase in temperature usually decreases analyte –matrix interactions. Although a number of SFs can be used for extraction purposes, for practical reasons only carbon dioxide has been widely used in most analytical SFE procedures. The main advantage of carbon dioxide is its accessible critical constants (Tc ¼ 318C, Pc ¼ 74 bar), its inertness, non-corrosive nature, low cost and toxicity (threshold limit value, TLV ¼ 5000 ppm), high purity and relatively high solubility parameter (dSF < 7 at 250 bar), which was thought to be close to pyridine [2]. However, more recent studies suggest that this value is inﬂated and, in line with experimental evidence, CO2 can solubilize only low molecular weight materials making the use of modiﬁers unavoidable in most extraction processes [3]. For technical applications, a variety of CO2-philic TABLE 10.1 The solubility, S, viscosity, h and diffusion coefﬁcient, D, for naphthalene in carbon dioxide under gas, liquid and supercritical conditions State

S (kg m23)

h (m Pa s)

D (m2 s21)

Gas (408C, 1 bar) Supercritical (408C, 100 bar) Liquid (238C, 500 bar)

2 632 1029

16 17 133

5.11026 1.41028 8.71029

314

Supercritical ﬂuid extraction

materials have already been developed in order to increase their solubility at lowest pressures, such as ﬂuorinated polymers [4]. According to Hawthorne [5] the extraction process can be considered to involve the three factors shown in the following SFE triangle:

First, the solute must be sufﬁciently soluble in the SF. If the solubility is not sufﬁcient, the addition of an organic modiﬁer or a complexing agent will improve the extraction yield. Second, solutes must be transported rapidly by diffusion from the interior of the matrix pores or surface and it is controlled by the diffusion coefﬁcient. This parameter is very important because extraction time depends on its square. Third, solutes must be released from the matrix by desorption. This step is slow because analyte removal is rather difﬁcult, depending on the matrix characteristics and strongly on pressure; their inﬂuence can be reduced by adding polar modiﬁers. Finally, trapping of the recovered extract by SF decompression is of primary importance, and the efﬁciency of the collection system depends on the analyte volatility. Several systems have been developed in order to accomplish this purpose (see Section 10.2). Although early studies discarded the use of SFs in the extraction process, different strategies have been developed for the metal ions and organometallic species extraction from different matrices. These strategies include either a derivatization, to transform organometallic ionic species into a non-ionic compound readily soluble in the ﬂuid, or the formation of stable chelates or complexes by adding a suitable ligand. However, as noted earlier, a requirement for an effective extraction is the solubility of the chelate or the derivatized organometallic in the ﬂuid. This can be measured experimentally (see Section 10.2) or estimated from chromatographic retention data. The latter method is much faster and can be tested under a variety of experimental conditions. The method relies on the assumption that an inverse relationship exists between the chromatographic retention obtained by SFC and the solubility in the mobile phase [6]. The degree of retention of a solute in SFC, as measured by the capacity factor, k0 ; is at least qualitatively inversely related to the solvation power of the mobile phase for that solute: the more soluble it is in the mobile phase, the less it will be retained. The relationship used is: S ¼ C=k0

ð10:2Þ

where S is the solubility (per unit volume), k0 is the chromatographic capacity factor deﬁned in terms of the retention volume, VR and the void volume of the

315

R. Alzaga et al.

column and the connecting tubes, VM by: k0 ¼ ðV R 2 V M Þ=V M

ð10:3Þ

and C is a constant for a particular column, solute and temperature given by: C ¼ Aexp ½B=RT

10:4Þ

In Eq. (10.4), Aexp and B are constants for a particular column and temperature. The validity of these equations has been tested with ferrocene and a nickel complex by making a small number of direct measurements [6]. The precision of the results is estimated to vary from 5% at low pressures to 20% at higher pressures, where high solubilities and low retention times give rise to higher experimental errors. 10.2

INSTRUMENTATION

An analytical SFE system consists of a source of ﬂuid (i.e. CO2), a pumping system (which delivers a ﬂuid at constant pressure), and a heated extraction chamber, where the sample vessel is located. An outlet valve (an on/off valve) allows for the isolation of the extraction system under SF conditions without any ﬂow through the extraction cell and is known as the static mode. By activation of the former valve, the ﬂuid percolates through the extraction cell and ﬂushes the extracted sample to the collection system, which is termed the dynamic mode. A ﬂow restrictor or other backpressure regulation device inserted between the on–off valve and the extract collection device keeps the system pressurized. Optionally, a modiﬁer pump is used, which delivers an organic solvent (modiﬁer) to the SF (see Fig. 10.2). Typical modiﬁers include dichloromethane, methanol, ethanol, hexane, etc. Fused silica capillaries are the most used ﬂow restrictors in SFE. These restrictors are fabricated from appropriate lengths of capillary tubing of small internal diameter (i.e., 50 mm). The ﬂow rate is dependent upon the internal diameter and the length of the restrictor. Unfortunately, fused silica restrictors become brittle with the use of certain modiﬁed ﬂuids, such as methanol or chelating agents. This results in a short-lived restrictor, which necessitates frequent replacement. Outlet valves and restrictors are prone to plugging, which results from loss in analyte solubility as the density of the SF drops during its decompression. Cooling due to the Joule –Thompson effect as the ﬂuid expands can further exacerbate this plugging. Although heating diminishes this problem, it is sometimes difﬁcult to heat thoroughly and consistently this part of the SFE system. In order to circumvent this drawback, variable restrictors have been developed, allowing a constant ﬂow independent of the extraction pressure. Off-line extraction. In this extraction procedure, a few grams or milliliters of sample are extracted and the supercritical carbon dioxide (SF-CO2) is decompressed within a solvent. Because the solubility of the analytes decreases

316

Supercritical ﬂuid extraction

Fig. 10.2. SFE instrumentation.

rapidly with dropping pressure, the analytes are partitioned into the solvent. The trapped extract can then be analyzed using various analytical techniques. The main advantage of this extraction procedure is its high ﬂexibility because the extract can be cleaned prior to analysis or submitted to different analytical techniques. Several collection systems have been used in SFE depending on the physical– chemical properties of the analyte. Solvent trapping is wide because of its simplicity, provided the analyte is not volatile which is the case of metal ions or organometallic species. Another trapping approach used off-line is to decompress the carbon dioxide in an appropriate support (i.e., glass or stainless steel beads) or sorbent (i.e., silica, Florisil, C18), usually at subambient temperature when eluted with a solvent. On-line extraction. In this extraction procedure, a few milligrams of sample are extracted, the extract is trapped and transferred to a detection system, such as UV, FT-IR, atomic absorption spectrometry (AAS) [7] or further processed using separation methods (i.e., GC or SFC) by use of an appropriate interface. The main disadvantage compared to off-line techniques is the risk of contamination of the chromatographic system because no cleanup steps are undertaken, memory effects in the case of incomplete desorption and small amount of sample is needed (ca. mg range), which demands high sample homogeneity. However, when an extraction is coupled to the detection system without chromatography, the interface is less sophisticated because no cryogenic trapping is necessary; this is particularly useful for method

317

R. Alzaga et al.

development. Detection limits at the picomol level have been reported in the determination of As, Cd, Cu, Mn, Pb, Se or Zn by AAS. 10.2.1 Experimental solubility measurements Different approaches have been developed in order to determine ligand or metal complex solubility in SF-CO2. In situ spectroscopic determinations incorporated into the extraction cell are achieved using UV–visible measurements with a ﬁber-optic system. This approach allows rapid determination and requires small amounts of analyte. However, the cell design (which requires a precision-engineered view cell), path lengths and clean window protocols are critical during solubility determinations [8]. Alternatively, FT-IR spectroscopy has also been used for the determination of ligand –metal complexes [9]. The spectrometer was ﬁtted with a high-pressure stainless steel reﬂectance cell. The resulting metal spectra were subtracted from that for pure CO2 obtained at the same pressure and temperature. Off-line determinations include gravimetric, spectroscopic or chromatographic methods. These methods are time consuming and exhibit a lack of sensitivity, especially for low-solubility analytes compared to solubility determinations made on line. Taylor et al. [10] developed a system with a recirculating pump, which isolates and maintains the extraction cell at constant extraction conditions. The function of this pump is to ensure complete saturation of SF with the ligand/chelate of interest. After equilibration (i.e., 30 min) an aliquot of the ﬂuid is analyzed by LC –UV. 10.3

SFE OF TRACE ELEMENTS

Direct extraction of metals ions by SF-CO2 is inefﬁcient because of the weak van der Waals interactions between metal ions and the non-polar carbon dioxide. This weak interaction has apparently discouraged efforts to perform SFE of metals from different matrices. Initially, metal extraction using SFs has been considered not to be feasible owing to the charge neutralization requirements. By addition of a complexing agent (ligand) to the SF phase, the charge on the metal ion can be neutralized and then lipophilic groups are introduced, forming the metal–complex system (Table 10.2). Solubilization of the metal complexes into the SF is then possible, and extraction and separation can then be accomplished. Rapid complexation kinetics between metal ion– ligand and a high stability constant for the resulting neutral complex can obviously enhance the extraction process. Typical chelating agents used in SFE include dithiocarbamates, b-diketones, organophosphorus reagents, hydroxamic acid, crown ethers containing O, N, P, and S functional groups and ligands having ﬂuorine substituents (Tables 10.2 and 10.3). Their structure is shown in Fig. 10.3.

318

Supercritical ﬂuid extraction TABLE 10.2 Ligands and chelating agents used in SFE of metals Ligand

Abbreviation

Ionizable-crown ethers Dicyclohexylbistriazolo-crown ether Dibenzobistriazolo-crown ether Tert-butyl-substituted dibenzobistriazolo-crown ether

Crown I Crown II Crown III

b-Diketones (non-ﬂuorinated/ﬂuorinated) Acetylacetone Hexaﬂuoroacetylacetone

AcAc HFA FOD

2,20 -Dimethyl-6,6,7,7,8,8,8-heptaﬂuoro-3,5-octanedione Triﬂuoroacetylacetone Thenyltriﬂuoroacetone 2,2,6,6-tetramethyl-3,5-heptanedione Benzylacetonate

TFA TTA THAc BzAc

Dithiocarbamates (non-ﬂuorinated/ﬂuorinated) Diethyldithiocarbamate Bis(triﬂuoroethyl)dithiocarbamate Dibutyldithiocarbamate Pyrrolidinedithiocarbamate

DDC FDDC DBDTC PDTC

Organophosphorus reagents Triphenylphosphine oxide Tributylphosphate Tributylphosphine oxide Trioctylphosphine oxide Bis(2,4,4-trimethylpentyl)phosphinic acid Bis(2,4,4-trimethylpentyl)monophosphinic acid Bis(2,4,4-trimethylpentyl)dithiophosphinic acid Bis(2-ethylhexyl)monothiophosphoric acid

TPPO TBP TBPO TOPO Cyanex 272 Cyanex 302 Cyanex 301 D2EHTPA

Fluorinated hydroxamic acid Perﬂuorooctanohydroxamic acid N-methylperﬂuorooctanohydroxamic acid Heptaﬂuorobutyrichydroxamic acid N-methylheptaﬂuorobutyrichydroxamic acid

PFOHA MPFOHA HFBHA MHFBHA

Others 7-(1-vinyl-3,3,5,5-tetramethylhexyl)-8-hydroxyquinolin

Chelex 100

319

320

TABLE 10.3 Conditions used for the extraction of trace elements Metal

Matrix

Modiﬁer (ligand)

Extraction conditions

Reference

Cu2þ

Ceolite and aqueous solution

FDDC

[57]

La3þ, Eu3þ, Lu3þ

Cellulose-based ﬁlter paper (spiked, wet, pH ¼ 6.5)

FOD

Hg2þ

Cellulose-based ﬁlter paper (wet, spiked)

FDDC

(UO2)2þ

Cellulose-based ﬁlter paper (spiked), wet (10 mL water, pH ¼ 1.0 and 6.5) Freeze-dried bovine liver (NIST SRM 1557a) in 10% (w/v) KOH in 20% (v/v) methanol/water Cellulose-based ﬁlter paper and sand (spiked), topsoil, water (spiked), mine-water (pH ¼ 3.5, buffered)

FOD

CO2, T: 35–458C, P: 80 bar, S: 50 min (St), R: 45–99 CO2 þ MeOH (5 mol%), T: 608C, P: 150 bar, S: 10 min, D: 10 min R: 91–99% CO2 þ MeOH (5 mol%), T: 508C, P: 100 bar, S: 20 min, D: 10 min, R: .99% CO2 þ MeOH (5 mol%), T: 608C, P: 150 bar, S: 10 min, D: 10 min, R: 99% CO2, T: 508C, P: 240.5 bar, S: 20 min, D: 30 min, R: 91–94%

Th4þ, (UO2)2þ

TBA-DBDTC

FOD, TTA, HFA, TFA, AcAc, TBP

CO2, T: 608C, P: 150 bar, S: 10 min, D: 10 min, RSp: .95%, RN: 77–91%

[53]

[18]

[7]

[27]

R. Alzaga et al.

Zn2þ, Cd2þ, Cu2þ

[18]

TABLE 10.3 (continuation) Matrix

Modiﬁer (ligand)

Extraction conditions

Reference

La3þ, Eu3þ, Lu3þ

Cellulose-based ﬁlter paper and sand (wet, spiked), Water (spiked, pH ¼ 4 buffered) Nitric acid solution

HFA, TTA, FOD and/or TBP

CO2 þ MeOH (5 mol%), T: 608C, P: 150 bar, S: 10 min, D: 10 min, RSS: 92–98%, RWS: 75– 89%

[25]

TBP, TBPO, TOPO, TPPO

CO2, T: 608C, P: 200 bar, S: 15 min, D: 15 min, R: .96% CO2 þ MeOH (5 mol%), T: 608C, P: 200 bar, S: 10 min, D: 15 min, R: .96% CO2, T: 608C, P: 200 bar, S: 10 min, D: 20 min, R: .91– 95% CO2 þ methanol (5 mol%), T: 608C, P: 200 bar, S: 10 min, D: 15 min, R: 95– 98% CO2 þ ethanol (5% v/v), T: 608C, P: 200 bar, S: 10 min, D: 15 min, R: 93– 100% CO2, T: 40– 508C, P: 206 bar, S: 20 min, D: 60 min, R: 2 –13.9

[58]

Th4þ, (UO2)2þ

Zn2þ, Cd2þ, Pb2þ, Cu2þ, As3þ, Pd2þ Au3þ, Ga3þ, Sb3þ La3þ, Eu3þ, Lu3þ, Th4þ, (UO2)2þ

Cellulose-based ﬁlter paper and sand. (wet, spiked)

FDDC

Sand (wet, spiked)

TTA and TBP

Hg2þ

Cellulose-based ﬁlter paper and sand (wet, spiked)

Crown I, II, III

(UO2)2þ

Polypropylene/ polyester glasswool, cotton, kaolin (spiked)

FOD and TBP

Cd2þ, Cu2þ, Zn2þ

Bound to metallothionein from rabbit liver

TBA-DBDTC

[59]

[59]

[60]

[61]

[62]

continued

Supercritical ﬂuid extraction

Metal

321

322

TABLE 10.3 (continuation) Matrix

Modiﬁer (ligand)

Extraction conditions

Reference

Cellulose-based ﬁlter paper, soil and sand (wet, spiked)

FDDC, PDTC, DDC, TFA, HFA, TTA

[63]

Cu2þ, Pb2þ, Zn2þ, Cd2þ

Cellulose-based ﬁlter paper and bentonite clay (acid added, spiked)

Cyanex 272, Cyanex 302, Cyanex 301, D2EHTPA

Ni2þ, Cu2þ, Cr2þ

Water (spiked)

HFA, FDDC, DDC, AcAc

Pd2þ, Rh3þ

Sand and humic acid (spiked)

AcAc, THAc, BzAc

Fe3þ

Cellulose-based ﬁlter paper (wet, spiked)

PFOHA, HFBHA, MPFOHA, HFBHA

Cellulose-based ﬁlter paper (wet, spiked)

AcAc, HFA, DDC, FDDC

Co2þ

Filter paper–glass bead (wet, spiked)

HFA, AcAc

Pb2þ, Cu2þ, Ni2þ, Cr6þ, Zn2þ

Soil (wet)

HFA

CO2, T: 458C, P: 100– 250 bar, S: 15 min, D: 15 min, R: .4– 96% CO2, T: 608C, P: 200– 300 bar, S: 20 min, D: 20 min, R: .96% CO2, T: 608C, P: 200– 400 bar, S: 10 min, D: 30 min, R: .2– 82% CO2, T: 608C, P: 15 and 400 bar, R: 75–99% CO2, T: 708C, P: 300 bar, S: 20 min, D: 10 min, R: .98% CO2, T: 608C, P: 400 bar, S: 20 min, D: 20 min, R: .91% CO2, T: 608C, P: 400 bar, S: 20 min, D: 30 min, R: .95% CO2, T: 608C, P: 400 bar, S: 20 min, D: 30 min, R: .11 –30%

Metal 2þ

Cd

2þ

, Pb

2þ

, Hg

177

Lu(III)

[10]

[64]

[65]

[21]

[11]

[11]

R. Alzaga et al.

Co2þ, Pb2þ,

[17]

TABLE 10.3 (continuation) Metal

Matrix

Modiﬁer (ligand)

Extraction conditions

Reference

Aqueous solution (spiked)

Cyanex 302

[24]

Gd3þ

Metal oxides

TBP and HNO3

La3þ, Ce3þ, Nd3þ, Sm3þ, Eu3þ, Yb3þ

Aqueous solution (spiked, 1 M HNO3)

TBP, D2EHTPA

Pb2

Sodium alginate, blood, urine, human milk

DDC

Mo4þ

Irrigation water and pasture

AcAc

Cu2þ (as Cu(BzAc)2)

Diatomaceous earth (spiked)

BzAc

Cellulose-based ﬁlter paper and soil. (wet, spiked)

HFA, TBP

CO2, T: 408C, P: 83–138 bar, S: 20 min, D: 120 min (St), R: 49–60% CO2, T: 408C, P: 120 bar, S: 20 min, D: 90 min (St), R: 49–60% CO2, T: 408C, P: 250 bar, S: 90 min (St), R: 90% CO2 þ MeOH (50%), T: 1608C, P: 172 bar, S: 90 min, D: 2 min, R: .90% CO2 þ MeOH (30%), T: 1408C, P: 69 bar, S: 30 min, D: 3 min, R: 30% CO2 þ MeOH (5%v/v) þ Triton X-100, T: 608C, P: 250 bar, S: 10 min, D: 28 min, R: 90.5% CO2 þ MeOH (5% mol), T: 608C, P: 200 bar, S: 10 min, D: 15 min, R: 96–100%

Zn

2þ

177

Lu)

[66]

[33]

[33]

[32]

[23]

323

continued

Supercritical ﬂuid extraction

La3þ, Lu3þ (140La,

[30]

324

TABLE 10.3 (continuation) Matrix

Modiﬁer (ligand)

Extraction conditions

Reference

(UO2)

Contaminated soils (wet)

HFA and TOPO

[23]

Cu2þ, Pb2þ, Zn2þ, Cd2þ

Cellulose-based ﬁlter paper and sand (spiked)

Pb2þ, Zn2þ

Contaminated soil (wet/dry)

Kelex 100, Cyanex 301, Cyanex 302, Cyanex 272, D2EHTPA Cyanex 302

CO2 þ MeOH (5% mol), T: 608C, P: 200 bar, S: 10 min, D: 20 min, R: 72–88 CO2, T: 608C, P: 300 bar, S: 20 min, D: 20 min, R: 80–100% CO2, T: 608C, P: 400 bar, S: 20 min, D: 20 min, R: 40–84%

[20]

Metal 2þ

[20]

R. Alzaga et al.

T: extraction temperature; P: extraction pressure; S: static conditions; D: dynamic conditions; St: stirring; R: recovery; Sp: spiked; N: non-spiked; SS: solid sample; WS: water sample.

Supercritical ﬂuid extraction

325

Fig. 10.3. Ligand structure.

R. Alzaga et al.

Wide solubility ranges have been observed for metal complexes. The solvation is inﬂuenced by several parameters, including the pressure and temperature of SFs, modiﬁer effects in the SF, the ligand used, the pH, the identity of the metal and its oxidation state and the ligand functional group. 10.3.1 Ligand solubility in SFs The relevance of ligand solubility to metal extraction feasibility is obvious, in the way that a reasonable quantity of ligand must be solubilized to be able to access the metal ions in the various matrices. Ligand solubility depends on its structure and also on the extractant agent composition as well as pressure and temperature. Generally, solubility increases with SF density, and these extraction conditions can be achieved at high pressures and low temperatures, close to the critical point. This effect is shown in Table 10.3, where low temperature (i.e., 608C) is required for the extraction conditions. Moreover, temperature can affect the ligand itself. By increasing the temperature up to 1008C, using HFA as a complexing agent may increase the degradation of the ligand, which tends to decrease the extraction efﬁciency of the SFE process [11]. The solubility of ligands in SFs can be estimated by the group contribution method [12,13]. For instance, the solubility parameters (d2) of dithiocarbamate ligands (in their hydrogen form) with different alkyl substitutions were calculated using the group contribution method according to the following equation: X 1=2 .X d2 ¼ ð10:5Þ DUi DVi where Ui is the molar cohesive energy and Vi the molar volume of the ith group of a ligand molecule. Generally, an increase in chain length of the alkyl group of the dithiocarbamate ligands tends to lower the solubility parameter of the ligand [14]. The change in the solubility parameter value per carbon chain number is large when the carbon number is small and gradually levels off when the carbon number becomes greater than 10. Increasing the alkyl chain length substitution (i.e., dithiocarbamates) can increase the solubility of the ligand in SF. For organophosphorus ligands (Table 10.2) alkyl or aryl substitution dramatically affects the ligand solubility. The TOPO ligand (octyl substitution) shows solubilities approximately one order of magnitude higher than those of the TPP ligand (phenyl substitution). Ligand solubility is increased when ﬂuorine substitution is employed. The solubility of the ﬂuorine-substituted ligands is greater than that of the non-ﬂuorinated ligands. Increasing the amount of ﬂuorine substitution (low polarity) also increases the solubility in SF. Ligands can exhibit acidic –basic properties (i.e., b-diketones), and ﬂuorine substitution can signiﬁcantly increase the acidity of the ligand (i.e., AcAc pka ¼ 8:67; HFA pka ¼ 4:46). This reduction in the basic strength of the ﬂuorinated

326

Supercritical ﬂuid extraction

Fig. 10.4. Keto–enol equilibrium for the b-diketone ligand.

ligand can be advantageous to water (required in the SF process) in contact with the SF-CO2, which depends on temperature and pressure but is strongly acidic (e.g. pH ¼ 2.80–2.95) [15]. The b-diketone ligands are used almost exclusively in their protonated forms. The b-diketones are rendered acidic by the tautomeric equilibrium established between the enol and the keto forms (Fig. 10.4). The ﬂuorinated ligands have been found to be almost exclusively in the enol form under the high pressures and temperatures relevant to SFE. The non-ﬂuorinated ligand (e.g. AcAc) is found to be partly in each form in the SF phase. The acidity of the ligand originates from the enol form and the deprotonated b-diketones involved in the metal complexation process. It is well known that the presence of electron-withdrawing ﬂuorine substituents can signiﬁcantly increase the ligand acidity. In general, it appears that the most favorable complex for high SF solubility is a ﬂuorine-substituted ligand system. For analytical applications, the ﬂuorine-substituted system is clearly the most suitable. However, the high cost of ﬂuorine-substituted ligands will probably limit their use to the analytical scale. For hydrocarbon-based ligands, aliphatic-substituted systems show high solubilities, approaching those of ﬂuorine-substituted systems, and these may have potential for larger scale applications. Phenyl-substituted ligands show the lowest solubilities and are not likely to ﬁnd many SF applications. In general, the ligand solubility is increased signiﬁcantly when SF is modiﬁed by adding a low percentage (e.g. 5%) of an organic polar solvent (generally methanol). Determination of ligand solubility in SF-CO2 media is indispensable from both fundamental and practical viewpoints. First, for elucidating and formulating an extraction reaction in which the distribution equilibrium of ligand can be expressed by the ratio of ligand solubilities in both phases (when aqueous matrices are extracted), the distribution equilibrium of the ligand itself between aqueous and SF-CO2 phases should be taken into account. Secondly, solubility of ligand in SF-CO2 is, in general, substantially lower than that in a conventional organic solvent, which may restrict the preparation of SF-CO2 media containing the ligand at sufﬁciently high concentration for optimum extraction performance [16].

327

R. Alzaga et al.

10.3.2 Complex – SF solubility A primary requirement for the use of a ligand as an extractant in SF is that the ligand and the resulting metal complexes should be reasonably soluble in the ﬂuid phase. However, little information is available on this subject area, because metal chelate solubility depends on the metal ion, ligand and extraction conditions (i.e., temperature and pressure). For instance, DDC (see Table 10.2) is a widely used chelating agent for trace metal extraction, which forms stable complexes with over 40 metals and metalloids. Different metal–DDC chelates exhibit limited solubilities in SF-CO2, reﬂected by their poor SFC behavior. In parallel with the “free” ligands, complexed metal systems show the highest solubility for ﬂuorine, followed by aliphatics, and the lowest for phenyl-substituted systems [17]. Generally, the solubilities of ﬂuorine complexes are about 2 –3 orders of magnitude higher than those of the corresponding metal – non-ﬂuorinated chelates. The ligand selection can determine the extraction efﬁciency. Solubilities of ﬂuorinated b-diketonates (i.e., La(FOD)3) are 1–2 orders of magnitude greater than the corresponding ﬂuorinated dithiocarbamate chelates [18]. The introduction of organic solvents such as methanol into the SF as modiﬁers can improve the solubility of metal chelates (cf. Table 10.3). Fluorinated dithiocarbamate-based metal chelates also have higher stability constants than those of the corresponding non-ﬂuorinated analogues [19]. Changes in the density of SF at constant temperature signiﬁcantly inﬂuence the solubility of metal –ligand chelates, the latter increasing with pressure. The most remarkable change in the ﬂuid solubility with increased pressure is obtained near the critical point (cf. Fig. 10.1). This fact is considered operationally important in extraction systems, where changes in the physical properties of the SF can be utilized to inﬂuence both the extraction selectivity and the recovery itself. A general trend also exists in that trivalent complexes are more soluble than divalent complexes, which in turn are more soluble than monovalent complexes. Increasing ligand hydrocarbon tail lengths increases solubility [17]. 10.3.3 SFE process The SF must be able to dissolve both the ligand and the metal complex in quantities sufﬁcient to achieve extraction on a reasonable time scale. Ligand and metal complex must be sufﬁciently stable to be adequately solvated in the SF. The ligand must be able to access and react with the metal within the sample matrix and the metal ion must be efﬁciently chelated so charge neutralization is achieved. The resulting metal complex must be sufﬁciently soluble and stable in the SF to be rapidly extracted and effectively swept out of the extraction cell. Finally, the metal complex must be transported in the SF without signiﬁcant decomposition [20,21].

328

Supercritical ﬂuid extraction

The extraction of metals using ligands suffers from a number of difﬁculties that are not found with SFE of organic compounds, due to the introduction of complexing agents into the SF stream. This can be achieved by passing the SF (a) over the solid complexing agent, (b) through a liquid complexing agent or (c) passing the complexing agent directly into the SFE cell. In all cases, it can be difﬁcult to determine the concentration of the complexing agent in the SF during the extraction process. The extraction of metals also presents problems regarding the chemical stability of the extraction cell because the ligand may corrode its inner walls. Conventional SFE systems are constructed using stainless steel due its relative inertness, ready availability, tensile strength and low cost. Bartle and coworkers [11] have demonstrated an evident corrosion process of the stainless steel SFE system during extraction of cobalt (608C, 400 bar, HFA as ligand, 30 min ext.), obtaining a high amount of iron in the collection vessel. The corrosion of the SFE system was higher when extraction from soil was attempted, and low recoveries were achieved. Some advantages can be realized with the development of metal-free extraction systems, such as those using PEEK (i.e., polyetheretherketone). This reduces the possibility of contamination of the extracts with foreign materials [19]. However, PEEK can only be reliably operated at a maximum temperature and pressure of 508C and 200 bar, respectively [22]. These limited pressure and temperature ranges are undesirable, as high SF temperatures and pressures are known to increase the analyte solubility and therefore extraction yield, especially when non-spiked samples are extracted. The vessel position (vertical or horizontal) and the outlet valve (inside–outside of the oven) in the extractor system can also affect the SFE recovery [23] Although SFE is mostly applied to solid matrices, in situ chelation –SFE for the extraction of a wide range of metal ions from aqueous solution has been investigated. The use of a special liquid extraction vessel is required in order to carry out the extraction. A study of the extraction kinetics revealed that the extraction of metal ions from aqueous solutions is a rapid process; quantitative extraction can be achieved within 20 min. Tai et al. [24] determined the effective mass-transfer coefﬁcient and found an increase with the stirring rate but decrease with pressure when Cyanex 302 was used as a ligand for the SF extraction of Zn2þ from an aqueous solution. As a result, pressure affects the extraction efﬁciency and the mass-transfer rate but the two are oppositely inﬂuenced, thus the pressure needs to be optimized for practical purposes. For SFE of metal ions from solid samples, a small amount of water is usually added to the solid matrices to improve the extraction efﬁciencies. Water probably facilitates the formation and migration of metal chelates by blocking the active sites in solid matrices holding the metal ions and acting as a matrix modiﬁer [23]. A strong synergistic effect was reported in the SFE of lanthanides [25] and actinides [18] from solid samples when a mixture of tributylphosphate (TBP) and ﬂuorinated b-diketones were used. TBP probably competes with the matrix

329

R. Alzaga et al.

for the unoccupied coordination sites by forming adducts with the actinide- or lanthanide-b-diketone complexes in SF-CO2, making them easier to move from the solid phase into the SF. The extraction efﬁciency obtained from the synergistic extraction of lanthanides and actinides by a mixture of b-diketones and TBP increased in the order AcAc , TAA , FOD ¼ TTA , HFA (cf. Table 10.2). The pKa values for these b-diketones are 9.0, 6.7, 6.7, 6.2 and 4.4, respectively [26]. The pKa order follows the extraction efﬁciency, suggesting a signiﬁcant inﬂuence of the Lewis acidity of the metal chelates on adduct formation. Perﬂuorinated b-diketones are probably more efﬁcient for the synergistic extraction of metal species. The additional electron-withdrawing effect of ﬂuorine in the ligands coordinated with the metal ions enhances the Lewis acidity of the metal chelate. This gives lanthanide- and actinide-bdiketones a stronger tendency to form adducts with the Lewis basic substrate, TBP. Fluorination of the ligands also effectively increases the solubility of the metal chelates. Using the mixed ligand approach, no modiﬁer is needed for the extraction of lanthanides and actinides by SF-CO2. Matrix effects are signiﬁcant for the chelation –SFE of lanthanides and actinides. For the extraction of uranyl ions from sand and ﬁlter papers (TTA þ TBP) or (HFA þ TBP) can extract uranyl ions effectively. However, the extraction of uranyl ions from soil by (TTA þ TBP) is not efﬁcient. A stronger chelating cocktail, HFA combined with TBP, is needed to effectively extract uranyl ions from soil [27]. As noted earlier, a stronger Lewis acid, such as lanthanide- or actinide-bdiketone, can form more stable adducts with the Lewis base, TBP, in SF-CO2, which is why a large synergistic effect is observed. In the same manner, a stronger Lewis base should form a more stable adduct with a Lewis acid, a lanthanide- or actinide-b-diketone. TOPO, a stronger Lewis base than TBP, shows greater synergistic extraction ability than TBP, when mixed with HFA or TTA, for the extraction of lanthanide and uranyl ions from soil and water samples. In uranium contaminated soil samples, even using TOPO þ HFA as a ligand mixture, quantitative extraction was not achieved [23]. The extraction of indigenous uranium was not as rapid as that of spikes added to the samples. The difference is most likely due to the differing chemical nature and accessibility to the SF for the naturally occurring analyte (uranium speciation and matrix effect). Extracted metal chelates are typically transported to the restrictor and trapped in a collection device, where they are often back-extracted with nitric acid solution and analyzed by different techniques (i.e., atomic absorption, emission, or neutron activation). Analytes extracted by this method can potentially be lost through a variety of mechanisms. These include: (a) decomposition of metal chelates in SF-CO2 during their extraction and transport [7,28,29]; (b) precipitation of metal chelates on the wall or end ﬁttings of the extraction vessel, outlet valve, and connecting tubes due to the dead volume and (c) deposition of metal chelates in the outlet valve or restrictor

330

Supercritical ﬂuid extraction

due to decreased solubility during depressurization of CO2 [21,23]. If the extraction is assessed only on the basis of the collection efﬁciency data, the effectiveness of the ligand can be signiﬁcantly underestimated. Conversely, if only the extraction efﬁciency data are used in the assessment, then the recovery of the metal could be signiﬁcantly overestimated. These individual assessments are of limited value, as they do not take into account what happens during the transport of the metal complex through the extraction apparatus, namely the potential for metal complex dissociation. Therefore, to fully understand the extraction process, a simple mass balance of the metal before and after the extraction is necessary, and this can be achieved by sequentially ﬂushing the apparatus with solvents to recover any metal residue. Typically, a recovery discrepancy from 1 to 15% can be obtained. For instance, the nonﬂuorinated metal b-diketones and dithiocarbamates have a greater tendency to dissociate and/or degrade in an SF (i.e., 14–15%) than the corresponding ﬂuorinated metal complexes (i.e., 1–6%) [21]. The kinetics of metal extraction is very important for optimization of both analytical and industrial scale SFE. The rate of metal extraction is affected by several factors, including afﬁnity of ligands and metal chelates to SF and the kinetics of the chelation reaction. Therefore, the selection of a suitable ligand system requires not only the formation of stable metal chelates, but also fast kinetics for chelate formation and sufﬁcient solubility of both ligand and metal chelate in SF-CO2. Short time periods are suggested in the static mode (i.e., 5– 10 min) during SFE performance in order to increase extraction efﬁciency, which is reﬂected by the data in Table 10.3. A highly selective extraction of metals ions can be achieved by SFE. Tomioka et al. [30] found selective extraction of Gd in the presence of Sr and Zr oxides using TBP and HNO3 in the SFE conditions. This supported their subsequent application to the selective cleaning of materials contaminated with radioactive oxides, such as UO2/PuO2, in order to minimize the amount of solvent waste generated during the decontamination step. Chelex 100 was found to be very selective for the extraction of Cu2þ in front of Pb2þ, Zn2þ and Cd2þ from cellulose support [31]. Reverse micelle formation is a recent approach that involves non-ionic surfactants (e.g., Triton X-100) for improving metal extraction efﬁciency with SF-ligand-CO2 [32]. Nearly all SFEs of metals are conducted at low temperature (i.e., 608C), however, there is a study which reports use of high temperatures (1608C) for the extraction of Pb2þ and Mo6þ in non-spiked samples using DDC and AcAc as ligands, respectively [33]. Sequential extraction methods (successive leaching of a sediment/soil by increasing solvent strength) are useful tools for determining the partitioning of contaminants (metals) to sediments/soils and these values can be related to their bioavailability (see Sequential Extraction, Chapter 39). Clark and coworkers [34] have studied the partitioning of stable (Ca, Fe, Y) and radioactive (U) elements

331

R. Alzaga et al.

in two different “native” sediments using solvent extraction (SE) combined with SFE/SE (extraction conditions: 20– 22 g sample, 1.2 – 2 g of TTA, 200 bar, CO2 þ TBP (2%), extraction times: 30 min/30 min static/dynamic mode). In the ﬁrst sediment, SFE removed more metals and had a greater impact on the subsequent distributions of the remaining metals (50–90% decrease in metal), compared to the other (not affected). SFE-metal performance is closely related to the matrix effect. Different extraction conditions give different recoveries, and can be correlated to their availability. Ligand-assisted SFE-CO2 could be proposed as an alternative solvent extraction technique for the selective removal of heavy metals and radionuclides from both liquid and solid waste matrices. SFE could be employed as a separation technique for extraction of key metals and radionuclides to reduce their concentrations to a level below established regulatory limits that would result in waste re-classiﬁcation (i.e., from high-level radioactive waste to a lowlevel radioactive waste). A waste stream could be effectively processed, resulting in signiﬁcant waste volume minimization with minimal production of secondary wastes, compared to traditional techniques. In the waste management area, ligand-assisted SFE has the potential to selectively remove metals to the parts-per-billion range or lower. Solid materials, such as tools, personal protective equipment, etc., can easily be surface decontaminated using a ligand-assisted SFE process, reducing the total mass of radioactive materials entering the waste stream. Liquid waste streams having extreme pH, high salt, organic or solid content can be treated using the ligand-assisted SFE technique without the need for prior chemical neutralization or solid separation. Wastewaters treated with ligand-assisted SFE processes would be of high enough quality to be re-used as a solvent, thus reducing the overall quantity of water needed in a process. 10.4

ORGANOMETALLIC COMPOUNDS

The procedures developed for SFE of organometallics are similar to those exposed for trace elements. However, in this section the extraction procedures will be presented according to the speciﬁc element and matrix. SFE has been evaluated in speciation studies of several elements, such as tin, lead, mercury and arsenic. The aim in the application of SFE to speciation studies is a reduction in the number of analytical steps and a minimization in the usage of hazardous acids and solvents which are commonly used in speciation studies of abiotic matrices [35]. 10.4.1 Organotin compounds The most commonly determined organotin compounds are tetraorganotins (R4Sn), triorganotins (R3SnX), diorganotins (R2SnX2) and monoorganotins

332

Supercritical ﬂuid extraction

(RSnX3), where R can be methyl, butyl, cyclohexyl and phenyl and X is the counter ion, which can be a halogen, OH2, S22, etc. The general aspects of sample preparation regarding tin and arsenic speciation are presented in detail in Chapters 30 and 31, respectively. In this chapter, only methods based on SFE are presented. 10.4.1.1 Abiotic matrices SFE of organotins ðRm SnX42m ; m ¼ 1 – 3Þ from aqueous matrices cannot be accomplished directly by CO2 because of the requirement for charge neutralization and the weak solute –SF interaction. However, when ionic compounds are bound to organic ligands or by ion-pair formation, their solubility in SF-CO2 is signiﬁcantly increased, thus enabling the extraction of these compounds from environmental samples. Consequently, direct extraction of organometallic species from aqueous samples has not been attempted, but the combination of solid-phase extraction (SPE) followed by SFE of the alkylated organotin derivatives has been successful (cf. Table 10.4). By using this analytical procedure, two different approaches have been developed: (i)

Extraction by SPE of the organotin chlorides followed by Grignard derivatization (EtMgBr) on the SPE membrane [36] (ii) Aqueous-phase derivatization with NaBEt4 followed by SPE [37] The former has only been successful for the determination of di- and tributyltin (TBT) species, but the latter has permitted simultaneous speciation of butyl-, phenyl-, and cyclohexyltins. In both analytical procedures, the derivatized organotin species in the SPE membrane were extracted quantitatively with neat SF-CO2 by combining a static and dynamic extraction at 300 bar and 408C. The different extraction behaviors between these analytical procedures can be accounted for by the low efﬁciency of extraction of ionic TABLE 10.4 SFE conditions for recovery of organotin compounds from aqueous matrices Preconcentration

Derivati- Extractant zation agent

Temperature Pressure Recovery Reference (8C) (MPa) (%)

C18-Empore disk C18-Empore disk Cation exchange

EtMgBr

CO2

40

10

92– 102a

[36]

NaEt4B

CO2

40

30

80–110b

[37]

CO2 –HCO2H 60

25

90c

[40]

–

a

Recovery of DBT and TBT chlorides from spiked seawater. Sea water spiked with MBT, DBT, TBT, MPhT, DPhT, TPhT, DCT and TCT chlorides. Spiked with TPhT, DPhT and TCT chlorides.

b c

333

R. Alzaga et al.

organotin species in the hydrophobic C18 membranes (ﬁrst procedure) while the ethylated organotins obtained in the aqueous phase (second procedure) are easily retained in the extraction membrane because of their increased hydrophobicity following the derivatization step. Although SPE preconcentration of organotins from aqueous matrices has been performed with C18 membranes followed by solvent elution [38], it only shows an effective recovery for TBT. Therefore, the advantages of SPE – SFE, in comparison with conventional SPE using liquid solvents, are a reduction in the solvent volume, broader range of analytes extracted and extraction selectivity. Therefore, organic extracts obtained by SFE can be analyzed without further clean up even using analytical techniques having medium selectivity, such as GC-FPD. These analytical procedures have been applied to the determination of organotins in marinas and continental waters from the western Mediterranean [37]. Another approach, not fully investigated, is the preconcentration of phenyl and cyclohexyltin compounds from aqueous samples on cationic exchange resins followed by SFE with CO2 –HCO2H (cf. Table 10.4). The main limitation of this methodology is the impossibility of its application to samples having a moderate to high ionic strength, because of the low capacity of the cationic exchange resins in such aqueous matrices. In this regard, the determination of organotin compounds from seawater is not feasible. No reports have been published regarding the SFE of other organometallics from aqueous samples. Extraction of organotins by SFE has been evaluated in a variety of solid matrices, such as food products (potato, almonds) and abiotic samples (paint coatings, soils, sediments, cf. Table 10.5). The most important variables in the optimization of the SFE methodology are: i) extraction agents, ii) pressure and temperature effects, iii) complexing agent effects and iv) derivatization reactions. Extraction agents. These are key variables in the optimization of SFE. The extraction of ionic organotins requires modiﬁed CO2, otherwise their recovery, even under conditions of high solubility (high pressure, low temperature) is very poor. In this regard, some investigations indicate that carbon dioxide modiﬁed with 5% methanol is a better extraction ﬂuid than neat carbon dioxide [39]. Typically, modiﬁed carbon dioxide has been demonstrated to be effective with different matrices. Formic acid modiﬁed CO2 (0.2% v/v) has been proven to be effective for foodstuffs (organotin spiked into potato or almond matrix) [40]. High concentrations of modiﬁer (20% v/v HCl – MeOH in CO2) become necessary for the recovery of TBT from lacustrine and marine sediments, attributable to its stronger interaction with these matrices [41]. Nevertheless, the TBT degradation products, namely dibutyltin (DBT) and monobutyltin (MBT), are not quantitatively extractable using the TBT extraction conditions. Other acidic modiﬁers, such as formic or propionic acids, only increase the recovery of a few organotin compounds.

334

TABLE 10.5 SFE conditions for recovery of organotin species from solid non-biotic matrices Matrixa

Foodstuffs marine paint

Sediments,n Sediments,n

Sedimentn Biotas Musseln, ﬁshn a

s, spiked; n, native.

Butyl, phenyl, cyclohexyl Butyl Butyl, phenyl Butyl Methyl, ethyl, butyl, cyclohexyl, phenyl Butyl, phenyl Butyl Butyl, phenyl

Extractant agent

Complexing/ derivatization agents

Determination

Reference

CO2 –HCO2H

–

SFE–SFC-FID

[40]

CO2 –MeOH– HCl CO2

– HexMgBr

GC-FPD GC-FPD

[41] [42]

CO2 –MeOH CO2 –MeOH

NaDDC NaDDC

GC-AED GC-AED

[43] [39]

CO2 –HCOOH

APDC, NaDDC – Tropolone

GC-FPD

[44]

SFC–ICP-MS GC-FPD

[47] [48]

CO2 –MeOH CO2 –MeOH, CO2 –HOAc

Supercritical ﬂuid extraction

Sediments,n Sediments,n

Analytes

335

R. Alzaga et al.

Pressure and temperature effects. As mentioned earlier, underivatized organotin compounds cannot be extracted with neat CO2 because of their low solubility. However, if SFE is performed on derivatized organotins with neat CO2, high pressure and low temperatures are required [42]. Pressures in the range of 300 –500 bar and temperatures in the range of 50 –808C have little effect on the extraction efﬁciency of organotin compounds when polar or acidic modiﬁers are used [39,43,44]. At lower pressures (100 –300 bar), there is a strong effect on the extraction recovery for the less soluble mono-, di- alkyl and phenyltins. Complexing agent effects. Six tetraalkyltin compounds spiked to topsoil samples were extracted by SFE with recoveries in the range of 90–110%. Recoveries ranging from 70 to 90% were obtained for most ionic organotin compounds when sodium diethyldithiocarbamate (NaDDC) ligand was used [22]. Bayona’s investigation [44] found that complexing agents [i.e., diethylammonium diethyldithiocarbamate (DEA-DDC), APDC], in the presence of polar modiﬁers, do not improve the extraction efﬁciency of native organotins, but an increase in the extraction efﬁciency of organotin compounds from spiked matrices has been observed [39,43]. Soil and sediment samples, mixed with DEA-DDC were extracted with 5% methanol modiﬁed CO2 at 608C and 450 bar [39]. The extracted analytes were treated with pentylmagnesium bromide to convert the ionic organotin compounds into their neutral derivatives, which were later determined by GC-AED. These results indicate that SFE followed by GC-AED detection is a promising technique for the determination of trace organotin compounds in environmental soil and sediment samples. Furthermore, complexing agents (i.e., APDC, DDC) in the presence of acidic modiﬁers (AcOH) do not improve their extraction efﬁciency [44], attributable to a rather limited stability of complexing agents under SFE conditions in the presence of acid modiﬁers. The use of neat acetic acid as CO2 modiﬁer in SFE provided the highest extraction efﬁciency of native organotins from sediment due to the combined effects of a displacement mechanism of the analytes from the matrix by HAOc and the formation of organotin acetate, which is readily soluble in CO2 (cf. Table 10.5). Evidence for the counter ion displacement under SFE conditions is given by the same retention in SFC, independent of the counter ion when formic acid is used as a CO2 modiﬁer [40]. Derivatization reactions. Butyl and phenyltin compounds can be derivatized in the extraction cell with hexylmagnesium bromide, yielding the hexylated derivatives of organotin compounds prior to their SFE with neat CO2 [42]. This analytical procedure saves time because the alkylated derivatives are formed in the extraction cell and they are ready to be determined by GC techniques. However, it suffers some limitations, such as (i) small amount of sample (500 mg), (ii) sample dryness is very critical, (iii) requires a large volume of derivatization reagents because they react with other components of the matrix and (iv) alkylsulﬁde formation which can

336

Supercritical ﬂuid extraction

interfere with the determination by GC-FPD. No reports are available describing the derivatization reactions under SFE conditions. Most of the above mentioned extraction procedures are useful for the quantitative extraction of di- and tri- butyl- and phenyltin compounds when certiﬁed reference materials (CRMs) are analyzed. However, the extraction efﬁciency of monoalkyl and monoaryltin compounds achieved with SFE appears to be lower than values obtained by LSE techniques. A round robin test with eight participants was conducted on three spiked soils and sediments and one unspiked sediment using the same analytical methodology (CO2 – MeOH, NaDDC) but using different SFE instrumentation with different collection systems (dichloromethane and C18), and the organotin determination was achieved with AED [45]. Results were independent of the analytical instrumentation and dependent on the methodology, but satisfactory results were obtained for tetra- and tri-substituted organotin compounds. 10.4.1.2 Biotic matrices SF-CO2 was used for the extraction of organotin compounds present in ﬁsh. A sample of tuna ﬁsh was spiked with trimethyltin chloride, TBT chloride and triphenyltin chloride [46]. Extraction conditions were optimized (pressure: 415 bars; temperature, 808C; static extraction time 30 min; dynamic extraction time, 15 min) by measuring total metal content by liquid chromatographyinductively coupled plasma mass spectrometry (LC-ICP-MS). Extraction efﬁciencies up to 80% for trimethyltin and 40% for triphenyltin were obtained with ﬁsh tissue CRM when 1% water was added as a modiﬁer and ammonium salts of diethyldithiocarbamic acid or pyrrolidinecarbothioic acid were added as complexing agents. SFE can be applied to freeze-dried homogenates of biotic tissues [47,48]. Alternatively, the extraction can be carried out on fresh tissue homogenates mixed with an adsorbent (Hydromatrix, anhydrous Na2SO4), thereby avoiding the freeze-drying process [49]. Organometallic compounds occurring in biotic matrices have fewer interactions compared to abiotic matrices [35]. Consequently, extraction can be achieved under milder conditions, which improves the selectivity of the extraction, leading to less extensive clean up steps following the extraction. Early reports wherein CO2 modiﬁed with MeOH was used resulted in low recoveries of TBT [44]. More recently, HOAc acid has been found more effective than MeOH for the SFE of native DBT and TBT from a mussel tissue, and complexing agents, such as tropolone, improve the extraction of butyl and phenyltins from ﬁsh tissue. Furthermore, an experimental design approach was applied for systematic optimization of temperature, pressure, modiﬁer volume and static time. Extraction efﬁciency was positively correlated with pressure and the mutual interaction between modiﬁer volume and temperature. A negative correlation with temperature occurred, suggesting that high solubility conditions are needed for extraction from this matrix. A cross-

337

R. Alzaga et al.

comparison between the results obtained using the developed SFE procedure and those obtained with an established liquid-solvent extraction of ﬁsh and clam tissues demonstrates that SFE is more effective for TBT but gives poorer results for TPhT [47]. Further research is needed to improve the extraction efﬁciency of the remaining analytes. 10.4.2 Organomercury compounds Optimization of SFE of methylmercury (MeHg) in marine samples followed by GC determination with electron capture detection (GC-ECD) was also developed [50]. An experimental design approach was used for the simultaneous determination of various SFE parameters, such as CO2 ﬂow rate and density, temperature, pressure, static extraction time, the amount of HCl and the contact time between HCl and the sample prior to extraction. A review summarized the SFE conditions for extraction of methylmercury from aquatic sediments. The analytical procedures for methylmercury determination were based on GC-ECD [51]. A method employing SFE and GC coupled with microwave-induced plasma atomic emission spectrometry (GC –MIP-AES) was used for the determination of methylmercury in sediments [52]. Butylmagnesium chloride was used to derivatize the target compound to methylmercury, rendering it amenable to GC. The studies of Liu et al. [39] illustrated the application of SF-CO2 modiﬁed with 5% of MeOH for the extraction of individual organomercury compounds, including CH3HgCl, C6H5HgCl, (C6H5)2Hg and inorganic mercury compounds (i.e., HgCl2, HgO, HgS) from soil and sediment samples, followed by their capillary GC-AED determination. Diorganomercury can be determined without pretreatment, whereas monoorganomercury and inorganic mercury compounds have to be soluble in SF-CO2. This can be achieved by complexation of monoorganomercury with NaDDC. Cellulose, spiked with Hg2þ can be extracted with CO2 and methanol containing the chlorinated chelating agent lithium bis(triﬂuoroethyl)dithiocarbamate (LiFDDC) because the Hg(FDDC)2 complex has a high solubility in CO2 [53]. Other mercury species (MeHgCl and Me2Hg) can be extracted with SF-CO2 without the need for a chelating agent and modiﬁer. Further research is needed to apply this extraction procedure to real samples where organomercury species interact strongly with the matrix. Baxter obtained reasonable recoveries for the MeHgþ from sediments containing elemental sulfur when using neat CO2 and attributed this to the formation of methylmercury sulﬁdes, which are soluble in CO2 [53]. No MeHg formation is reported in SFE from soil and sediments. A method for extracting methylmercury from seafood using SF-CO2 is described by Holak [54]. This method was validated through the analysis of CRMs, and applied to commercial samples and spikes. Extraction of methylmercury from seafood is difﬁcult because the compound is not present

338

Supercritical ﬂuid extraction

in free state, but bound to proteins. Thus, MeHg must be cleaved from protein binding sites before extraction, usually by treatment with hydrochloric acid. For extraction by SF-CO2, the matrix must be degraded by pretreatment with base (1 N NaOH). The extraction is performed under the following conditions: pressure, 200 bars; temperature, 508C; time, 20 min. After extracting and trapping MeHg in a thiosulfate solution (0.01 N Na2S2O3), the determination can be accomplished by any suitable method, such as cold-vapor atomic absorption spectrometry. 10.4.3 Organolead compounds Spikes of trialkyl- and dialkyllead (methyl and ethyl) added to sediment dust have been quantitatively extracted using modiﬁed CO2. Methanol was found to be the most effective modiﬁer, compared to acetone, and water and methanol was also used for analyte collection. Extraction efﬁciency increased with temperature (55–808C) and pressure (100–450 bars). Under optimum extraction conditions, recoveries of 96, 106 and 80% were obtained for trimethyllead, triethyllead and diethyllead, respectively, from spiked sediment. The validation of the analytical procedure for trimethyllead was demonstrated in an intercalibration exercise using urban dust in the framework of the Standards, Measuring and Testing Program of the EU [55]. 10.4.4 Arsenic compounds Organic dimethylarsenic and monomethylarsenic (DMA and MMA) and inorganic arsenic species [As(III) and As(V)] can be extracted from spiked samples with CO2 in the presence of thioglycolic acid methylester [56]. The derivatization reaction is accomplished in supercritical CO2, leading to the formation of derivatives that can be reproducibly determined by GC (DMA and MMA). Reported recoveries for MMA and DMA under optimum extraction conditions range from 90 to 103%, compared to a liquid-solvent extraction technique, with R.S.D. values of 1–8%. However, this procedure has not been validated with CRMs. 10.5

CONCLUSION

A large variety of chelating agents, including ionizable crown ethers, bdiketones, dithiocarbamates, organophosphorus and ﬂuorinated hydroxamic acid have been evaluated for the extraction of trace elements (i.e., Fe3þ, Cu2þ, Zn2þ, Cd2þ, As3þ, Pd2þ, La3þ, Co2þ, (UO2)2þ, Rh3þ) with SF-CO2. Among them, ﬂuorinated chelating agents are particularly effective because they possess high solubility in SF-CO2. Moreover, the addition of TBP exerts a synergistic effect, probably by competition with the matrix for the unoccupied sites. Nevertheless, the high cost of these ligands restricts their applications to the

339

R. Alzaga et al.

analytical scale. Further research is needed to develop stable ligands at pressures and temperatures typical to SFE with higher solubility in SF-CO2. Although most of the ligand-assisted SFEs of trace elements have been accomplished in spiked inert matrices (i.e., cellulose, ﬁlter paper, glass beads), real samples and reference materials have rarely been evaluated. Therefore, validation of the extraction procedures needs further attention. Extraction selectivity is another aspect of future development. In fact, preliminary studies have shown that extraction can be accomplished selectively in the presence of additives with Lewis acid– base properties. Technological applications arising from extraction selectivity present an area of emerging interest to the ﬁeld of cleaning materials contaminated with radionuclides. The main advantages, in comparison to conventional methods of cleaning, are the cost-effective recycling of CO2 and low toxicity versus methods based on solvents or water. Corrosion is one of the limitations encountered with SFE instrumentation, constructed of conventional stainless steel, when a cocktail of chelating agents and additives are used for the extraction of trace elements in environmental samples. Inert materials stable at moderate temperatures and pressures for trace level analytical applications need to be developed. Applications of ligand-assisted SFE with CO2 in the ﬁeld of speciation of tin, mercury, arsenic and organolead compounds have been partially successful when they are applied to real samples. The main limitations are desorption of the analyte from the matrix and the different solubilities of the chemical species in the extracting ﬂuid, thus affecting the recovery. Up to now, a great deal of attention has been devoted to mercury and tin in different environmental matrices and successful results for several species have been obtained when validated against reference materials. Finally, sequential SFE of trace elements can be of interest in remediation studies because the bioavailable fraction can be extracted by increasing pressure, temperature, or the ligand and organic modiﬁer concentrations. This procedure could be complementary to sequential solvent extraction, which is time consuming and difﬁcult to automate.

REFERENCES 1 2 3 4 5 6 7

340

M.L. Lee and K.E. Markides, Analytical Supercritical Fluid Chromatography and Extraction. Chromatography Conferences, Inc. Provo, UT, 1990. J.C. Giddings, M.N. Meyers and R.A. Keller, Science, 67 (1969) 162. R. Alzaga, J.M. Bayona and D. Barcelo´, J. Agric. Food Chem., 43 (1995) 395. J.M. DeSimone, Z. Guan and C.S. Elsbern, Science, 257 (1992) 945. S.B. Hawthorne, Anal. Chem., 62 (1990) 633A. C.M. Cowey, K.D. Bartle, M.D. Burford, A.A. Clifford, S. Zhu, N.G. Smart and N.D. Tinker, J. Chem. Eng. Data, 40 (1995) 1217. J. Wang and W.D. Marshall, Anal. Chem., 66 (1994) 3900.

Supercritical ﬂuid extraction 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45

M.J. Carrott and C.M. Wai, Anal. Chem., 70 (1998) 2421. K.E. Laintz, C.M. Wai, C.R. Yonker and R.D. Smith, J. Supercrit. Fluids, 4 (1991) 194. M. Ashraf-Khorassani, M.T. Combs and L.T. Taylor, Talanta, 44 (1997) 755. ¨ zel, K.D. Bartle, A.A. Clifford and M.D. Burford, Anal. Chim. Acta, 417 M.Z. O (2000) 177. R.F. Fedors, Polym. Eng. Sci., 14 (1974) 147. R.F. Fedors, Polym. Eng. Sci., 14 (1974) 472. C.M. Wai, S. Wang and J. Yu, Anal. Chem., 68 (1996) 3516. K.L. Toews, R.M. Shroll, C.M. Wai and N.G. Smart, Anal. Chem., 67 (1995) 4040. Y. Meguro, S. Iso, T. Sasaki and Z. Yoshida, Anal. Chem., 70 (1998) 774. N.G. Smart, T.E. Carleson, T. Karst, A.A. Clifford, M.D. Burford and C.M. Wai, Talanta, 44 (1997) 137. Y. Lin, R.D. Brauer, K.E. Laintz and C.M. Wai, Anal. Chem., 65 (1993) 2549. Y. Lin, N.G. Smart and C.M. Wai, Trends Anal. Chem., 14 (1995) 123. S. Elshani, N.G. Smart, Y. Lin and C.M. Wai, Sep. Sci. Technol., 36 (2001) 1197. ¨ zel, A.A. Clifford, K.D. Bartle, Y. Lin, C.M. Wai and M.D. Burford, M.Z. O N.G. Smart, Analyst, 124 (1999) 609. Y. Liu, V. Lopez-Avila, M. Alcaraz, W.F. Beckert and E.M. Heithmar, J. Chromatogr. Sci., 31 (1993) 310. Y. Lin, H. Wu, N.G. Smart and C.M. Wai, Sep. Sci. Technol., 36 (2001) 1149. C.Y. Tai, G.-S. You and S.-L. Chen, J. Sep. Fluid, 18 (2000) 201. Y. Lin and C.M. Wai, Anal. Chem., 66 (1994) 1971. K.L. Cheng, K. Ueno and T. Imamura, Handbook of Organic Analytical Reagents. CRC Press, Boca Raton, FL, 1982 Y. Lin, C.M. Wai, F.M. Jean and R.D. Brauer, Environ. Sci. Technol., 28 (1994) 1190. C. Fujimoto, H. Yoshida and K. Jinno, J. Microcolumn Sep., 1 (1989) 19. K. Jinno, H. Mae and C. Fujimoto, J. High Resolut. Chromatogr., 13 (1990) 13. O. Tomioka, Y. Enokida and I. Yamamoto, Prog. Nucl. Energy, 37 (2000) 417. N.G. Smart, T.E. Carleson, S. Elshani, S. Wang and C.M. Wai, Ind. Eng. Chem. Res., 36 (1997) 1819. J. Liu, W. Wang and G. Li, Talanta, 53 (2001) 1149. V. Arancibia, R. Segura, J.C. Leiva, R. Contreras and M. Valderrama, J. Chromatogr. Sci., 38 (2000) 21. S.M. Loyland, M. Yeh, C. Phelps and S.B. Clark, J. Radioanal. Nucl. Chem., 248 (2001) 493. ´ balos, J.M. Bayona, R. Compan˜o´, M. Granados, C. Leal and M.D. Prat, M. A J. Chromatogr. A, 788 (1997) 1. R. Alzaga and J.M. Bayona, J. Chromatogr. A, 655 (1993) 51. Y. Cai and J.M. Bayona, J. Chromatogr. Sci., 33 (1995) 89. O. Evans, B.J. Jacobs and L. Cohen, Analyst, 116 (1991) 15. ´ vila, M. Alcaraz and W.F. Beckert, Anal. Chem., 66 (1994) 3788. Y. Liu, V. Lo´pez-A J.W. Oudsema and C.F. Poole, Fresenius J. Anal. Chem., 344 (1992) 426. J. Dachs, R. Alzaga, J.M. Bayona and P. Quevauviller, Anal. Chim. Acta, 286 (1994) 319. Y. Cai, R. Alzaga and J.M. Bayona, Anal. Chem., 66 (1994) 1161. Y.K. Chau, F. Yang and M. Brown, Anal. Chim. Acta, 304 (1995) 85. ´ balos and J.M. Bayona, Appl. Organomet. Chem., 12 (1998) 577. Y. Cai, M. A V. Lo´pez-Avila, Y. Liu and W.F. Beckert, J. Chromatogr. A, 785 (1997) 279.

341

R. Alzaga et al. 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66

342

N.P. Vela and J.A. Caruso, J. Anal. At. Spectrom., 11 (1996) 1129. U.T. Kumar, N.P. Vela, J.G. Dorsey and J.A. Caruso, J. Chromatogr. A, 655 (1993) 340. I. Ferna´ndez-Escobar and J.M. Bayona, Anal. Chim. Acta, 355 (1997) 269. J.M. Bayona, in: E.D. Ramsey (Ed.), Analytical Supercritical Fluid Extraction. Kluwer Academic, Dordrecht, 1998, p. 72. R. Cela-Torrijos, M. Miguens-Rodriguez, A.M. Carro-Dı´az and R.A. LorenzoFerreira, J. Chromatogr. A, 750 (1996) 410. R.A. Lorenzo, M.J. Va´zquez, A.M. Carro and R. Cela, Trends Anal. Chem., 18 (1999) 410. H. Emteborg, E. Bjorklund, F. Odman, L. Karlsson, L. Mathiasson, W. Frech and D.C. Baxter, Analyst, 121 (1996) 19. C.M. Wai, Y. Lin, R.D. Brauer and S. Wang, Talanta, 40 (1993) 1325. W. Holak, J. AOAC Int., 78 (1995) 1124. M. Johansson, T. Berglo¨f, D.C. Baxter and W. Frech, Analyst, 120 (1995) 755. B.W. Wenclawiak and M. Krah, Fresenius J. Anal. Chem., 351 (1995) 134. K.E. Laintz, C.M. Wai, C.R. Yonker and R.D. Smith, Anal. Chem., 64 (1992) 2875. Y. Lin, N.G. Smart and C.M. Wai, Environ. Sci. Technol., 29 (1995) 2706. C.M. Wai, Anal. Sci., 11 (1995) 165. S. Wang, S. Elshani and C.M. Wai, Anal. Chem., 67 (1995) 919. K.G. Furton, L. Chen and R. Jaffe´, Anal. Chim. Acta, 304 (1995) 203. J. Wang and W.D. Marshall, Analyst, 120 (1995) 623 –627. C.M. Wai, S. Wang, Y. Liu, V. Lopez-Avila and W.F. Beckert, Talanta, 43 (1996) 2083. B. Wenclawiak, T. Hees, C.E. Zo¨ller and H.P. Kabus, Fresenius J. Anal. Chem., 358 (1997) 471. J.D. Glennon, S. Hutchinson, A. Walker, S.J. Harris and C.C. McSweeney, J. Chromatogr. A, 770 (1997) 85. S.N. Joung, S.J. Yoon, S.Y. Kim and K.-P. Yoo, J. Supercrit. Fluids, 18 (2000) 157.

Chapter 11

Accelerated solvent extraction of organometallic and inorganic compounds John L. Ezzell

11.1

ACCELERATED SOLVENT EXTRACTION AS A SAMPLE PREPARATION TECHNIQUE

11.1.1 Introduction In order to assess the inorganic analyte content of a solid sample, the compounds of interest are normally extracted or digested into the liquid state for introduction into quantifying instrumentation. There has been a trend in recent years toward a reduction in the time and volume of hazardous solvents used in these sample preparation techniques. Alternatives to traditional techniques have been introduced and met with varying levels of acceptance in the sample preparation area. Automated Soxhlet, supercritical ﬂuid, and microwave assisted extraction/digestion techniques have all been developed in an attempt to solve the challenges of sample preparation. In each of these techniques, temperature is used as a fundamental parameter. Accelerated solvent extraction (ASEw, registered trademark of Dionex Corporation) is a technique developed for the extraction of solid and semi-solid sample matrices using liquid solvents at temperatures above their normal (atmospheric) boiling points. By taking advantage of an increase in analyte solubility and solvent strength, which result from an increase in temperature, ASE is able to increase the efﬁciency of the extraction process. This results in extractions performed in less time, and using less solvent than traditional methods. With ASE, most extractions are performed in the 75–1508C range. In order to use common aqueous and organic solvents in this temperature range, pressure is applied to maintain the ﬂuids in their liquid state. A typical ASE extraction is performed in 15 min, using between 15 and 45 ml of solvent for sample sizes ranging from 1 to 30 g. Since its development and commercialization as ASE by Richter and Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

343

J. L. Ezzell

co-workers in 1995 [1], ASE has gained rapid acceptance in the environmental, polymer, food, and pharmaceutical industries. 11.1.2 Basic principles of ASE operation In the area of inorganic sample preparation, digestion with strong acids is commonly used. Extraction, by deﬁnition, is a different approach because the target analyte(s) are separated (solubilized) away from the sample by the extraction solvent, rather than dissolving the entire sample matrix. In order to extract a target analyte from a solid matrix into a liquid solvent, there are interactions which must be overcome, and interactions which must be formed. First, the interaction between the solute molecule (target analyte) and its neighboring solute and/or matrix molecules must be disrupted. This interactive force can be attributed to hydrogen bonding, Van der Waals interactions, and dipolar interactions among the molecules. The energy required to break these interactions can be called the lattice energy, the heat of sublimation or the heat of vaporization. This value will generally increase with the polarity of the solute molecule. Second, once the solute is free from the matrix, it must be solubilized into the liquid extraction solvent. Initially, this requires the breaking of interactions between solvent molecules. This can occur in two ways, ﬁrst, the solvent interactions must be weakened to the point that it can “wet” the solid surface of the matrix. This allows the solvent to exert a force in disrupting the solute –matrix interaction described above. Second, the solvent network is weakened to the point that the solute can be accommodated into the solvent environment. The energy required for this process is largest in polar solvents, and can be estimated by reference to such parameters as surface tension and boiling point. Once solubilized into the solvent, the solute molecule now forms a new interaction with the solvent molecules, as a stable solvent cage is formed around it. The stability of this interaction results in a release of energy, which is largest when the interaction of the solute and solvent is more polar. In order to achieve maximum solubility in the shortest time period, the energies described above, which act as barriers to solvation (and therefore extraction) must be overcome. Increasing the temperature of the process should act to increase the rate of the required steps necessary for efﬁcient extraction to occur. An increase in temperature will act to decrease the magnitude of hydrogen bonding interactions and dipolar interactions occurring on the high energy surfaces of the sample matrix. Thermal energy will also assist in disrupting stable lattice structures and impart a degree of kinetic energy to the solute molecules. In addition, a general decrease in solvent–solvent interactions will more easily accommodate solute molecules into the solvent environment. Through the application of increased temperature, ASE is able to maximize solubilization by overcoming the energy barriers associated with extraction, and thereby increase the efﬁciency of the process. Pressure is a required

344

Accelerated solvent extraction of organometallic and inorganic compounds

component of ASE, since most of the extraction solvents used have atmospheric boiling points lower than the temperatures at which they are commonly used in this technique. Increased pressure can also have a positive role in the extraction process by forcing the solvent into sample matrix pores and increasing the collision probability of solute and solvent molecules. 11.1.3 ASE instrumentation A schematic diagram of a ASE system is shown in Fig. 11.1. The ASE extraction procedure consists of a combination of dynamic and static ﬂow of the solvent through a heated extraction cell containing the sample. These cells must be capable of safely withstanding the pressure requirements of the system, and are normally constructed of stainless steel, with frits in the end caps to allow the passage of solvent while maintaining the solid sample within. The pore size of the frit should not allow passage of the matrix particles (5 –10 mm is typical). The sample cell is interfaced to the solvent ﬂow path, where it is ﬁlled with solvent. It is important to ensure that all of the void volume has been ﬁlled with solvent in order to ensure good contact between the sample matrix and the solvent, and to avoid possible analyte oxidation which may occur in the presence of air at elevated temperatures. The sample cell is then heated by direct contact with a heat source. Heating the cell prior to solvent introduction can result in the loss of volatile compounds [2]. In order to accurately assess the effect of the extraction temperature, the inside of the cell must be allowed to reach thermal equilibrium with the heating source. In order to maintain the extraction solvents in their liquid state, a pressure source must be applied. The system pressure must be above the threshold required to maintain the liquid state of the solvent at the set temperature and be able to move the solvent through the sample cell in a reasonable time period. This is normally accomplished with an HPLC type pump, which can maintain a constant ﬂuid

Fig. 11.1. Schematic diagram of ASE extraction system.

345

J. L. Ezzell

pressure of 1000 –3000 psi (6.9 –20.7 MPa). When heating a liquid solvent in a closed cell, the solvent will expand and result in an increase in pressure. This increase must be regulated without the loss of solvent from the cell (which contains target analytes). In order to accomplish this, a valve is positioned between the extraction cell and the collection vial. This valve is pulsed at intervals during the heating process, allowing small volumes of ﬂuid to escape from the cell into the vial. Due to the constant pressure applied by the pump, an additional amount of fresh solvent is introduced to the head of the cell as the valve pulses, maintaining the cell volume. Once thermal equilibrium has been reached, the sample cell is maintained at temperature for an additional time period, which is usually equal to the heat time, or approximately 5 min. During this static phase, analyte diffusion from the matrix into the solvent is believed to occur. Following this static hold step, the outlet valve is opened and a measured volume of fresh solvent, usually 50 –70% of the cell volume, is allowed to ﬂush over the sample, discharging the previous volume into the collection vial. Lastly, compressed nitrogen is used to force all of the solvent from the lines and cell, into the vial. It is important that all of the liquid solvent used in the extraction process be collected for analysis. The collection vials normally used are standard 40 or 60 ml vials sealed with a (Teﬂonw, registered trademark of Dupont) coated septa. This allows the extracts to be collected and maintained in a sealed, inert environment (under a nitrogen blanket) in order to prevent sample loss while waiting for quantiﬁcation. ASE is normally accomplished with a ﬂuid ﬂow from the top to the bottom of the cell, as this facilitates purging of the liquid with the compressed gas source, however, ﬂow direction is not critical. Due to the high operating temperatures of the system, solvents with unusually low auto-ignition points, such as carbon disulﬁde (CS2), should be avoided. 11.1.4 ASE methods development 11.1.4.1 Sample preparation Sample preparation is a signiﬁcant part of every solvent based extraction procedure. While many sample types can be efﬁciently extracted without any pre-treatment, other samples will require some manipulation for an efﬁcient extraction to occur. The ideal sample for extraction is a dry, porous, ﬁnely divided solid. Whatever can be done to make the sample approach this deﬁnition (within appropriate cost and time constraints) will have a positive impact on the results of the extraction. In general, the same sample preparation that is done prior to traditional extraction methods should be done prior to extraction by ASE. 11.1.4.2 Grinding For an efﬁcient extraction to occur, the solvent must be able to make physical contact with the target analytes. The more surface area that can be exposed in a

346

Accelerated solvent extraction of organometallic and inorganic compounds

sample, the faster that this contact will occur. Samples with large particle sizes should be ground prior to extraction. The minimum particle size required may differ based on the sample type, but should be generally smaller than 1.0 mm. Grinding can be accomplished with a conventional mortar and pestle, or with electric grinders and mills. Since quantitative transfer of ground material can be difﬁcult, it is recommended that a large, representative sample be ground, and weighed aliquots of the ground sample be used for extraction. 11.1.4.3 Dispersing It is possible that the interaction of sample particles may prevent efﬁcient extraction. In these cases, dispersing the sample with an inert material such as sand or diatomaceous earth (DE) will assist in the extraction process. Dispersion will also assist the extraction process when dealing with samples which tend to compact in the extraction cell outlet. These samples normally contain very small particles (ﬁnes) that can adhere tightly together under pressure. When using a dispersing agent, it is good practice to periodically run blank extractions of the material to verify its cleanliness. 11.1.4.4 Drying Many environmental samples contain water which can act as a barrier, preventing non-polar organic solvents from reaching the target analytes. While the use of more polar solvents (acetone, methanol, water) or solvent mixtures (hexane/acetone, methylene chloride/acetone) can be used to assist in the extraction of wet samples, sample drying prior to extraction is the most efﬁcient way to handle these sample types. Drying is normally accomplished by direct addition of a material such as sodium sulfate or DE. The choice of drying agent depends on the preference of the user. While sodium sulfate works well for soil and sediment samples, pelleted DE is good choice for wet tissue samples. The use of magnesium sulfate is not recommended in ASE, due to its potential for melting at higher temperatures. Sodium sulfate should not be used with methanol or other polar solvents, as it will be solubilized at elevated temperatures. Oven drying and freeze drying are other viable alternatives for drying samples containing inorganics prior to extraction; however, the recovery of volatile compounds may be compromised by these procedures. 11.1.4.5 Extraction parameters Solvent In order for an efﬁcient extraction to occur, the solvent must be able to solubilize the target analytes, while leaving the majority of the sample matrix intact. The polarity of the extraction solvent should therefore closely match that of the target compounds. Mixing solvents of differing polarities can be used to extract broader ranges of compound classes. Generally, if a particular solvent has been shown to work well in a conventional procedure, it will also work well in ASE. Compatibility with the post-extraction analytical technique, the need for

347

J. L. Ezzell

extract concentration (solvent volatility), and the cost of the solvent should all be considered. While many ASE methods recommend solvents or solvent mixtures for speciﬁc analyte classes, there may be alternatives which better ﬁt the needs of a particular laboratory. Solvents that exhibit marginal results at ambient conditions may perform adequately at higher temperatures. Most all liquid solvents, including water and buffered aqueous mixtures, can be used in ASE. Strong acids (HCl, HNO3, H2SO4) are not recommended for use, due to their ability to react with the stainless steel in most systems. When acidic conditions are required, weaker acids, such as acetic or phosphoric acid, should be used, and are normally added to aqueous or polar solvents in the 1–10% (v/v) range. When using the same solvent system as a traditional method, analyte recovery as well as co-extractable content should be very similar, although the ASE extract will be in a concentrated form. Therefore, if a traditional extraction approach requires an extract clean-up step prior to quantiﬁcation, the ASE extract will most likely require the same treatment. Temperature Temperature is the most important parameter used in ASE extraction. When developing a new method, it is recommended to start at 1008C, or if the target analytes have a known thermal degradation point, start at 208C below this level. An increase in temperature will generally have a positive impact on the extraction, but only up to the point where analyte or matrix degradation begins to occur. These problems, however, are generally not observed in the normal operating ranges of ASE (75–1508C). For sample matrices which tend to melt at elevated temperatures, cellulose extraction thimbles can be inserted into the extraction cells, in order to facilitate sample loading and removal. Pressure The effect of pressure is, ﬁrst, to maintain the solvents in their liquid states while above their atmospheric boiling points, and second, to move the ﬂuids through the system efﬁciently. The pressures normally used in ASE are well above the thresholds required to maintain the solvents as liquids (1000– 3000 psi, 6.9 –20.7 MPa), so adjustments for changing solvents are not required. Changing the pressure will have very little impact on analyte recovery, and it is not considered a critical experimental parameter. Most ASE extractions are performed between 1000 and 2000 psi (6.9 –13.8 MPa), with 1500 psi (10.3 MPa) the most common value. Cycles The use of static cycles was developed in order to introduce fresh solvent during the extraction process, which helps to maintain a favorable extraction equilibrium. This effectively approximates dynamic extraction conditions without the need for troublesome ﬂow restrictors to maintain pressure. When more than one cycle is used in a method, the ﬂush volume is divided

348

Accelerated solvent extraction of organometallic and inorganic compounds

by that number. When the ﬁrst static time is complete, the divided portion of the ﬂush volume is delivered to the cell, with the “used” solvent directed to the collection vial. The system then holds the sample and solvent for a second static period. The nitrogen purge step is initiated only after the ﬁnal static cycle. Since the original ﬂush volume has only been divided, no additional solvent is used for the extraction. Static cycles have proven useful for sample types with a very high concentration of analyte, or samples with difﬁcult to penetrate matrices. The static time can be adjusted to minimize the total extraction time. For example, 3, 3-min static cycles can be used in place of 1, 10-min static step. When low temperature extractions are desired (, 758C), multiple static cycles should be used to compensate for the lack of fresh solvent normally introduced during the heat-up step, as the static valve pulses to regulate the pressure. Time Certain sample matrices can retain analytes bound, encapsulated or otherwise trapped within pores or other structures. Increasing the static time at elevated temperatures can allow these compounds to diffuse into the extraction solvent. The effect of static time should always be explored in conjunction with static cycles, in order to produce a complete extraction in the most efﬁcient way possible. 11.1.5 Application areas 11.1.5.1 Extraction of arsenicals and arsenic speciation Due to the species dependent toxicity of arsenic, speciation of arsenicals is a growing area of interest in risk assessment research. While the inorganic species, arsenite (AsIII) and arsenate (AsV), have been categorized as carcinogenic, the methylated forms, mono-methylarsonic acid (MMA) and dimethylarsinic acid (DMA), have more recently been identiﬁed as possible cancer promoters. It is believed that arsenobetaine (AsB) and arsenocholine (AsC), which are substituted organoarsenicals, are relatively non-toxic. Two major sources of arsenic exposure are drinking water and dietary ingestion. According to the 1986–1991 FDA Total Diet Study [5], typical daily intake of aresenic is 50 ug/day for a normal adult, with seafood as the major source of total arsenic exposure. Therefore, the extraction of arsenicals from marine samples is an area of increasing environmental signiﬁcance. Due to the anionic, cationic, and neutral nature of the arsenic compounds, quantiﬁcation can be performed using ion exchange and reversed phase chromatography, although AAS and ICP-MS are preferred due to the sensitivity levels which can be obtained. Selection of an extraction solvent should be based, in part, on the extract analysis method to be employed. McKiernan [6] compared ASE and sonication extraction to traditional digestion procedures for the analysis of total arsenic from a variety of ﬁsh tissues. Vela [7] used ASE for the extraction of arsenic species present in freeze-dried carrots which were collected from

349

J. L. Ezzell

several areas around the United States. Carrot samples were mixed with Ottawa sand (acid rinsed to remove any arsenic contamination) and extracted using methanol –water mixtures at 1008C followed by LC– ICP-MS determination. Recoveries ranging from 84 to 111% were obtained for AsIII, Asv, MMA, DMA and AsB, at concentration levels from 24 to 16,200 ng/g. Gallagher [8,9] extracted arsenosugars using ASE with methanol/water (30/70) at ambient temperature. Extracts were cleaned using C18 SPE cartridges followed by analysis by IC-ICP-MS and IC-ESI-MS, which permitted additional structural information. 11.1.5.2 Direct extraction of metals Direct extraction of metals using ASE has been published by Sadik [10] using water/EDTA mixtures. Recovery of lead was found to be 97.3% of the reference value for a certiﬁed soil, with a relative standard deviation of 2.7% ðn ¼ 5Þ: This approach was also applied to the extraction of a range of toxic metals, including lead, manganese, nickel, gold, iron, copper, and cadmium. Recoveries ranged from 78 to 101% of certiﬁed values with a relative standard deviation of 5% ðn ¼ 5Þ: The total extraction solvent volume used was 13 ml for a 5 g sample. Sample matrices included industrial sludge, river sediments, coal ﬂy ash, and spent copper plating baths. The advantage of this approach was the complete elimination of concentrated acids from the preparation procedure. 11.1.5.3 Extraction of organometallic compounds Organotin compounds are used widely throughout the world as insecticides, fungicides, bactericides, acaricides, wood preservatives, and antifouling agents. They are therefore routinely found in water, sediments, biological tissues, and sewage sludge. Due to their high toxicity in marine organisms, the application of tributyltin (TBT) and triphenyltin (TPT) has been restricted in recent years. Despite these restrictions, TBT and TPT, as well as their major metabolites, dibutyltin (DBT), monobutyltin (MBT), diphenyltin (DPT), and monophenyltin (MPT), are still found in waters and sediments at signiﬁcant levels. Arnold [11] used ASE to extract organotins from freeze-dried sediments at 1008C using a solvent mixture of 1 M sodium acetate and 1 M acetic acid in methanol prior to GC –MS analysis. At spike levels of 10 and 1000 ng/g, recoveries ranged between 87 and 105%. Chiron [12] used a solvent mixture of methanol with 0.5 M acetic acid and 0.2% (w/v) tropolone prior to LC– ICP-MS analysis. For spiked samples, limits of detection ranged from 0.7 to 2.0 ng/g and relative standard deviations were found to be between 8 and 15%. 11.1.5.4 Extraction of phosphorus Phosphorus was extracted from lake sediments by Waldeback [13] by using a buffered dithionite solution at 258C followed by water at 1008C in sequential extractions. The ﬁrst extraction was designed to extract redox sensitive iron adsorbed phosphorus and the second designed to recover phosphorus

350

Accelerated solvent extraction of organometallic and inorganic compounds

compounds released by oxidative hydrolysis in the hot water. This approach was used to characterize the depth and extent of phosphorus presence in a natural lake water. 11.1.6 Summary Compared to conventional extraction times ranging from 4 to 48 h in length, ASE extractions are normally performed in 12–20 min per sample. While the decrease in extraction time is favorable for most laboratories in general, it can be critical for those industries where laboratory analytical data is used in feedback control of production cycles and manufacturing process quality control. The volume of solvents used in ASE can be 10– 20 times lower than traditional extraction methods. When factors such as safety and analyst exposure, solvent purchase and disposal costs, are considered, the beneﬁts of ASE can be quite substantial for most laboratories. When compared directly to traditional extraction methods, the recoveries generated by ASE normally equal or slightly exceed the alternative method. ASE has been extensively compared to conventional methods used in the environmental industry, and standard methods are available for most analyte classes [1,3,4]. Once an ASE method has been developed for a class of analyte, that same method can be successfully applied to a variety of matrix types without adjustment to extraction parameters. This lack of matrix dependency has allowed a very small set of standard methods to be applied to a large range of sample types. The increased temperatures used in ASE increase the efﬁciency of the extraction process. Generally, an increase in extraction temperature will result in an increase in the amount of material extracted from a sample matrix. There is a point, however, at which increased temperature begins to have a negative effect on analyte recovery. This is normally due to thermal degradation or thermally induced rearrangement of target compounds. Burning or charring of sample matrices can also occur as the temperature is increased. Extensive evaluations and comparisons of ASE have shown that, in the normal operating range of 75–1508C, the extraction efﬁciency is increased without these negative effects. As the temperature is increased towards the 2008C level, however, stability problems with analytes and matrices are more common. It seems unlikely, therefore, that future generation ASE systems will incorporate higher temperature capabilities, and methods development in high temperature extraction (.2008C) will be most likely be limited to very speciﬁc application areas. ASE is generally considered as an exhaustive extraction technique, and under the appropriate conditions all of the extractable content of a matrix will be solubilized. At the temperatures used in ASE, this will include the moisture content of the sample matrix. If the extraction solvent has a degree of miscibility with water (methanol, acetone, etc.) the analyst may not observe water in the collection vial. However, when non-polar solvents are used, an aqueous layer may be present. This is normally eliminated from the extract

351

J. L. Ezzell

prior to concentration or analysis by addition of sodium sulfate directly to the vial, shaking and decanting. In contrast to traditional extraction approaches, all of the basic steps of ASE are amenable to automation, freeing the analyst from the labor intensive nature of most sample preparation protocols. Automated ASE systems can extract up to 24 sample cells, and have built in the necessary safety considerations for unattended operation. This level of system operation results, however, in a capital cost higher than traditional glassware based approaches.

REFERENCES 1 2 3 4 5

6 7 8 9 10 11 12 13

352

B. Richter, J. Ezzell, D. Felix, K. Roberts and D. Later, Am. Lab., (February) (1995) 24–28. B. Richter, B. Jones, J. Ezzell, N. Porter, N. Avdalovic and C. Pohl, Anal. Chem., 68 (1996) 1033–1039. J. Ezzell, B. Richter, D. Felix, S. Black and J. Meikle, LC/GC, (May) 13(5) (1995) 390– 398. M. Schantz, J. Nichols and S. Wise, Anal. Chem., 69 (1997) 4210–4219. R. Schoof, L. Eickhoff, L. Yost, D. Crecelius, D. Cragin, D. Meacher and D. Menzel, Third International Conference on Arsenic Exposure and Health Effects. Elsevier, Amserdam, 1999, pp. 81–88. J. McKiernan, J. Creed, C. Brockhoff, J. Caruso and R. Lorenzana, J. Anal. At. Spectrom., 14 (1999) 607– 613. N. Vela, D. Heitkemper and K. Stewart, Analyst, 126 (2001) 1011 –1017. P. Gallagher, X. Wei, J. Shoemaker, C. Brockhoff and J. Creed, J. Anal. At. Spectrom., 14 (1999) 1829–1840. P. Gallagher, X. Wei, J. Shoemaker, C. Brockhoff and J. Creed, Fresnius J. Anal. Chem., 369 (2001) 71– 80. O. Sadik, A. Wanekaya and S. Myung, Analyst, 127 (2002) 3–7. C. Arnold, M. Berg, S. Mueller, U. Domman and P. Schwarzenbach, Anal. Chem., 70 (1998) 3094–3101. S. Chiron, S. Roy, R. Cottier and R. Jeannot, J. Chromatogr. A, 879 (2000) 137– 145. M. Waldeback, E. Rydin and K. Markides, Int. J. Environ. Anal. Chem., 72(4) (1998) 9 –17.

Chapter 12

Sonication as a sample preparation method for elemental analysisq Kevin Ashley

12.1

INTRODUCTION

Chemical effects induced by ultrasonic energy in solution can be employed for sample preparation purposes. Ultrasound consists of pressure waves exceeding 18 kHz; acoustic waves into the gigahertz range can be experimentally generated by mechanical means. When imparted to solutions, ultrasonic energy causes acoustic cavitation, or bubble formation and subsequent implosion [1,2]. The collapse of bubbles created by ultrasonication (usually abbreviated to “sonication”) of solutions generates extremely high local temperature and pressure gradients, conditions which have been employed extensively in sonochemistry [1 –5]. On a timescale of , 10210 s, effective local energies and pressure gradients of , 1 eV and , 104 atm, respectively, are generated as a result of bubble implosion [1–3]. Hence ultrasonic energy serves to create localized “hot spots” that form most readily at interfaces between phases, notably at junctions between solutions and solids. These energetic interfacial effects of ultrasonic energy can be easily observed in ultrasonic cleaners, which are used to remove oxide layers tarnishing jewelry. Concomitant with bubble collapse in aqueous solution is the generation of hydrogen peroxide and hydroxide radicals [6 –8]. These highly oxidative species aid signiﬁcantly in the use of ultrasound for analytical extraction purposes. Applications of ultrasonic extraction (UE) in sample preparation have shown considerable promise for speeding up and simplifying sample treatment procedures.

q Disclaimer: Mention of company or product names does not constitute endorsement by the Centers for Disease Control and Prevention. Note: This article was prepared by U.S. Government employees as part of their ofﬁcial duties, and legally may not be copyrighted in the United States of America.

Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

353

K. Ashley

Apart from chemistry speciﬁcally induced by cavitation, ultrasound offers a convenient means for the efﬁcient mechanical agitation of solutions. In this way mass transport in solution is greatly enhanced, thereby favoring increased reactivity. With or without cavitation, ultrasonic energy can often be used to quickly and easily perform effective sample dissolutions. For instance, ultrasonic energy has been used successfully to extract organic analytes from environmental matrices, and sonication forms the basis of a United States Environmental Protection Agency (EPA) method for the extraction of organic compounds from soil samples [9]. However, UE has been comparatively underutilized by analytical chemists for purposes of extracting metal species from solid sample matrices [10,11]. Concerning elemental determinations, the sample dissolution procedure is ordinarily the most time-consuming step of the overall analysis. For elemental analysis, standard methods for solid sample treatment ordinarily involve hot plate or microwave digestion or similar “wet ashing” techniques [12]. To assist with sample dissolution, decompositions using concentrated strong acids, and high temperatures and/or pressures (e.g., via microwave digestion), are routinely carried out in analytical laboratories. Many of these traditional sample preparation methodologies are described in other chapters of this book. As an alternative to these more conventional digestion techniques, in many cases UE can offer beneﬁts such as convenience, speed and safety, while also maintaining acceptable analytical recoveries for numerous elements in a variety of sample types. In this chapter, an overview of UE is provided, with an emphasis on the use of ultrasound to effect the dissolution of target elements for subsequent analytical determination. Speciﬁc examples are given where UE has been used successfully for analysis purposes. Although UE remains a ﬂedgling technique as far as elemental analysis is concerned, an increasing number of papers have recently appeared wherein ultrasonic energy has been utilized in inorganic analysis. The goals of this chapter are to provide the reader with some useful background on the subject of UE, and to shed light on various attributes as well as limitations of this sample dissolution technique. It is intended that after consideration of the material herein, the reader will be able to ascertain the potential of UE and adapt it to his/her own analytical application(s). 12.2

METHODOLOGICAL CONSIDERATIONS

The familiar laboratory ultrasonic bath has been used extensively for sample preparation purposes. Ultrasonic baths are available in a wide range of dimensions and power ratings, and many offer temperature controls. Baths are normally constructed so that ultrasonic transducers are fastened to the outer surface of the bottom and/or sides of the bath’s walls, enabling transmission of ultrasonic vibrations into the liquid within the bath. The most common

354

Sonication as a sample preparation method for elemental analysis

transducers for laboratory ultrasonic baths are comprised of piezoelectric materials, i.e., materials that mechanically expand and contract in response to an alternating applied electric ﬁeld. Some ultrasonic transducers are based on materials that expand and contract as a result of applied alternating magnetic ﬁelds; these are often seen in large-scale baths used for industrial purposes. Transducers are usually attached directly to the outside walls of the bath in such a manner that the walls themselves act as vibrating diaphragms. Typical laboratory ultrasonic baths are equipped with transducers providing vibrations in the frequency range of 20–40 kHz. For a more extensive and extremely useful discussion of ultrasonic transduction, instrumentation, and applications of ultrasound, the reader is referred to Berliner’s highly informative website: http://home.att.net/~Berliner-Ultrasonics. A critical parameter of interest for ultrasonic energy is power density, which is described in terms of the bath’s power rating (in Watts) and dimensions (square centimeters). Power densities of laboratory sonicators are ordinarily less than 1 W/cm2. For such “low power” ultrasonic devices, longer sonication times may be required to achieve complete dissolution of target analytes, e.g., lead [13]. High intensity ultrasound can be generated by means of ultrasonic horns, which serve to effectively amplify ultrasonic energy. These horns, as ultrasonic transduction tools, can be immersed into solution if properly contained within sealed housings [14]. The use of immersible ultrasonic transducers provides not only high intensities, but also allows for speciﬁc placement of ultrasonic probes in desired locations within reaction/extraction vessels. High ultrasonic intensities produce more widespread cavitation, which may or may not be necessary for sample dissolution purposes. High intensity ultrasonic probes have been used extensively in sonochemistry. An extremely informative overview on the subject of chemistry and ultrasound is presented in Suslick’s website: http://www.scs.uiuc.edu/~suslick. Vendors and manufacturers of ultrasonic devices can be accessed conveniently through the web page of the Ultrasonic Industry Association: http://www.ultrasonics.org. Besides the instrumental hardware that is necessary for carrying out UE (Fig. 12.1), different target analytes require a variety of extraction solutions and strengths. Extraction of heavy metals is typically performed in strong acid solutions and/or mixtures of acids (e.g., HCl, HNO3, H2SO4, HClO4), and varying acid concentrations may be necessary for different sample matrices [15]. As in hot plate digestion or microwave digestion, dissolution of silicate materials using laboratory ultrasonic baths requires the use of hydroﬂuoric acid (HF), but in the case of UE no deliberate heating is necessary [16]. UE of chromium is conveniently carried out in basic buffer solutions, and chromium speciation can be subsequently performed [17]. In some cases, UE with dilute acid can be used for on-site sample preparation using ﬁeld-portable instrumentation [11]. Factors which should be considered for choice of the UE dissolution solution(s) include not only the analyte(s) of interest, but also

355

K. Ashley

Fig. 12.1. Two basic ultrasonic systems for operation at frequencies of 20–80 kHz. Left: low-intensity bath system, ,1 to ,2 W/cm2; Right: high-intensity ultrasonic probe system, .100 W/cm2.

sample mass, particle size distribution, sample medium (e.g., soil, air ﬁlter, biological material, commercial product(s)), and so forth. Other sample preparation aspects must be considered besides the extraction procedure itself. For instance, in the case of some sample matrices such as soil or paint, signiﬁcant sample work up may be required prior to application of UE in order to homogenize the sample and reduce the mean particle size as much as possible. In other situations, isolation or separation steps must be conducted in order to separate the dissolved analyte of interest from other species that might interfere with subsequent instrumental analysis. While the focus of this chapter is on the extraction procedure itself, it is recognized that sample dissolution may represent only one segment of the overall sample preparation process (Fig. 12.2). In the following sections, a brief history of UE for purposes of elemental analysis is presented. Also, examples of recent applications of UE for elemental determinations in a variety of sample matrices are given.

Fig. 12.2. Schematic of overall sampling and analysis process employing UE. Dashed lines indicate where sample work up preliminary to extraction, and/or sample aliquot treatment subsequent to extraction, may possibly be bypassed.

356

Sonication as a sample preparation method for elemental analysis

12.3

HISTORICAL BACKGROUND

In an early application of UE, trace airborne lead collected onto ambient air ﬁlters was measured in an interlaboratory investigation which was overseen by the US EPA [18]. Airborne lead recoveries from ultrasonication in nitric acid were compared with recoveries obtained by hot plate digestion in this acid; several different instrumental analysis techniques were used. The performance of UE was found to be equivalent to that of hot plate digestion, and the procedure has become an EPA reference method for the measurement of lead in ambient air samples [19]. In a follow-up study by Harper et al., UE was applied to the dissolution of heavy metals from a National Institute of Standards and Technology (NIST) Standard Reference Material (SRM) [20]. Urban Particulate Matter, SRM #1648, was subjected to UE in various acid mixtures for different time durations within a laboratory ultrasonic bath. Experimental variables were changed systematically in order to optimize observed recoveries of numerous heavy metals; metal species were determined by atomic spectrometry. It was found that sonication for 1 h in concentrated HNO3 at elevated temperature provided quantitative recoveries for up to 10 elements having certiﬁed NIST SRM concentrations. But despite the success of this seminal work, few additional investigations of UE for elemental dissolutions appeared in the literature in the next several years. Some important early work on UE for elemental dissolution was carried out in Russia. Investigations by Kumina et al. demonstrated the utility of UE for the extraction of several metals from plant tissues and soils [21,22]. Elemental recoveries from UE in nitric – hydrochloric acid mixtures were compared against results obtained from a standard digestion procedure. Recoveries of ﬁve elements (Cu, Zn, Mn, Co, Ni) were discovered to be comparable to the recoveries obtained using the established sample preparation procedure. Also, good recoveries by UE were obtained from reference material soils. The authors lauded the simplicity and efﬁciency of the ultrasonic dissolution method. In France, UE has been used for years in the measurement of metals in workplace air samples [23]. The French method speciﬁes UE with nitric and hydroﬂuoric acids [24]. HF is used in addition to HNO3 since it is desired to measure metal species that are bound up in silicate materials, and would not otherwise be soluble, even in other acid solutions. The dissolution procedure is carried out directly within the sampler, which consists of a plastic sampling cassette housing a quartz ﬁber ﬁlter. Air particles are collected onto the ﬁlter by means of a low-volume, battery-powered sampling pump. Nitric and hydroﬂuoric acids are introduced into the sampler, which is then immersed into an ultrasonic bath and subjected to sonication. Metals in the extract are subsequently measured by inductively coupled plasma-atomic emission spectrometry (ICP-OES).

357

K. Ashley

The French UE HF/HNO3 method was used to measure a suite of metals in aerosols collected from a variety of work environments, where it was shown that a signiﬁcant portion of the collected airborne particulate matter was retained on the inside walls of the samplers [25]. Sonication within the sampler afforded a means to perform the extraction in such a way that particles collected within the sampler, but not retained on the ﬁlter, were nonetheless accounted for. In this way, sample losses which would have resulted from extraction and analysis of the ﬁlter alone were avoided.

12.4

APPLICATIONS—SONICATION AND SAMPLE PREPARATION

Rather surprisingly, ultrasonic energy for extraction purposes has been largely overlooked by analytical chemists as a sample preparation technique for elemental analysis [10]. However, some workers have recognized the potential of UE for dissolving target elemental analytes, and have engaged in investigations of UE for elemental determination in a variety of sample types, e.g., environmental and biological. In the following sections, examples of applications of UE for elemental analysis are presented, and successes as well as limitations are highlighted. 12.4.1 Environmental analysis Several groups have reported on investigations of UE in efforts to reduce sample preparation times for elemental analysis of soils, sediments, and related samples. In recent work, Bendicho et al. used focused ultrasound (by means of ultrasonic horns) for agitation purposes in sequential extraction procedures that were applied to sludge and sediment samples [26,27]. Elemental recoveries were determined by atomic spectrometry after ultrasonic irradiation. For sewage sludge samples, UE was applied to “acid soluble,” “reducible,” and “oxidizable” sample fractions [26]; for sediments, UE was used on “exchangeable,” “carbonate-bound,” “oxide-bound,” and “organic-bound” fractions [27]. Following optimization of sonication parameters (i.e., power and time) for each of the stages of the extraction, elemental recoveries were compared to recoveries obtained by the conventional mechanical shaking technique. In the case of sewage sludge samples, for each fraction, elemental recoveries of Cu, Cr, Ni, Pb, and Zn after sonication compared extremely well with recoveries of these metals using the conventional method [26]. On the other hand, UE of river sediment samples subjected to a sequential extraction treatment gave varying results for different metals and different fractions [27]. Yet total metal recoveries after UE were quantitative. In related work, sequential extraction schemes using ultrasonic agitation were evaluated for the dissolution of Cu, Mn, Zn, Fe [28], and Sr [29] in soil and sediment samples.

358

Sonication as a sample preparation method for elemental analysis

Recoveries of Cu, Mn, Zn, and Sr after UE of each fraction were comparable to recoveries obtained using conventional means. However, Fe recoveries from UE of various fractions differed greatly from recoveries of this metal after using the conventional mechanical shaking technique. These disparate results indicate that design-based sample preparation procedures (i.e., procedures that are speciﬁed vs. performance-based) can be successfully modiﬁed in some instances by using ultrasonic agitation, but not in others. For many situations, UE is well suited for quantitative extraction of several elements from environmental media. UE of soil samples immersed in concentrated strong acids and mixtures thereof has been found to give good recoveries for several metals, including As, Cd, Cu, Mn, Pb, and Zn [15,16,21, 30]. As mentioned previously, use of HF is required for complete dissolution of silicate-bound metal species [16,25] (including radionuclides [31]), but the use of this acid is often avoided owing to its highly hazardous nature. Ultrasoundassisted extractions by means of slurry suspensions have been shown to yield high elemental recoveries from samples of sediments and soils [32,33]. Also, UE of soil samples in slightly basic buffer solution has been demonstrated to give acceptable recoveries of Cr(VI) [34]. In general, efﬁciency of UE decreases with increasing particle size and/or organic content [35,36], so these factors relating to the sample medium must be considered and dealt with in sample preparation strategies. It must be noted that while good elemental recoveries may be obtained from certiﬁed reference materials (which have been ground to small particle sizes), poor recoveries could be obtained from real-world samples comprised of larger particles. Also, longer extraction times may be required for samples having high organic contents. UE is ideally suited to the extraction of metals from airborne particles collected onto ﬁlters [15,16,18]. This is largely due to the smaller particle size of airborne particulate matter, which enables efﬁcient elemental extraction by sonication. About a decade ago, Carneiro et al. [37] employed sonication in a mixture of nitric, sulfuric, and hydroﬂuoric acids to extract Ni, Sb, and V from air ﬁlter samples. These workers reported quantitative recoveries of these three elements from NIST SRM ﬂy ash and urban particulate matter. In a thorough Finnish study, UE of ﬁlter materials spiked with NIST SRMs was performed in aqua regia and also in acid solutions containing HF; atomic spectrometry was used for instrumental analysis of extracts [38]. Quantitative recoveries were found for many elements when HF is used during the extraction; yet this observation is not terribly surprising, for the method essentially constitutes a modiﬁcation of the procedure used by the INRS in France [23,24]. In an extensive investigation by Eyckmans et al. [39], sonication was used for the purpose of measuring trace metals and inorganic ions in airborne particulate ﬁlter samples. In this work, results from UE were compared to data obtained using a “leaching” method. After extractions, atomic spectrometry was used for instrumental elemental analyses, while extracted ions were measured by ion chromatography [39]. It was reported that

359

K. Ashley

elemental recoveries from UE were generally greater than those obtained by the leaching technique. In another related study, UE in acid solution was used to dissolve Cd, Fe, Pb, and Zn collected from atmospheric aerosols onto cellulose ﬁlters; anodic stripping voltammetry (ASV) was employed for elemental determinations [40]. This UE/ASV procedure was used previously for the determination of lead in air ﬁlter samples [41]. The method was tried on ambient air samples in order to evaluate ultrasonic dissolution of additional elements (besides Pb) which can be monitored by ASV. Elemental recoveries obtained from UE/ASV or by neutron activation analysis were compared to recoveries from a reference method which employed a high-temperature digestion technique and elemental determination by atomic spectrometry [40]. Similar to what has been observed in UE of environmental CRMs (mentioned previously), good recoveries by UE/ASV were observed for Cd, Pb, and Zn. However, monitoring the ionic content of ultrasonic extracts can be problematic owing to anion production during the sonication process [42]. In a great body of work carried out in Compton’s group at Oxford, ultrasonic energy has been combined with electroanalytical techniques in efforts to increase mass transport and alleviate electrode passivation [43,44], thereby opening up new possibilities for studies of electrode reactions and applications of electroanalysis. In recent work, sonoelectrochemical methods employing modiﬁed electrodes have been used to increase the sensitivity of trace metal analysis in environmental samples [45,46]. Sonoelectroanalysis has also been successfully accomplished in the presence of surfactants which have impeded electrochemical detection in the past [47]. It should be a simple matter to devise sonoelectrochemical cells wherein UE and electroanalysis can be performed almost simultaneously. The ﬁeld of sonoelectrochemistry could revitalize electroanalysis, and offers considerable promise for future environmental analysis applications. Table 12.1 gives a summary of UE for elemental determinations in environmental applications. 12.4.2 Industrial hygiene In recent years, UE has seen increasing application as a dissolution technique for airborne elemental species that are sampled in occupational environments. In a large study by Butler and Howe, UE of laboratory-generated aerosol samples deposited onto ﬁlter materials was evaluated in an interlaboratory trial [48]. One purpose of this round-robin trial was to evaluate the UE technique as a sample preparation method, prior to its inclusion in an international standard for the determination of metals and metalloids in workplace air. Compared to results from various hot plate and microwave digestion protocols, UE gave satisfactory recoveries of a number of elements, although HF was used in the dissolutions [48]. A subsequent study examined elemental

360

Sonication as a sample preparation method for elemental analysis TABLE 12.1 Ultrasonic extraction of environmental samples for elemental analysis Matrix(es)

Elemental analyte(s)

Reference(s)

Soil Soil Soil Soil Soil

As, Cd, Cu, Mn, Pb, Zn Co, Cu, Mn, Ni, Zn Cu, Fe, Mn, Zn Sr Al, Co, Cr, Cu, Fe, Mn, Ni, Pb, Sr, Zn Cr(VI) Al, Ca, Cu, Fe, Mg, Mn, P, S Radionuclides As, Cd, Cr, Cu, Ni, Pb Cd, Cr, Cu, Mn, Ni, Pb, Zn As, Ba, Cd, Co, Cr, Cu, Fe, Mg, Mn, Ni, Pb, V, Zn Cu, Cr, Ni, Pb, Zn As, Ba, Cd, Co, Cr, Cu, Fe, Mn, Mo, Ni, Pb, Sr, Ti, V, Zn Ni, Sb, V

[21] [22] [28] [29] [30]

Soil Soil Soil, sediment Soil, sediment Soil, sludge, sediment Soil, sediment, urban particulate matter Sludge, sediment Urban particulate matter Urban particulate matter, ﬂy ash, air ﬁlters Urban particulate matter, ﬂy ash, air ﬁlters Air ﬁlters, ﬂy ash Airborne particles Airborne particles Airborne particles

Al, As, Be, Ca, Cd, Co, Cr, Cu, Fe, Mg, Mn, Mo, Ni, Pb, Sb, Sr, Ti, Tl, U, V, Zn Cr(VI) Pb Al, Ca, Cl, Cu, F, Fe, K, Na, Pb Cd, Fe, Pb, Zn

[34] [35] [31] [32] [16] [15] [26,27] [20] [37] [38]

[17] [18] [39] [40]

recoveries from air ﬁlter samples using UE in acid solutions sans HF [15]. It was discovered that several elements captured on air ﬁlter materials could be measured quantitatively, with recoveries for most metals increasing as the acid concentration was increased. UE has also been employed in efforts to develop ﬁeld-portable monitoring methods for various metallic elements that may be present in toxic concentrations in workplace air. A ﬁeld-portable UE/ASV method was evaluated for the on-site determination of lead in workplace air samples [49]. Ultrasonic baths are easily transported to ﬁeld sites, unlike the conventional “high heat” sample preparation equipment. Also, portable hand-held electroanalytical devices for measuring lead are commercially available. It was found

361

K. Ashley

that the UE/ASV method performed equivalently to the reference analytical method, i.e., hot plate strong acid digestion and atomic spectrometric measurement of lead [49]. This method, which was later tested in an interlaboratory evaluation [50] and in a ﬁeld trial [51], has been promulgated in the NIOSH Manual of Analytical Methods as a deﬁnitive procedure for the on-site monitoring of workplace lead exposures [52]. UE is also included as an optional technique for lead dissolution in a newly-published ASTM standard which describes occupational sampling and analysis of airborne lead [53]. Workplace monitoring of hexavalent chromium, Cr(VI), is of concern due to the carcinogenic properties of this species when it is inhaled. To enable on-site monitoring of Cr(VI), a ﬁeld-portable method for this species in workplace air samples was developed and evaluated [54]. The method entails the following steps: (1) sampling of airborne Cr(VI) onto inert ﬁlter materials using battery-powered personal sampling pumps; (2) UE of the ﬁlters in basic buffer solution in order to dissolve Cr(VI); (3) solid-phase extraction of extract aliquots to isolate Cr(VI) from Cr(III) and other metal cations; (4) elution of isolated Cr(VI), and reaction of the eluate with 1,5-diphenylcarbazide; and (5) determination of the highly colored Cr-diphenylcarbazone product using ﬁeld-portable spectrophotometry. The portable method was validated using performance evaluation materials consisting of welding fume samples having certiﬁed Cr(VI) loadings [54]. The method was further ﬁeldtested on-site in a workplace environment, where Cr(VI) exposures were measured as a function of work task [55]. Also, the performance of the ﬁeld method was compared to that of reference ﬁxed-site laboratory methods for the determination of airborne Cr(VI) [56]. This work demonstrated the utility of UE to effect the quantitative dissolution of Cr(VI) from airborne particulate matter. Table 12.2 summarizes applications of UE for elemental determinations in the industrial hygiene ﬁeld.

TABLE 12.2 Ultrasonic extraction of industrial hygiene samples for elemental analysis Matrix(es)

Elemental analyte(s)

Reference(s)

Air ﬁlters Air ﬁlters Air ﬁlters Air ﬁlters Air ﬁlters, indoor dust Air ﬁlters, indoor dust

Cr(VI) Pb As, Be, Cl, Cr, Cu, F, Fe, Mn, Ni, Pb, Sb, Sn, Zn Cd, Co, Cr, Cu, Fe, Mn, Mo, Ni, Sn,W, Zn As, Ba, Cd, Co, Cr, Cu, Fe, Mg, Mn, Ni, Pb, V, Zn Pb

[15,54,56] [49] [25] [48] [15] [50,51]

362

Sonication as a sample preparation method for elemental analysis

12.4.3 Biological tissues and ﬂuids UE has been recognized to be an efﬁcient tool for elemental extraction from materials of biological origin, and numerous research papers on the subject have appeared in the recent literature. In one study, sonication of biological samples for 40 min was performed using a commercial 20-kHz ultrasonic probe [57]; a sulfuric acid/hydrogen peroxide solution was used to dissolve heavy metals in NIST SRM oyster tissue and pine needles. Atomic spectrometry was used for instrumental analysis. Good recoveries of Cd, Cu, Mn, Sr, and Zn were obtained, although some metal contamination emanating from the titanium probe tip of the ultrasonic horn was detected [57]. In other work, mussel tissue samples were subjected to sonication for 2 h in mixtures of hydrogen peroxide and nitric and hydrochloric acids [58]; this was followed by elemental analysis using atomic spectrometric techniques. Quantitative recoveries were reported for numerous elements: Ca, Cu, Fe, K, Mg, Mn, Na, V, and Zn; Cd, Co, Cr, Rb, and Se were partially dissolved. These workers noted that control of ultrasonic bath temperature was unimportant to elemental recoveries, and they praised the speed, simplicity, and safety aspects of the UE procedure [58]. Related studies have reported on UE of arsenicals in ﬁsh [59] and seafood [60], Se in seafood [61], and various elements in lobster tissue [62]. Various attributes of sonication which have already been mentioned were cited in these papers. Lima et al. conducted an extensive study [63] in which an ultrasonic probe was used to assist in extracting trace Cd, Cu, and Pb from numerous reference materials, many of biological origin. Results from ultrasound-assisted extraction were compared to data obtained from slurry sampling and microwave digestion. The ultrasound-assisted procedure was reported to give recoveries of these metals that agreed with certiﬁed concentrations, and served to dramatically reduce sample preparation times [63]. Related work has been conducted in Bendicho’s group to ultrasonically extract cadmium [64] and lead [65] from biological materials, including animal tissues. In work by Caroli et al. [66], samples of honey were subjected to sonication, and numerous elements were determined by ICP-AES and inductively coupled plasma-mass spectrometry (ICP-MS). Ranges of trace levels of As, Cd, Cr, Cu, Fe, Mn, Ni, Pb, Pt, Sn, and Zn in honey samples from various sources were reported. In a thorough investigation, Bermejo-Barrera et al. [67] used a PlackettBurman design to optimize UE conditions for Ca, Cu, Fe, Mg, Mn, and Zn from samples of human hair. Experimental variables were optimized for seven factors, including nitric acid, hydrochloric acid, and hydrogen peroxide concentrations, sonication time and temperature, solution volume, and hair particle size. Certiﬁed reference materials were also analyzed using the optimized method. In contrast with some other studies of UE, temperature was found to be a crucial variable [67]. It was also discovered that dilute acid could be used for efﬁcient metal extraction from hair samples by ultrasound.

363

K. Ashley

UE of elements from plant tissues has been reported by several groups. Earlier investigations demonstrated the utility of ultrasound for extracting iron [68,69] and copper [69] from plant materials. In a series of papers, Bendicho and co-workers [70 –73] recently investigated UE of elements from plant tissues. Sonication in acid solutions was found to give acceptable recoveries of several heavy metals (Cu, Mn, Mg, Pb, Zn) from plant samples [70,71], and good analytical precision was achieved. UE in basic medium containing ethylenediaminetetraacetic acid (EDTA) also gave good recoveries of several elements (Ca, Cd, Mg, Mn, Pb, Zn) from plant tissues [72]. Arsenic speciation in plant certiﬁed reference materials was also demonstrated using an ultrasonically assisted procedure [73]. In related work, UE was applied to the extraction of heavy metals from sprouts [74] and vegetables [75]. UE as a sample preparation method can be useful in assessing heavy metal contamination in plant foods meant for human consumption. A summary of applications of UE for elemental determinations in biological matrices is given in Table 12.3.

12.4.4 Other applications Some additional applications of UE have been reported besides those mentioned thus far. UE has been employed as a ﬁeld-portable sample preparation procedure for the on-site determination of lead in paint [13,49–51,76,77] and dust wipe [78] samples. An ASTM UE method for lead has been published [79]. Lead extractability problems associated with large particle sizes [13,51,77], sample mass [13], and high organic content [13,78] were cited; the importance of these factors for UE of soil and sediment samples was already addressed above. In other work, UE has been employed in metallurgical applications for the dissolution of nickel [80] and chromium [81] from ore samples. The authors pointed to the potential of UE for speeding up assays of metallurgical interest. Elsewhere, ultrasound was used to recover metals from electrolytes [82]; these metals can exist as by-products of industrial processes. UE was proposed as a method to improve recoveries of metals from electrolyte solutions, enabling recycling of the metals. Sonication of metal surfaces and particles in solution results in metallic deposition onto electrode surfaces [83], an observation which could offer metallurgical and industrial applications. In an interesting application, recording discs were subjected to UE in acid solution in order to recover metals from the recording media [84]. Ultrasonic irradiation was found to be an effective means for extracting metal species from recording media. Table 12.4 presents an overview of other applications of UE for elemental analysis besides those already summarized in Tables 12.1– 12.3.

364

Sonication as a sample preparation method for elemental analysis TABLE 12.3 Ultrasonic extraction of biological samples for elemental analysis Matrix(es)

Elemental analyte(s)

Reference(s)

Plants Plants Plants

Cu, Co, Mn, Ni, Zn Fe Ca, Cd, Cu, Fe, Mn, Mg, Pb, Zn Al, Cd, Cr, Cu, Fe, Ni, Pb Arsenicals Cd, Cu, Mn, Sr, Zn Cu Ca, Mg, Mn Ca, Co, Cr, Cu, Cs, Fe, K, Mg, Mn, Na, Rb, Sc, Se, V, Zn Cd, Pb, Cu

[22] [68] [70–72]

Plants Plants, ﬁsh Pine needles, oyster tissue Palm oil Vegetables Mussel tissue Mussel tissue, liver tissue, grass, bread, plankton, ﬁsh, etc. Mussel tissue, liver tissue, ﬁsh, plants Mussel tissue, plants, ﬁsh tissues Fish Seafood Seafood Lobster tissue Honey Hair

[74] [73] [57] [69] [75] [58] [63]

Cd

[64]

Pb Arsenicals As Se As, Cd, Co, Mo, Se As, Cd, Cr, Cu, Fe, Mn, Ni, Pb, Pt, Sn, Zn Ca, Cu, Fe, Mg, Mn, Zn

[65] [59] [60] [61] [62] [66] [67]

TABLE 12.4 Ultrasonic extraction for elemental determinations in paints, ores, products, etc. Matrix(es)

Elemental analyte(s)

Reference(s)

Paint chips Dust wipes Metallurgical samples Ores, “beneﬁciated” products Electrolytic baths Recording media

Pb Pb Ni Cr Cu, Zn Co, Fe, Tb

[13,51,76,77] [78] [80] [81] [82] [84]

365

K. Ashley

12.5

SUMMARY

Although UE has largely been underutilized by analytical chemists, many laboratories are beginning to recognize some of the attributes of ultrasonic energy for sample preparation purposes. In many cases, sonication can be applied for short time periods in order to simply, quickly, safely, and effectively extract elemental species for their subsequent analytical determination. UE has shown promise for elemental analysis of environmental, workplace, biological, and industrial samples, and many applications remain unexplored to date. In instances where ultrasound alone is insufﬁcient to effect quantitative extraction of target elements, sonication can be used in concert with other traditional sample preparation techniques in order to help speed up the overall sample preparation process. Aside from other beneﬁts already noted, the use of UE is often less expensive than traditional high-temperature and/or pressurized equipment (e.g., microwave digestion apparatus) that is used for preparation of samples. Although there are limitations (e.g., problems due to particle size effects, limits on sample mass, organic interferences), it is predicted that UE will become a more commonly used technique for routine sample preparation purposes in the analytical laboratory.

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13

366

K.S. Suslick (Ed.), Ultrasound: Its Chemical, Physical and Biological Effects. VCH Publishers, Weinheim, Germany, 1988. K.S. Suslick, Y.T. Didenko, M.M. Fang, T. Hyeon, K.J. Kolbeck, W.B. McNamara, M.M. Mdleleni and M. Wong, Philos. Trans. R. Soc. Lond. A, 357 (1999) 335. K.S. Suslick, Kirk-Othmer Encyclopedia of Chemical Technology, 4th edn, Vol. 26. Wiley, New York, 1998, p. 517. T.J. Mason and J.-L. Luche, in: R. Van Eldick and C.D. Hubbard (Eds.), Chemistry Under Extreme or Non-classical Conditions. Wiley, New York, 1996, p. 317. M.A. Margulis, Sonochemistry and Cavitation. Gordon and Breach, Newark, NJ, 1995. I. Hua and M.R. Hoffman, Environ. Sci. Technol., 31 (1997) 2237. S.R. Souza, M. Korn, L.R.F. de Carvalho, M.G.A. Korn and M.F.M. Tavares, Ultrasonics, 36 (1998) 595. H. Destaillats, T.M. Lesko, M. Knowlton, H. Wallace and M.R. Hoffman, Ind. Eng. Chem. Res., 40 (2001) 3855. EPA SW-846 Method #3550, Test Methods for the Analysis of Solid Wastes. US Environmental Protection Agency (EPA), Washington, DC, 1994 M.D. Luque de Castro and M.P. da Silva, Trends Anal. Chem., 16 (1997) 16. K. Ashley, Trends Anal. Chem., 17 (1998) 366. J.A. Dean, Analytical Chemistry Handbook. McGraw-Hill, New York, 1995, p. 1.29. W.J. Rossiter, B. Toman, M.E. McKnight and M.B. Anaraki, Factors Affecting Ultrasonic Extraction of Lead from Laboratory-Prepared Household Paint Films (NIST Internal Report #6834). National Institute of Standards and Technology (NIST), Gaithersburg, MD, 2002.

Sonication as a sample preparation method for elemental analysis 14 15 16 17 18 19 20 21 22 23

24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47

D. Ensminger, Ultrasonics: Fundamentals, Technology, Applications. Marcel Dekker, New York, 1988. K. Ashley, R.N. Andrews, L. Cavazos and M. Demange, J. Anal. At. Spectrom., 16 (2001) 1147. J. Sa´nchez, R. Garcı´a and E. Milla´n, Analusis, 22 (1994) 222. J. Wang, K. Ashley, E.R. Kennedy and C. Neumeister, Analyst, 122 (1997) 1307. S.J. Long, J.C. Suggs and J.F. Walling, J. Air Pollut. Control Assoc., 29 (1979) 28. US EPA, 40 CFR Part 50 (Appendix G), Fed. Regist., 44 (1979) 564. S.L. Harper, J.F. Walling, D.M. Holland and L.J. Pranger, Anal. Chem., 55 (1983) 1553. D.M. Kumina, A.V. Karyakin and I.F. Gribovskaya, J. Anal. Chem. USSR, 40 (1985) 930. D.M. Kumina, E.N. Savinova, T.V. Shumskaya, M.D. Alybaeva and A.V. Karyakin, J. Anal. Chem. USSR, 44 (1989) 459. M. Demange, Me´thode de Mise en Solution des Poussie`res Pre´leve´es en Vue de Leur Analyse. Institut National de Recherche et de Se´curite´ (INRS), Vandœuvre-lesNancy, France, 1988, (internal report). INRS, Evaluation de l’exposition professionelle: Me´thodes de pre´le`vement et d’analyse de l’air (Fiche 003—Me´taux-Me´talloı¨des). INRS, Paris, 2000. M. Demange, J.C. Gendre, B. Herve´-Bazin, B. Carton and A. Peltier, Ann. Occup. Hyg., 34 (1990) 399. B. Pe´rez-Cid, I. Lavilla and C. Bendicho, Anal. Chim. Acta, 360 (1998) 35. B. Pe´rez-Cid, I. Lavilla and C. Bendicho, Int. J. Environ. Anal. Chem., 73 (1999) 79. C.M. Davidson and G. Delevoye, J. Environ. Monit., 3 (2001) 398. A. Elik and M. Akc¸ay, Int. J. Environ. Anal. Chem., 80 (2001) 257. R. Al-Merey, M.S. Al-Masri and R. Bozou, Anal. Chim. Acta, 452 (2002) 143. D. Dale, G. Brooks and M. Monagle, J. Radioanal. Nucl. Chem., 236 (1998) 199. W. Klemm and G. Bombach, Fresenius J. Anal. Chem., 353 (1995) 12. L. Amoedo, J.L. Capelo, I. Lavilla and C. Bendicho, J. Anal. At. Spectrom., 14 (1999) 1221. B.R. James, J.C. Petura, R. Vitale and G.R. Mussoline, Environ. Sci. Technol., 29 (1995) 2377. J. Eriksen, R.D.B. LeFroy and G.J. Blair, Soil Biol. Biochem., 27 (1995) 1005. G. Zhang and I. Hua, Chemosphere, 46 (2002) 59. M.C. Carneiro, R.C. Campos and A.J. Curtius, Talanta, 40 (1993) 1815. L.M. Jalkanen and E.K. Ha¨sa¨nen, J. Anal. At. Spectrom., 11 (1996) 365. K. Eyckmans, J. Zhang, J. DeHoog, P. Joos and R. Van Grieken, Int. J. Environ. Anal. Chem., 80 (2001) 227. M. Ochsenku¨hn-Petropoulou and K.-M. Ochsenku¨hn, Fresenius J. Anal. Chem., 369 (2001) 629. K. Ashley, Electroanalysis, 7 (1995) 1189. L.R.F. Carvalho, S.R. Souza, B.S. Martinis and M. Korn, Anal. Chim. Acta, 317 (1995) 171. R.G. Compton, J.C. Eklund and F. Marken, Electroanalysis, 9 (1997) 509. A.J. Saterlay and R.G. Compton, Fresenius J. Anal. Chem., 367 (2000) 308. A.M.O. Brett, C.M.A. Brett, F.-M. Matysik and S. Matysik, Ultrason. Sonochem., 4 (1997) 123. Y.C. Tsai, J. Davis, R.G. Compton and N. Ono, Electroanalysis, 13 (2001) 7. J.L. Hardcastle, G. Hignett, J.L. Melville and R.G. Compton, Analyst, 127 (2002) 518.

367

K. Ashley 48 49 50 51 52

53

54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76

77 78

368

O.T. Butler and A.M. Howe, J. Environ. Monit., 1 (1999) 23. K. Ashley, K.J. Mapp and M. Millson, Am. Ind. Hyg. Assoc. J., 59 (1998) 671. K. Ashley, R. Song, C.A. Esche, P.C. Schlecht, P.A. Baron and T.J. Wise, J. Environ. Monit., 1 (1999) 459. A. Sussell and K. Ashley, J. Environ. Monit., 4 (2002) 156. P.M. Eller and M.E. Cassinelli (Eds.), NIOSH Manual of Analytical Methods, 4th edn, Method No. 7701. National Institute for Occupational Safety and Health (NIOSH), Cincinnati, OH, 1994, 1998 suppl. ASTM D6785, Standard Test Method for Determination of Lead in Workplace Air Using Flame or Graphite Furnace Atomic Absorption Spectrometry. ASTM, West Conshohocken, PA, 2002. J. Wang, K. Ashley, D. Marlow, E.C. England and G. Carton, Anal. Chem., 71 (1999) 1027. D. Marlow, J. Wang, T.J. Wise and K. Ashley, Am. Lab., 32(15) (2000) 28. J.M. Boiano, M.E. Wallace, W.K. Sieber, J.H. Groff, J. Wang and K. Ashley, J. Environ. Monit., 2 (2000) 329. S. Mamba and B. Kratochvil, Int. J. Environ. Anal. Chem., 60 (1995) 295. H. El Azouzi, M.L. Cervera and M. de la Guardia, J. Anal. At. Spectrom., 13 (1998) 533. J.W. McKiernan, J.T. Creed, C.A. Brockhoff, J.A. Caruso and R.M. Lorenzana, J. Anal. At. Spectrom., 14 (1999) 607. C. Santos, F. Alava-Moreno, I. Lavilla and C. Bendicho, J. Anal. At. Spectrom., 15 (2000) 987. H. Mendez, F. Alava, I. Lavilla and C. Bendicho, Anal. Chim. Acta, 452 (2002) 217. J.A. Brisbin and J.A. Caruso, Analyst, 127 (2002) 921. ´ .C. Lima, F. Barbosa, F.J. Krug, M.M. Silva and M.G.R. Vale, J. Anal. At. E Spectrom., 15 (2000) 995. J.L. Capelo, I. Lavilla and C. Bendicho, J. Anal. At. Spectrom., 13 (1998) 1285. L. Amoedo, J.L. Capelo, I. Lavilla and C. Bendicho, J. Anal. At. Spectrom., 14 (1999) 1221. S. Caroli, G. Forte, A.L. Iamiceli and B. Galoppi, Talanta, 50 (1999) 327. P. Bermejo-Barrera, O. Mun˜oz-Naveiro, A. Moreda-Pin˜eiro and A. BermejoBarrera, Forensic Sci. Int., 107 (2000) 105. F. La´zaro, M.D. Luque de Castro and M. Valca´rcel, Anal. Chim. Acta, 242 (1991) 283. M.I. Saleh, M.S. Jab, I.A. Rahman and S. Norasiah, Analyst, 116 (1991) 743. I. Lavilla, B. Pe´rez-Cid and C. Bendicho, Int. J. Environ. Anal. Chem., 72 (1998) 47. A.V. Filgueiras, J.L. Capelo, I. Lavilla and C. Bendicho, Talanta, 53 (2000) 441. A.V. Filgueiras, I. Lavilla and C. Bendicho, Fresenius J. Anal. Chem., 369 (2001) 451. J.L. Capelo, I. Lavilla and C. Bendicho, Anal. Chem., 73 (2001) 3732. A. Elik, M. Akc¸ay and M. So¨kmen, Int. J. Environ. Anal. Chem., 77 (2000) 133. C. Nascentes, M. Korn and M.A.Z. Arruda, Microchem. J., 69 (2001) 37. U.S. EPA, Evaluation of the Performance of Reﬂectance and Electrochemical Technologies for the Measurement of Lead in Characterized Paints, Bulk Dusts, and Soils (EPA 600/R-95-093). U.S. EPA, Research Triangle Park, NC, 1996. K. Ashley, M. Hunter, L.H. Tait, J. Dozier, J.L. Seaman and P.F. Berry, Field Anal. Chem. Technol., 2 (1998) 39. K. Ashley, W. Mercado, T.J. Wise and D.B. Parry, J. Hazard. Mater., 83 (2001) 41.

Sonication as a sample preparation method for elemental analysis 79

80 81 82 83 84

ASTM E1979, Standard Practice for Ultrasonic Extraction of Lead in Paint, Dust, Soil, and Air Samples for Subsequent Determination of Lead. ASTM, West Conshohocken, PA, 2002. B. Pesic and T. Zhou, Metall. Trans. B, 23 (1992) 13. U. Patnaik and J. Muralidhar, Talanta, 42 (1995) 553. R. Walker, Ultrason. Sonochem., 4 (1997) 39. N.A. Madigan, C.R.S. Hagan, H. Zhang and L.A. Coury, Ultrason. Sonochem., 3 (1996) S239. T. Tanaka, Q.W. Zhang and F. Saito, J. Chem. Eng. Jpn., 35 (2002) 173.

369

This page intentionally left blank

Chapter 13

Solid phase microextraction as a tool for trace element determination Zolta´n Mester and Ralph Sturgeon 13.1

INTRODUCTION

Extensive use of organic solvents in analytical laboratories is no longer tolerated because of the associated health risks and disposal concerns. As a result, many solvent-free extraction methods have been described or rediscovered in the last decade. These can be classiﬁed according to the nature of the extraction phase, as shown in Fig. 13.1, i.e., gas, membrane or sorbent [1]. Other low solvent consumption methods, such as single droplet extraction, are becoming more popular [2]. One solvent-free extraction approach is solid phase microextraction (SPME) which, as is evident from Fig. 13.1, is a sorbent extraction technique similar to solid phase extraction (SPE). With SPME, the sorbent material is attached to the surface of a ﬁber rather than packaged into a cartridge (tube) or used on the surface of a ﬂat disk. Whereas the SPE technique has been primarily designed for use with liquid matrices and exhaustive extraction, SPME can be used in liquid (aqueous) or gaseous matrices and primarily aims for partial or equilibrium extraction of the analyte. The principal approach of SPME is the use of a small volume of extracting phase, usually less than 1 ml. The extracting phase can be a high molecular weight polymeric “liquid” or a solid sorbent, typically a high surface area porous material. Figure 13.2 illustrates the structure of a commercially available SPME unit. A small diameter fused silica ﬁber, coated with the extraction phase, is mounted in a syringe-like device for protection and ease of handling. The needle serves to conveniently pierce septa during sample extraction and desorption operations. Using the syringe-like mechanism of the holder unit, the ﬁber can be extruded from the needle to expose the extraction phase to the sample (headspace or liquid). After the sampling period, the same mechanism can be used to withdraw the ﬁber inside the needle. During the extraction and desorption periods, the ﬁber is thus exposed by sitting outside of the needle; during transfer of the SPME unit to a desorption apparatus, the polymeric end of the ﬁber is inside the protective needle. Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

371

Z. Mester and R. Sturgeon

Fig. 13.1. Classiﬁcation of solvent-free sample preparation methods [supercritical ﬂuid extraction (SFE), SPE, SPME].

Fig. 13.2. Commercial SPME device: (A) SPME ﬁber holder; (B) SPME holder and ﬁber assembly—section view [32] (reprinted with permission of Supelco Bellefonte, PA 16823, USA).

372

Solid phase microextraction as a tool for trace element determination

13.2

GENERAL DESCRIPTION OF SOLID PHASE MICROEXTRACTION

There are several books [3] and reviews [1,4,5] on SPME, describing in detail the extraction theory and process. A short overview is only presented here. The transport of analytes from the matrix into the extraction medium begins as soon as the coated ﬁber has been placed in contact with the sample. In most cases, SPME is considered to terminate when the analyte concentration has reached distribution equilibrium between the sample matrix and the ﬁber coating. In practice, this means that once equilibrium is reached, the amount extracted is constant, within the limits of experimental error, and is independent of further increase of extraction time. Simplicity and convenience of operation make SPME a superior alternative to more established techniques in a number of applications. In some cases, the technique facilitates unique investigations. The most dramatic advantages of SPME exist at the extremes of sample volumes. Because the setup is small and convenient, coated ﬁbers can be used to extract analytes from very small samples. For example, SPME devices are used to probe for substances emitted by a single ﬂower bulb during its life span; the use of sub-micrometer diameter ﬁbers permits the investigation of single cells. Because SPME is an equilibrium technique and therefore does not extract target analytes exhaustively, its presence in a system should not result in signiﬁcant disturbance. In addition, the technique facilitates speciation in natural systems, as the presence of a minute ﬁber, which removes small amounts of analyte, is not likely to disturb chemical equilibria. It should be noted, however, that the fraction of analyte extracted increases as the ratio of coating volume to sample volume increases. Complete extraction can be achieved for small sample volumes when distribution constants are large. This observation can be used to advantage if exhaustive extraction is required. SPME also allows rapid sample extraction and transfer of analyte to the analytical instrument. These features result in additional advantage when investigating intermediates in the system. 13.2.1 Extraction modes Two basic types of extractions can be performed using SPME: direct extraction and headspace extraction. Figure 13.3 illustrates the differences between these modes. In the direct extraction mode (Fig. 13.3B), the coated ﬁber is inserted into the sample medium and the analytes are transported directly to the extraction phase. To facilitate rapid extraction, some level of agitation is required to enhance transport of the analytes from the bulk of the solution to the vicinity of the ﬁber. For gaseous samples, natural convection and diffusion in the medium is sufﬁcient to facilitate rapid equilibration. For aqueous matrices, more efﬁcient agitation techniques, such as fast sample ﬂow, rapid ﬁber or vial movement, stirring or sonication, are required. These actions are

373

Z. Mester and R. Sturgeon

Fig. 13.3. SPME operation modes: (A) headspace sampling; (B) direct liquid phase (immersion) sampling.

undertaken to reduce the effect caused by the “depletion zone” which occurs close to the ﬁber as a result of ﬂuid shielding and slow diffusion of analytes in liquid media. In the headspace mode (Fig. 13.3A), the analytes need to be transported through a layer of gas before they can reach the coating. This approach primarily serves to protect the ﬁber coating from damage by high molecular weight and other non-volatile concomitants present in the liquid sample matrix, such as humic materials or proteins. This headspace mode also allows modiﬁcation of the matrix, such as a change of the pH, without damaging the ﬁber. Amounts of analyte extracted into the coating from the same vial at equilibrium using direct and headspace sampling are identical as long as sample and gaseous headspace volumes are the same. This is a result of the fact that the equilibrium concentration is independent of ﬁber location in the sample/headspace system. If the above condition is not satisﬁed, a signiﬁcant sensitivity difference between the direct and headspace approaches exists only for very volatile analytes. The choice of sampling mode has a signiﬁcant impact on extraction kinetics. When the ﬁber coating is in the headspace, the analytes are removed from the headspace ﬁrst, followed by indirect extraction from the matrix. Therefore, volatile analytes are extracted faster than semi-volatile components as they are present at a higher concentration in the headspace, which contributes to faster mass transport through the headspace. Temperature has a signiﬁcant effect on the kinetics of the process by determining the vapor pressure of analytes. In fact, the equilibration times for volatile components are shorter in the headspace SPME mode than for direct extraction under similar agitation conditions. This outcome occurs as a result of two factors: a substantial portion of analyte is in the headspace prior to extraction and diffusion coefﬁcients in the gas phase are typically four orders of magnitude larger than in liquid media. Though the concentration of semivolatile components in the gas phase at room temperature is small, mass

374

Solid phase microextraction as a tool for trace element determination

transfer rates are substantially lower and result in longer extraction times for such species. The situation can be improved by the use of even more efﬁcient agitation techniques, such as sonication, further reducing extraction time. The other option is to increase the temperature; this decreases the amount extracted at equilibrium but it may be acceptable if target limits of detection can still be attained. 13.2.2 Coatings The efﬁciency of the extraction process is dependent on the analyte distribution constant between coating and sample matrix. This characteristic parameter describes properties of a coating and its selectivity toward the analyte versus other matrix components. Speciﬁc coatings can be developed for a range of applications. Coating volume determines method sensitivity as well, but thicker coatings result in longer extraction times because diffusion is slow within the polymer extraction phase. Therefore, it is important to use the appropriate coating for a given application. Coating selection and design can be based on chromatographic experience. For example, a very pronounced difference in selectivity toward target analytes and interferences can be achieved by using surfaces common to afﬁnity chromatography or molecular imprinted polymers. To date, several experimental coatings have been prepared and investigated for a range of applications. In addition to liquid polymeric coatings, such as polydimethylsiloxane (PDMS) for general applications, other more specialized materials have been developed.

13.3

SOLID PHASE MICROEXTRACTION: STEP-BY-STEP METHOD DEVELOPMENT

13.3.1 Extraction mode selection Selection of the extraction mode is based on a consideration of the sample matrix composition and volatility of the analyte. Generally, for inhomogeneous matrices or pH or polarity incompatible matrices, the only choice is headspace sampling. For example, with typically used inorganic analytical methods, acid leaching is not compatible with the ﬁbers because their working range is limited in pH and therefore direct extraction cannot be performed. If the aqueous matrix contains large non-polar species, such as lipid components, the non-polar ﬁbers should not be exposed to them because they rapidly saturate the extraction phase. Obviously, for clean and compatible matrices, either headspace or direct sampling can be used.

375

Z. Mester and R. Sturgeon

13.3.2 Fiber coating selection The chemical nature of the analyte determines the type of polymer used for the extraction. Selection of coating is based primarily on the polarity and volatility of the target compound. As a ﬁrst consideration, the simple rule “like dissolve like” applies well for liquid polymeric coatings. PDMS is the most useful liquid type coating. It is very rugged and its extraction characteristics can be easily estimated from the wide gas chromatographic experience gained with this coating. However, for special cases, other solid adsorptive coatings can be considered. The distribution constant of the analyte and the coating thickness inﬂuence the sensitivity of the method. Thicker coatings require longer equilibration times and thus the coating of choice is the thinnest one that will provide sufﬁcient sensitivity for the determination. 13.3.3 Derivatization method selection Analyte derivatization may prove advantageous or necessary for several reasons. For example, if the target analyte is not suitable for gas chromatographic analysis or for the polarity-based extraction offered by the SPME coating (such as ionic compounds), derivatization before, during or after the SPME procedure can be used to enhance detection power by introducing select functional groups into the analyte [such as halogen groups for electron capture detection (ECD) or negative chemical ionization mass spectrometry]. Derivatization is especially important for speciation analysis, wherein most of the target compounds are in an ionic form. 13.3.4 Optimization of desorption conditions Optimal advantage of SPME methodology is achieved when the smallest possible desorption volume can be realized for use with either liquid or gas chromatographic sample introduction. The usual spilt/splitless types of GC injectors were designed for large volume sample introduction and are therefore normally equipped with a large volume glass insert sufﬁcient to accommodate the vapor volume arising from a few microliters of organic solvent. By introducing the analyte absorbed on or in a sorbent bed, there is no need for such a large desorption volume because there is no solvent expansion effect. Narrow bore injection liners can thus be used to enhance sensitivity by minimizing dispersion. 13.3.5 Sample volume optimization The sensitivity achieved with SPME methodology is dependent on the number of moles of analyte extracted from the sample. Provided the sample volume q the coating volume ðVc Þ; the amount of analyte extracted is

376

Solid phase microextraction as a tool for trace element determination

independent of the volume of sample if distribution equilibrium is achieved. If the available sample volume is not signiﬁcantly greater than the coating volume, conventional care is needed for the sample volume measurement. Care must also be taken to avoid analyte losses via evaporation, adsorption or microbiological activity. 13.3.6 Optimization of the extraction time An optimal approach to SPME analysis is to allow the analyte to reach equilibrium between the sample and the ﬁber coating. The equilibration time is deﬁned as the time after which the amount of analyte extracted remains constant and corresponds, within the limits of experimental error, to the amount extracted after inﬁnite time. Care should be taken when determining the equilibration time because, in some cases, a substantial reduction of the slope of the curve might be mistakenly interpreted as the point at which equilibrium has been achieved. Such phenomena often occur in headspace SPME analysis of aqueous samples, where a rapid rise of the equilibration curve, corresponding to extraction from the gas phase only, is followed by a very slow increase related to analyte transfer from the aqueous phase through the headspace to the ﬁber. Determination of the amount extracted at equilibrium allows calculation of the distribution constants. When equilibration times are excessively long, shorter extraction times can be used. However, in such cases, the extraction time and mass transfer conditions have to be strictly controlled to assure good precision. At equilibrium, small variations in the extraction time do not affect the amount of analyte extracted by the ﬁber. On the other hand, over the steep part of the curve, even small variations in the extraction time may result in signiﬁcant variations in the amount extracted. The relative error becomes larger when the extraction time is shorter. Autosamplers can reproduce the time very precisely, and the precision of analyte determination can then be very good, even when equilibrium is not reached in the system. However, this requires that the mass transfer conditions and the system temperature remain constant for all experiments. 13.3.7 Optimization of extraction conditions An increase in extraction temperature causes an increase in the extraction rate, but simultaneously a decrease in the distribution constant. In general, if the extraction rate is of major concern, the highest temperature which still provides satisfactory sensitivity should be used. Adjustment of the pH of the sample can improve the sensitivity of the method for basic and acidic analytes. This is related to the fact that unless ion exchange coatings are used, SPME can extract only neutral (non-ionic) species from water. By properly adjusting the pH, weak acids and bases can be converted to their neutral forms for extraction by the SPME ﬁber. To ensure

377

Z. Mester and R. Sturgeon

that at least 99% of the acidic compound is in the neutral form, the pH should be at least two units lower than the pKa of the analyte. For basic analytes, the pH must be larger than pKb by two units.

13.3.8 Determination of the linear dynamic range Modiﬁcation of the extraction conditions affects both the sensitivity and the equilibration time. It is advisable, therefore, to check the previously determined extraction time before proceeding to the determination of the linear dynamic range. This step is required if substantial changes of the sensitivity occur during the optimization process. SPME coatings include polymeric liquids, such as PDMS, which by deﬁnition exhibit capability for a very broad linear range. For solid sorbents, such as Carbowax/DVB or PDMS/DVB, the linear range is narrower because of a limited number of sorption sites on the surface, but it can still span several orders of magnitude for typical analytes in pure matrices. In some rare cases, when the analyte has extremely high afﬁnity towards the surface, saturation can occur at low analyte concentrations. In such cases, the linear range can be expanded by decreasing the extraction time. 13.3.9 Selection of the calibration method Standard calibration procedures can be used with SPME. A ﬁber blank should be checked ﬁrst to ensure that neither the ﬁber nor the instrument causes interferences with the determination. The ﬁber should be conditioned prior to ﬁrst use by desorption in a GC injector or other similarly designed conditioning device. This process ensures that the ﬁber coating itself does not introduce contaminants or interfering species. Fiber conditioning may have to be repeated after analysis of samples containing signiﬁcant amounts of high molecular weight compounds because such compounds may require longer desorption times (or higher desorption temperatures) than the analytes of interest. A special calibration procedure, such as isotope dilution (ID) or standard additions, should be used for more complex samples. With these methods, it is assumed that the target analytes behave similarly to enriched spikes added prior to the extraction. This is usually a valid assumption when analyzing homogeneous samples. However, it might not be true when heterogeneous samples are analyzed, unless the native analytes are completely released from the matrix under the conditions used or care is taken to ensure the added spikes are fully equilibrated with all components of the sample matrix. Moreover, whenever any of these methods are used, an inherent assumption is made that the response is linear in the concentration range between the original analyte concentration and the spiked concentration. While this is

378

Solid phase microextraction as a tool for trace element determination

usually true for ﬁbers extracting the analytes by absorption (PDMS, PA), and detectors with wide linear range are available, problems may arise when porous polymer ﬁbers (PDMS/DVB, Carbowax/DVB) are used, or when the detector applied has a narrow linear range. It is, therefore, important to verify the linearity of the response using standard solutions, before applying standard additions or ID for calibration. To improve the accuracy and precision, multipoint standard additions should be used whenever it is practical. 13.3.10 Precision of the method The most important factors affecting precision in SPME are presented below: – – – – – – – – – – – – – – – –

agitation conditions sampling time (if non-equilibrium conditions are used) temperature sample volume headspace volume vial shape condition of the ﬁber coating (cracks, adsorption of high MW species) geometry of the ﬁber (thickness and length of the coating) sample matrix components (salt, organic material, humidity, etc.) time between extraction and analysis analyte losses (adsorption on the walls, permeation through Teﬂon, absorption by septum) geometry of the injector ﬁber positioning during injection condition of the injector (integrity of septum) stability of the detector response moisture in the needle

To ensure good reproducibility of the SPME measurement, all experimental parameters listed above should be kept constant. 13.3.11 Automation of the method While SPME is a very powerful investigative tool, it can also be a technique of choice in many applications for processing large numbers of samples. To accomplish this task automation of the methodology is required. As automated SPME devices with more advanced features and capabilities become available, automation becomes easier. The currently available SPME autosampler from Varian enables direct sampling with agitation of the sample by ﬁber vibration and static headspace sampling. In some cases, custom modiﬁcations to the commercially available systems can facilitate operation of the method closer to optimum conditions.

379

Z. Mester and R. Sturgeon

13.4

SOLID PHASE MICROEXTRACTION FOR SPECIATION ANALYSIS

13.4.1 Volatile metal species—gas chromatographic determination Mester et al. [5] recently reviewed the determination of organometallic species using SPME. 13.4.1.1 Ethylation Most applications of SPME to speciation are based on classical gas chromatographic (GC) determinations. Table 13.1 summarizes the SPME –GC methods used for such speciation studies. Methods have been described for mercury, tin, lead, arsenic and selenium. Both headspace and direct extraction methods have been used for the sampling of organometallic compounds and, generally, some type of derivatization process is necessary to permit their GC separation. The typical derivatization method used for tin, lead and mercury species is ethylation using sodium tetraethylborate reagent (NaBEt4). As is evident from Table 13.1, the working pH range for the derivatization is 4.0 –5.3, depending on the target compounds. An advantage of NaBEt4 is that derivatization can be accomplished in an aqueous environment, the natural medium for most environmental and biological samples. There is no need to change phases, as in the case of Grignard reagents. Derivatization with NaBEt4 offers the unique possibility for multielemental speciation of tin, mercury and lead species. Moens et al. [6] reported a comparison of sensitivity between “conventional” liquid/liquid- and headspace SPME-extraction of butyltin and organolead compounds. They found that headspace SPME provided about 300-fold better sensitivity for butyltin compounds and about 35-fold enhancement for trimethyllead. Analysis of butyltin species based on SPME can thus be accomplished using a conventional ﬂame ionization detector (FID). One of the main limitations of this technique is that it cannot be applied to the separate identiﬁcation of species containing the ethyl ligand. For example, reaction of triethyllead and inorganic lead with NaBEt4 produces the same compound: tetraethyllead. Yu and Pawliszyn [7] overcame this limitation by utilizing deuterated NaBEt 4 for the derivatization of organolead compounds. A typical sampling procedure involves the following: for solid samples and for most biological tissues a “soft” digestion is used to drive the target compounds into the liquid phase. For homogeneous liquid samples (such as seawater), no digestion step is necessary. The aqueous sample is typically transferred to a 40 mL septum-sealed glass vial, pH adjusted to a predetermined value, the derivatization agent added and the vial tightly sealed. To facilitate mass transfer in the system, the solution is often continuously stirred by use of a magnetic stirrer. Aguerre et al. [8] examined the use of mechanical shaking for liquid phase extraction of organotin species. The SPME ﬁber is subsequently exposed to the headspace or the liquid phase of

380

TABLE 13.1 SPME methods for mercury, tin, lead, arsenic and selenium speciation with GC separation Derivatization

Fiber/ extraction time/ extraction mode

Chromatographic column/ temperature program

Detector

Detection limita

Reference

Water, ﬁsh tissue

NaBEt4/ acetate buffer pH 4.5

100 mm PDMS/ 5 min/HS

AFS

3.0 ng/l

[33]

(Me)2Hg

Gas condensate

100 mm PDMS/ 30 s/HS

MIP-AES

20 mg/l

[16]

Me2Hg Et2Hg

Soil

No derivatization (direct sampling) No derivatization (direct sampling)

15 m £ 0.53 mm £ 1.5 mm, DB1 column, 408C (30 s)—308C/min to 858C (1 min) 208C/min to 2008C (1 min) 15 m £ 0.53 mm £ 1.5 mm DB1, isotherm 308C

MIP-AES

3.5 mg/l

[34]

MeHgþ

Biological samples, sediments

Hydride generation (KBH4)/ acetate buffer pH 3

Fused-silica ﬁber (pretreated with cc. HF acid for 3.5– 4 h)/ 1.5– 2 h/HS

25 m £ 0.32 mm £ 0.25 mm HP-1, 408C (5 min)— 408C/min to 2008C (1 min) 10 m £ 0.25 mm with CPL-SIL 5CB coating, isotherm 408C

AAS (quartz tube)

Not reported

[9]

Hg MeHgþ

100 mm PDMS/ 20 min/HS

381

continued

Solid phase microextraction as a tool for trace element determination

Sample type

Species

382

TABLE 13.1 (continuation) Sample type

Derivatization

Fiber/ extraction time/ extraction mode

Chromatographic column/ temperature program

Detector

Detection limita

Reference

MeHg EtHg PhHg

Soil

Hydride generation (KBH4) acetate buffer pH 4

Fused-silica ﬁber (pretreated with cc. HF acid for 3.5– 4 h)/ 1.5 –2 h/HS

AAS (quartz tube)

Not reported

[10]

Hg2þ MeHgþ

Urine

NaBEt4/ buffer pH 4

100 mm PDMS/ 15 min/HS

30 m £ 0.32 mm £ 0.25 mm SPB-1, 508C (1 min)— 408C/min to 658C (1 min), 1508C (1 min), 2008C (1 min) 30 m £ 0.25 mm £ 0.25 mm HP-5, 508C (3 min)— 128C/min to 2808C

EI-MS

93 ng/l 303 ng/l

[35]

Sn TeMT TMT DMT MMT

Water, seawater

NaBEt4/ acetic acid buffer pH 4

100 mm PDMS/ 20 min/HS

FPD

41 ng/l 15 ng/l 8.4 ng/l 8.6 ng/l

[36]

River water

NaBEt4/ ethanolic acid buffer pH 4.8

100 mm PDMS/ 60 min/LPh

30 m £ 0.32 mm £ 1.8 mm DB-624, 558C (2 min)—108C/min to 1508C (2 min) 30 m £ 0.25 mm £ 0.25 mm (methylsiloxane), 708C (1 min)— 158C/min to 2708C (6 min)

FPD

2 ng/l 2 ng/l 4 ng/l 1 ng/l 2 ng/l 3 ng/l

[37]

MBT DBT TBT MPhT DPhT TPhT

Z. Mester and R. Sturgeon

Species

TABLE 13.1 (continuation) Sample type

Derivatization

Fiber/ extraction time/ extraction mode

Chromatographic column/ temperature program

Detector

Detection limita

Reference

MBT DBT TBT

Environmental, sediment

NaBEt4/ acetate buffer pH 4.0

100 mm PDMS/ 60 min/ HS

FID

1.0 mg/l 1.2 mg/l 0.9 mg/l

[38]

MBT DBT TBT MPhT DPhT TPhT MBT DBT TBT TeBT

Sediment, sewage sludge

NaBEt4/ ethanolic acid buffer pH 4.8

100 mm PDMS/ 60 min/ LPh

FPD

NaBEt4/ acidiﬁed with HCl

10 mm PDMS/ 45 min/ LPh

MIP-OES

0.031 ng/l 0.007 ng/l 0.006 ng/l 0.114 ng/l 0.167 ng/l 0.583 ng/l mg/l range

[8]

Slurry of sediment

30 m £ 0.25 mm £ 1.0 mm SPB-1, 408C (1 min)— 208C/min to 1408C (1 min)—208C/min to 2208C (1 min) 30 m £ 0.25 mm £ 0.25 mm (methylsiloxane), 708C (1 min)— 158C/min to 2708C (6 min) 25 m £ 0.32 mm £ 0.17 mm HP-1, 808C— 208C/min to 2308C (0.75 min)

Spiked water

NaBEt4/ acetate buffer pH 4.0

100 mm PDMS/ 15–20 min/ HS

EI-MS

0.2 mg/l

[15]

Pb Pb2þ

383

30 m £ 0.25 mm £ 0.25 mm Omegawax 408C (1 min)—208C/min to 1208C (1 min)

[39]

continued

Solid phase microextraction as a tool for trace element determination

Species

384

TABLE 13.1 (continuation) Sample type

Derivatization

Fiber/ extraction time/ extraction mode

Chromatographic column/ temperature program

Detector

Detection limita

Reference

Pb2þ

Blood, urine

NaBEt4/ acetate buffer pH 4.0

100 mm PDMS/ 15 min/ HS

EI-MS

2 –3 mg/l

[40]

Pb2þ TML TEL TeEL

–

DeuteratedNaBEt4/ acetate buffer pH 4.0

100 mm PDMS/ 10 min/HS

30 m £ 0.25 mm £ 0.25 mm SPB 5 408C (1 min)— 208C/min to 1208C (4 min) 30 m £ 0.25 mm £ 0.25 mm SPB 5, 708C isotherm

EI-MS

95 ng/l 130 ng/l 83 ng/l 90 ng/l

[7]

River and tap water

Piaselenol formation

Not reported/ 30 min/HS

30 m £ 0.25 mm £ 0.25 mm 808C (2 min)— 108C/min to 2808C

EI-MS

6 ng/l

[41]

Urine

Thioglycolmethylate derivatization

100 mm PDMS/ 40 min/LPh

15 m £ 0.25 mm £ 0.25 mm SPB 5, 1108C (1 min)—208C/min to 2308C

EI-MS

0.29 mg/l 0.12 mg/l

[13]

Se Se(IV) total Se

As DMA MMA

Z. Mester and R. Sturgeon

Species

TABLE 13.1 (continuation) Species

Derivatization

Fiber/ extraction time/ extraction mode

Chromatographic column/ temperature program

Detector

Detection limita

Reference

Sediment

NaBEt4/ acetate buffer pH 5.3

100 mm PDMS/ 10 min/ HS

ICP-MS

NaBEt4/ acetate buffer pH 5.3

100 mm PDMS/ 10 min/ HS

Surface water, sediment

NaBEt4/ acetate buffer pH 5.0

100 mm PDMS/ 10 min/ HS

30 m £ 0.25 mm £ 0.50 mm (PDMS) 608C (1 min)— 208C/min to 2008C (0.5 min)

ICP-MS

0.34 ng/l 2.1 ng/l 1.1 ng/l 4.3 ng/l 0.19 ng/l 9 ng/l 13 ng/l 9 ng/l 22 ng/l 18 ng/l 7 ng/l 3.7 ng/l 0.38– 1.2 ng/l 0.13– 0.15 ng/l

[6]

Body ﬂuids

30 m £ 0.25 mm £ 0.50 mm (PDMS) 608C (1 min)— 208C/min to 2008C (0.5 min) 30 m £ 0.25 mm £ 0.50 mm VA-5, 408C (3 min)— 108C/min to 2508C

EI-MS-MS

[42]

[43]

TML, trimethyllead; TEL, triethyllead; TeEL, tetraethyllead; DMA, dimethylarsinic acid; MMA, monomethylarsonic acid; MBT, monobutyltin; DBT, dibutyltin; TBT, tributyltin; MPhT, monophenyltin; DPhT, diphenyltin; TPhT, triphenyltin; NaBEt4, sodium tetraethylborate; LPh, liquid phase; MeHg, methylmercury; EtHg, ethylmercury; PhHg, phenylmercury.

385

HS, headspace; AAS, atomic absorption spectrometry; AFS, atomic ﬂuorescence spectrometry; EI-MS, electron impact ionization mass spectrometry; FID, ﬂame ionization detection; FPD, ﬂame photometry detection; ICP-MS, inductively coupled plasma mass spectrometry; MIP-OES, microwave induced plasma atomic emission spectrometry. a

Detection limit normally based on the concentration of solution sampled by SPME.

Solid phase microextraction as a tool for trace element determination

Multielement MBT DBT TBT MeHg TML MBT DBT TBT MeHg, Hg2þ TML Alkylmercury, alkyltin, alkyllead

Sample type

Z. Mester and R. Sturgeon

the sample for a predetermined time. After extraction, the ﬁber is transferred to a suitable desorption unit. 13.4.1.2 Hydride generation Because of the high volatility of metal hydrides, few researchers have reported results on the headspace sampling/determination of metal hydrides using SPME techniques. Jiang and coworkers [9,10] developed a sampling method for organomercury species based on hydride generation using KBH4 reagent. Mester et al. [11] tested compatibility of two different ﬁbers for sampling volatile metal hydrides coupled with inductively coupled plasma mass spectrometry (ICP-MS) detection. An adsorption-based carboxen coating provided better sensitivity than an absorption-based extraction with a liquid type polymeric coating (PDMS). The success of absorptive sampling conﬁrms the relatively high stability of the studied metal hydrides (As, Se, Sn and Sb) because they survive diffusion into the polymeric liquid. 13.4.1.3 Other derivatization methods Guidotti [12] described the determination of Se(IV) by headspace SPME –GC/ MS following derivatization with piaselenol formation. The piaselenol complex is extracted from the liquid phase by immersing a 100 mm PDMS-coated ﬁber into the sample solution. The method can also be applied to Se(IV) and Se(VI) speciation by determining the Se(IV) content ﬁrst, followed by the total selenium concentration after converting all selenium species into the Se(IV) oxidation state. Mester and Pawliszyn [13] reported a method for the speciation of dimethylarsinic acid (DMA) and monomethylarsonic acid (MMA) by SPME – GC/MS. Thioglycolmethylate (TGM) was used for derivatization. The method is based on the known “afﬁnity” of the arsenic compound for the thiol group [14]. 13.4.1.4 Direct determination (no derivatization) Organometallic species, which are normally saturated (non-ionic) and sufﬁciently volatile, can be sampled by SPME and determined by GC without derivatization. Gorecki and Pawliszyn [15] described a simple sampling procedure for tetraethyllead, wherein a PDMS ﬁber was exposed to the headspace above an aqueous sample. GC –MS was used for quantitation. Similar studies were performed by Snell et al. [16] for determination of dimethylmercury in the headspace above natural gas condensates. Mester et al. [17] described an SPME method for methylmercury determination. Headspace SPME sampling was performed above a methylmercury solution, which was previously saturated with sodium chloride. This method was based on the relatively high vapor pressure of methylmercury chloride. A slightly polar solid coating (PDMS/DVB) was used for extraction.

386

Solid phase microextraction as a tool for trace element determination

Sample introduction into an ICP-MS was achieved with a unique thermal desorption interface consisting of a heated glass-lined splitless type GC injector placed directly at the base of the torch to minimize the length of needed transfer-line. This arrangement provided for fast desorption and high sample introduction efﬁciency. Direct liquid immersion and headspace extraction of methylmercury were studied. For clean solutions, immersion sampling SPME provided good sensitivity that was linear over two orders of magnitude whereas headspace sampling showed 15% lower sensitivity but a linear range of more than three orders of magnitude. Calibration by the method of additions using direct extraction revealed a severe matrix effect with biological tissue samples, diminishing the methylmercury response 70-fold; that obtained by headspace extraction was statistically indistinguishable from signals generated using matrix-free standards. The real novelty of the method was that speciation could be achieved without employing any separation method (GC or HPLC) and was based entirely on the selectivity of the extraction procedure and detection technique. A similar approach has been used for tributyltin determination from aqueous samples [18,19] or after heating directly from solid samples [20]. As a secondary result of these studies, a new sample introduction approach was developed which can be used with any type of atomic spectroscopic instrumentation. In addition to SPME, small volumes (1 –3 m L) of organic solvents may be introduced into the ICP with this device. A schematic diagram of the thermal desorption ICP-MS interface is shown in Fig. 13.4.

Fig. 13.4. Schematic illustration of the thermal desorption interface for SPME analyte introduction into ICP-MS.

387

Z. Mester and R. Sturgeon

13.5

SOLID PHASE MICROEXTRACTION AS AN INVESTIGATIVE TOOL

SPME can and has been used as a research tool. Because of the non-invasive nature of the sampling process, it is suitable for sampling in sensitive or difﬁcult environments and is also fast and easy to use with volatile and semivolatile compounds. For example, Barshick and coworkers [21–23] published several papers focusing on the determination of mercury species in soils using various detection methods. The applied sampling and sample introduction method were SPME based. Gorecki et al. [24] applied SPME for sample preparation and analyte introduction into a radio frequency glow discharge mass spectrometer. The distinctive fragmentation patterns obtained for tetramethyltin and tetraethyllead species offer the possibility of direct speciation without separation. Rosenkranz et al. [25] used SPME sampling and sample introduction for the study of transalkylation processes in different mercury species. Mester and Pawliszyn [26] employed in-tube SPME for sample introduction into an electrospray mass spectrometer. The main purpose of the study was the elucidation of the collision induced fragmentation patterns of different organolead species. 13.6

LIMITATIONS OF SOLID PHASE MICROEXTRACTION

The principal limitations of SPME technology are strongly linked to the main advantages of the technology. The volume of the polymer extraction phase is very small and, although responsible for the non-exhaustive nature of the extraction, requires extreme precision during manufacture of the coating so as to reproducibly produce the same coating quality. The reuse character of the SPME ﬁber is also a signiﬁcant analysis cost related advantage but, at the same time, some level of degradation of the ﬁber occurs during repeated usage, resulting in compromised accuracy and precision. Haberhauer Troyer et al. [27] recently presented a study of the surface characteristics of SPME coatings. Using electron microscopy, the surfaces of new and used PDMS and PDMS/ Carboxen ﬁbers were examined. Damage to the coatings was detected on the tip of the ﬁber and at the bond between the polymer-coated fused silica ﬁber and the ﬁber attachment tubing. The pore size ranges up to 40 mm on the surface of porous PDMS/Carboxen ﬁbers and the pore density is highly variable between ﬁbers. On the surface of PDMS ﬁbers, physical contamination was detected; metal particles were present, which likely came from the septum-piercing needle (the composition of the particles was determined and found to closely match the needle composition). Metallic contamination on the ﬁbers can pose a serious threat to the ﬁeld of trace metal analysis. Mester et al. [28] also reported data relating to the intra- and interﬁber reproducibility achieved during methylmercury analysis. The results for

388

Solid phase microextraction as a tool for trace element determination

intra- and inter-ﬁber performance obtained with ﬁve replicate headspace samplings of a 10 ng/ml sample solution of methylmercury using a PDMS/DVB ﬁber were reported. The three ﬁbers had not been used prior to this process; they were used as provided by the supplier. For any given ﬁber, the (intra-) reproducibility of measurement is quite good, averaging about 2.3% RSD. However, the RSD of the means for the three ﬁbers is greater than 20%. More alarming is the difference between the two extremes, which is more than 30%. These data are likely a reﬂection of the quality of manufacture of the ﬁbers. Surprisingly, over the past several years of commercial history of SPME and the hundreds of research papers (organic analysis) utilizing this technology, only few discussions have arisen concerning the quality of the results with respect to ﬁber-to-ﬁber performance. The unique aspect of manufacture of SPME ﬁbers is the extremely low volume of the extraction phase. Any irregularity or inhomogeneity in the polymer phase/surface may result in a signiﬁcant effect on its extraction characteristics. This effect could be more pronounced in the case of non-equilibrium extraction with solid coatings where the extraction is based on adsorption phenomena rather than absorption. The imprecision of the results obtained with different ﬁbers often makes it necessary to employ an internal standard or standard addition method which, unfortunately, can signiﬁcantly increase the analysis time. 13.7

ISOTOPE DILUTION CALIBRATION IN COMBINATION WITH SOLID PHASE MICROEXTRACTION

To improve the precision of the quantiﬁcation by SPME, ID techniques can be applied. Isotope dilution mass spectrometry (IDMS) has been widely employed for trace element analysis in a variety of sample matrixes and recently has been considered to be a primary (ratio) method of analysis as a consequence of its high accuracy and precision. IDMS addresses means for the correction for species conversion that might occur during sampling, manipulation and even analysis procedures. Unfortunately, its application to species-speciﬁc determinations has been limited by the non-availability of commercial enriched standards. If these are available, a known amount of spike of a known isotopic composition is added to a known amount of sample of known isotopic composition; the isotopic composition of the spike and the sample must be different. A number of advantages of ID accrue, including enhanced precision and accuracy; matrix effects are accounted for, as quantitation is done by ratio measurements; non-quantitative analyte recovery does not impact on the ﬁnal results; species alteration during sample workup can be assessed; and an alternative and comparative quantitation strategy is provided. Only recently has this approach been applied to SPME determination of organometallic compounds, such as butyltins [29,30] and methylmercury [31]. In most of these studies, ICP-MS and GC/MS are employed as detection systems.

389

Z. Mester and R. Sturgeon

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32

390

J. Pawliszyn, Trac Trends Anal. Chem., 14 (1995) 113. E. Psillakis and N. Kalogerakis, Trac Trends Anal. Chem., 21 (2002) 53. J. Pawliszyn, Solid Phase Microextraction: Theory and Practice. Wiley, New York, NY, 1997. J. Pawliszyn, J. Chromatogr. Sci., 38 (2000) 270. Z. Mester, R. Sturgeon and J. Pawliszyn, Spectrochim. Acta Part B At. Spectrosc., 56 (2001) 233. L. Moens, T. DeSmaele, R. Dams, P. VandenBroeck and P. Sandra, Anal. Chem., 69 (1997) 1604. X.M. Yu and J. Pawliszyn, Anal. Chem., 72 (2000) 1788. S. Aguerre, C. Bancon Montigny, G. Lespes and M. Potin Gautier, Analyst, 125 (2000) 263. B. He, G.B. Jiang and Z.M. Ni, J. Anal. At. Spectrom., 13 (1998) 1141. B. He and G.B. Jiang, Fresenius J. Anal. Chem., 365 (1999) 615. Z. Mester, R.E. Sturgeon and J.W. Lam, J. Anal. At. Spectrom., 15 (2000) 1461. M. Guidotti, J. AOAC Int., 83 (2000) 1082. Z. Mester and J. Pawliszyn, J. Chromatogr. A, 873 (2000) 129. Z. Mester, G. Horvath, G. Vitanyi, L. Lelik and P. Fodor, Rapid Commun. Mass Spectrom., 13 (1999) 350. T. Gorecki and J. Pawliszyn, Anal. Chem., 68 (1996) 3008. J.P. Snell, W. Frech and Y. Thomassen, Analyst, 121 (1996) 1055. Z.N. Mester, J. Lam, R. Sturgeon and J. Pawliszyn, J. Anal. At. Spectrom., 15 (2000) 837. Z. Mester and R.E. Sturgeon, Environ. Sci. Technol., 36 (2002) 1198. C. Bancon Montigny, P. Maxwell, L. Yang, Z. Mester, R.E. Sturgeon and J.W. Lam, J. Anal. At. Spectrom., 17 (2002) 1506. Z. Mester, J. Anal. At. Spectrom., 17 (2002) 868. C.M. Barshick, S.A. Barshick, M.L. Mohill, P.F. Britt and D.H. Smith, Rapid Commun. Mass Spectrom., 10 (1996) 341. C.M. Barshick, S.A. Barshick, P.F. Britt, D.A. Lake, M.A. Vance and E.B. Walsh, Int. J. Mass Spectrom., 178 (1998) 31. C.M. Barshick, S.A. Barshick, E.B. Walsh, M.A. Vance and P.F. Britt, Anal. Chem., 71 (1999) 483. T. Gorecki, M. Belkin, J. Caruso and J. Pawliszyn, Anal. Commun., 34 (1997) 275. B. Rosenkranz, J. Bettmer, W. Buscher, C. Breer and K. Cammann, Appl. Organomet. Chem., 11 (1997) 721. Z. Mester and J. Pawliszyn, Rapid Commun. Mass Spectrom., 13 (1999) 1999. C. Haberhauer Troyer, M. Crnoja, E. Rosenberg and M. Grasserbauer, Fresenius, J. Anal. Chem., 366 (2000) 329. Z. Mester, J. Lam, R. Sturgeon and J. Pawliszyn, J. Anal. At. Spectrom., 15 (2000) 837. L. Yang, Z. Mester and R.E. Sturgeon, J. Anal. At. Spectrom., 17 (2002) 944. C. Bancon Montigny, P. Maxwell, L. Yang, Z. Mester and R.E. Sturgeon, Anal. Chem., 74 (2002) 5606. L. Yang, Z. Mester and R.E. Sturgeon, J. Anal. At. Spectrom., 18 (2003) 431–436. Supelco Data Sheet No. T713019A, Instructions for the Supelco solid phase microextraction ﬁber holder for manual use.

Solid phase microextraction as a tool for trace element determination 33 34 35 36 37 38 39 40 41 42 43

Y. Cai, S. Monsalud, K.G. Furton, R. Jaffe and R.D. Jones, Appl. Organomet. Chem., 12 (1998) 565. S. Mothes and R. Wennrich, HRC J. High Resolut. Chromatogr., 22 (1999) 181. M. Guidotti and M. Vitali, HRC J. High Resolut. Chromatogr., 21 (1998) 665. Y. Morcillo, Y. Cai and J.M. Bayona, HRC J. High Resolut. Chromatogr., 18 (1995) 767. G. Lespes, V. Desauziers, C. Montigny and M. Potin Gautier, J. Chromatogr. A, 826 (1998) 67. E. Millan and J. Pawliszyn, J. Chromatogr. A, 873 (2000) 63. S. Tutschku, S. Mothes and R. Wennrich, Fresenius J. Anal. Chem., 354 (1996) 587. X.M. Yu, H.D. Yuan, T. Gorecki and J. Pawliszyn, Anal. Chem., 71 (1999) 2998. M. Guidotti, G. Ravaioli and M. Vitali, HRC J. High Resolut. Chromatogr., 22 (1999) 414. L. Dunemann, H. Hajimiragha and J. Begerow, Fresenius J. Anal. Chem., 363 (1999) 466. T. De Smaele, L. Moens, P. Sandra and R. Dams, Mikrochim. Acta, 130 (1999) 241.

391

This page intentionally left blank

Chapter 14

Solid-phase extraction Vale´rie Camel

14.1

INTRODUCTION

Despite the selectivity and sensitivity of analytical techniques such as atomic absorption spectrometry, there is a crucial need for the preconcentration of trace elements before their analysis due to their frequent low concentrations in many samples (especially natural waters). Additionally, because high levels of non-toxic components usually accompany analytes, a clean-up step is often required. Liquid –liquid extraction (LLE) is a classical method for preconcentrating metal ions and/or matrix removal. Solid-phase extraction (SPE) is another approach that offers a number of important beneﬁts. It reduces solvent usage and exposure, disposal costs, and extraction time for sample preparation. Consequently, in recent years, SPE has been used successfully for the separation and sensitive determination of metal ions, mainly in water samples. After outlining the theory of this technique, guidelines are given for the development of SPE-based methods for preconcentration of many trace elements. Finally, examples of applications are presented. 14.2

THEORY

The principle of SPE is similar to that of LLE, involving a partitioning of solutes between two phases. However, instead of two immiscible liquid phases, as in LLE, SPE involves partitioning between a liquid (sample matrix) and a solid (sorbent) phase. It is also very similar to liquid –solid extraction (LSE), which involves contacting the sample with a suitable adsorbent material (solid phase) while agitating the system for a given time. However, while LSE is adequate for samples containing cations in the ppm range, preconcentration afforded by SPE is useful when trace levels are to be analysed. Indeed, SPE is a sample treatment technique that enables the concentration and puriﬁcation of analytes from solution by sorption onto a solid sorbent. The basic approach involves passing the liquid sample through a column, a cartridge, a tube or a disc containing an adsorbent that retains the analytes. After all of the sample Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

393

V. Camel

has been passed through the sorbent, retained analytes are subsequently recovered upon elution with an appropriate solvent. The ﬁrst experimental applications of SPE started 50 years ago [1,2]. However, its growing development as an alternative approach to LLE for sample preparation started only in the mid-1970s. It has been extensively used in the past 15 years for the preconcentration of organic micropollutants, especially pesticides, in water samples [3]. However, numerous studies have also shown the great potential of this technique for speciation studies.

14.2.1 Presentation of the technique 14.2.1.1 Basic principles An SPE method always consists of three to four successive steps, as illustrated in Fig. 14.1. First, the solid sorbent should be conditioned using an appropriate solvent, followed by the same solvent as the sample solvent. This step is critical, as it enables the wetting of the packing material and the solvation of the functional groups. In addition, it removes possible impurities initially contained in the sorbent or the packaging. Also, this step removes the air present in the column and ﬁlls the void volume with solvent. The nature of the conditioning solvent depends on the nature of the solid sorbent. Typically, for reversed-phase sorbent (such as octadecyl-bonded silica), methanol is frequently used, followed by water or aqueous buffer whose pH and ionic strength are similar to that of the sample. Care must be taken not to allow the solid sorbent to dry between the conditioning and the sample treatment steps, otherwise the analytes will not be efﬁciently retained and poor recoveries will be obtained. If the sorbent dries for more than several minutes, it must be reconditioned.

Fig. 14.1. SPE operation steps.

394

Solid-phase extraction

The second step is the percolation of the sample through the solid sorbent. Depending on the system used, volumes can range from 1 ml to 1 l. The sample may be applied to the column by gravity, pumping, aspirated by vacuum, or delivered by an automated system. The sample ﬂow-rate through the sorbent should be low enough to enable efﬁcient retention of the analytes, and high enough to avoid excessive duration. During this step, the analytes are concentrated on the sorbent. Even though matrix components may also be retained by the solid sorbent, some of them pass through, thus enabling some puriﬁcation (matrix separation) of the sample. The third step (which is optional) may be the washing of the solid sorbent with an appropriate solvent having a low elution strength, to eliminate matrix components that have been retained by the solid sorbent, without displacing the analytes. A drying step may also be advisable, especially for aqueous matrices, to remove traces of water from the solid sorbent. This will eliminate the presence of water in the ﬁnal extract, which, in some cases, may hinder the subsequent concentration of the extract and/or the analysis. The ﬁnal step consists of the elution of the analytes of interest by an appropriate solvent, without removing matrix components still retained. The solvent volume should be adjusted so that quantitative recovery of the analytes is achieved with subsequent low dilution. In addition, the ﬂow-rate should be correctly adjusted to ensure efﬁcient elution. It is often recommended that the solvent volume be fractionated into two parts, and to allow the solvent soak the solid sorbent before the elution. 14.2.1.2 Retention of trace elements on the sorbent Adsorption of trace elements on the solid sorbent is required for preconcentration (see Fig. 14.2). The mechanism of retention depends on the nature of the sorbent, and may include simple adsorption, chelation or ion-exchange. Also, for trace elements, ion-pair SPE may be used. Adsorption Trace elements are usually adsorbed on solid phases through van der Waals forces or hydrophobic interaction. Hydrophobic interaction occurs when the solid sorbent is highly non-polar (reversed phase). The most common sorbent of this type is octadecyl-bonded silica (C18-silica). More recently, reversed polymeric phases have appeared, especially the styrene – divinylbenzene copolymer. Hydrophobic compounds are retained on such sorbents by hydrophobic interaction when aqueous samples are percolated through the sorbent, as well as by p–p interaction when p-electrons are present in the analyte [4]. Elution is usually performed with organic solvents, such as methanol or acetonitrile (ACN). Such interactions are usually preferred in online systems, as they are not too strong and thus they can be rapidly disrupted. However, because most trace element species are ionic, they will not be retained by such sorbents. Therefore, to retain trace elements, chelating agents

395

V. Camel

Fig. 14.2. Interactions occurring at the surface of the solid sorbent. ES: elution solvent; F: functional group; MI: matrix ions; ML: matrix ligands; MS: matrix solvent; TE: trace element.

may be added to the sample to form complexes, which are sufﬁciently hydrophobic to be further retained on the sorbent by hydrophobic interaction. The chelating agents may also be deposited on the hydrophobic sorbent, either by physical sorption or by chemical bonding, as discussed later.

396

Solid-phase extraction

Chelation Chelating resin may be prepared by treating a hydrophobic sorbent (such as C18-silica) with a reagent having a hydrophobic group (for sorption on the sorbent) along with a chelating group (for chelation of trace elements). Another way of obtaining such sorbents is to treat an ion-exchange resin with a reagent having both an ion-exchange group (for its sorption on the resin) and a chelating group. Hence, several reagents have been examined for preparing a chelating resin from an anion-exchange resin [5]. Best results were obtained using the sulfonic acid derivative of dithizone (DzS) as compared to other reagents such as dithizone (i.e. diphenylthiocarbazone) or thiosalicylic acid. The loading of 5-sulfo-8-quinolinol has also been reported on an anionexchange resin [6]. The complexation process in the resulting chelating resin was found to be the rate-limiting step for sorption of some trace elements (Co, Cd, Mn, Ni, and Zn). Sulfonated chelating reagents were also loaded on an anion-exchange resin (Dowex 1-X8) [7]. In such case, retention was based not only on ion-exchange, but also on molecular absorption between the resin matrix and the chelating agent. Thus, not all the chelating groups present in the loaded resin were available for further chelation of metal ions. Several functional group atoms are capable of chelating trace elements. The atoms most frequently used are nitrogen (e.g. N present in amines, azo groups, amides, nitriles), oxygen (e.g. O present in carboxylic, hydroxyl, phenolic, ether, carbonyl, phosphoryl groups), and sulfur (e.g. S present in thiols, thiocarbamates, thioethers). These groups may be introduced into the solid sorbent through different means: (1) the synthesis of new sorbents containing such groups (new sorbents), (2) the chemical bonding of such groups on existing sorbents ( functionalised sorbents), (3) the physical binding of the groups on the sorbent by impregnating the solid matrix with a solution containing the chelating ligand (impregnated, coated, or loaded sorbents). In practice, inorganic cations may be divided into three groups: –

–

–

Group I—“hard” cations: these preferentially react via electrostatic interactions (due to a gain in entropy caused by changes in orientation of hydration water molecules); this group includes alkaline and alkalineearth metals (Ca2þ, Mg2þ, Naþ) that form rather weak outer-sphere complexes with only hard oxygen ligands. Group II—“borderline” cations: these have an intermediate character; this group contains Fe2þ, Co2þ, Ni2þ, Cu2þ, Zn2þ, Pb2þ, Mn2þ. They possess afﬁnity for both hard and soft ligands. Group III—“soft” cations: these tend to form covalent bonds. Hence Cd2þ and Hg2þ possess strong afﬁnity for intermediate (N) and soft (S) ligands.

The nature of the functional group will give an idea of the selectivity of the ligand towards trace elements. For soft metals, the following order of donor atom afﬁnity is observed: O , N , S. A reversed order is observed for hard cations. For a bidentate ligand, afﬁnity for a soft metal increases with the

397

V. Camel

overall softness of the donor atoms: (O,O) , (O,N) , (N,N) , (N,S). The order is reversed for hard metals. In general, the competition for a given ligand essentially involves metals of Groups I and II for O sites, and metals of Groups II and III for N and S sites. The competition between metals of Groups I and III is weak. Different ligands immobilised on a variety of solid matrices have been used successfully for the preconcentration, separation, and determination of trace metal ions, as discussed later. As an example, carboxyphenylporphyrin (TCPP) has been investigated recently as a complexing agent for the preconcentration of some divalent metal ions [8]. The macroporous anion-exchange resin Amberlite IRA-904 was used as a solid support. Two different procedures were evaluated: TCPP complexation in the solution and further retention of the metal complexes; TCPP immobilisation on the solid support. In both cases, the selectivity order was as follows: Pb(II) . Ni(II) . Cu(II) . Cd(II) . Mg(II). It is well known that binding of metal ions to the chelate compound, either in solution or loaded on solid support, is dependent on several factors, such as: nature, charge and size of the metal ion; nature of the donor atoms present in the ligand and buffering conditions that favour certain metal extraction; and binding to active donor or groups. With a chelating exchanger, processes are slower than with an ion-exchanger, and strongly depend on the nature of the sorbent as well as on the degree of cross-linkage of polymeric sorbents. For an efﬁcient retention of metal species, either by chelation or sorption, the functional group should be accessible to the species without steric hindrance. In practice, many functional groups present on the sorbent surface may remain inactive in complexation because equilibrium cannot be attained. In some cases, the behaviour of immobilised chelating sorbents towards metal preconcentration may be predicted using the known values of the formation constants of the metals with the investigated chelating agent, as observed for Eriochrome black-T (ERT)-functionalised silica gel [9]. However, the presence of the solid sorbent may also have an effect and lead to results different from that observed in a homogeneous reaction. Thus, Chelex-100, a chelating ion-exchange resin having iminodiacetate groups, was less selective than was expected based on complexation results in aqueous solutions, and retained calcium at much lower pH than expected [10]. This was attributed to the formation of a 1:2 complex due to the large excess of active groups in the resin phase, whereas in solution only the 1:1 complex is reported. Similar observations were made for Amberlite CG-50, a chelating resin containing carboxylic active groups [11]. Also, the length of the spacer arm between the resin and the bound ligand was found to have an effect, due to a decrease in ligand ﬂexibility for short spacer arms, as observed with iminodiacetate-type resins [12]. In addition, the pore dimensions of silica gel have been shown to control the chemistry between a loaded chelating agent (dimethylglyoxine, DMG) and Ni(II) [13]. Thus, with microporous silica, the stoichiometry of the Ni(II)–DMG complex was 1:1 rather than 1:2 in aqueous solutions.

398

Solid-phase extraction

Ion-pairing When a non-polar sorbent is to be used, an ion-pair reagent (IP) can be added to the sorbent [14]. IP reagents contain a non-polar portion (such as a long aliphatic hydrocarbonated chain) and a polar portion (such as an acid or a base). The non-polar portion interacts with the reversed-phase non-polar sorbent, while the polar portion forms an ion-pair with the ionic species present in the matrix. As an example, different quaternary ammonium salts enabled the formation of ion-pairs with the Cu–8-hydroxyquinoline-5-sulfonic acid complex, and their subsequent retention on Amberlite XAD-8 [15]. Also, the ion-pairing reagent, sodium dodecylsulfate (SDS), has been loaded on C18-silica as the long alkyl chain is strongly adsorbed on this reversed phase, and the resulting phase enabled the retention and separation of several alkaline-earth metals [16]. Ion-exchange Ion-exchange sorbents usually contain cation or anion functional groups bonded to silica gel or to a polymer. Strong and weak sites refer to the fact that strong sites are always present as ion-exchange sites at any pH, while weak sites are only ion-exchange sites at pH values greater or lower than the pKa. Strong cation-exchange sorbents contain sites of sulfonic acid groups, and weak cation-exchange sorbents contain sites consisting of carboxylic acid groups. Strong anion-exchange sites are quaternary amines, and weak anion-exchange sites are primary, secondary, and tertiary amines. Ion-exchange sorbents derived from silica gel are not as rugged as polymeric ion-exchange resins because the silica matrix is susceptible to dissolution by strong acids or bases. In practice, a typical ion-exchange resin is based on a cross-linked styrene – divinylbenzene polymer. In general, ion-exchangers recover hydrated ions, charged complexes, and ions complexed by labile ligands. The most popular ion-exchanger has been Chelex-100 (or Dowex A1), with iminodiacetate groups. It offers very high distribution ratios for transition metals ions and for Hg2þ and Pb2þ, but not for alkali ions [17]. In addition, its chemistry can be well predicted by analogy with ethylene diamine tetraacetic acid (EDTA). Many other ion-exchangers have been successfully used. As an example, Dowex 1-X8, a strongly basic anionexchanger, enabled the retention of the negative Fe(III) –ferron complex, allowing the separation of labile Fe(III) from Fe(II) and its further determination [18]. Inorganic selenium could also be retained on a strong anion-exchanger (SAX); the subsequent selective elution enabled determination of both Se(IV) and Se(VI) [19]. An ion-exchanger may be characterised by its capacity, resulting from the effective number of active functional groups per unit of mass of the material. The theoretical value depends upon the nature of the material and the form of the resin. However, in the column operation mode, the operational capacity is usually lower than the theoretical one, as it depends on several experimental

399

V. Camel

factors such as ﬂow-rate, temperature, particle size, and concentration of the feed solution. In fact, retention on ion-exchangers depends on the distribution ratio of the ion on the resin, the stability constants of the complexes in solution, the exchange kinetics, and the presence of other competing ions. Usually, ionexchangers are of limited use for preconcentration of trace elements due to their lack of selectivity and their retention of major ions [17]. 14.2.1.3 Elution of trace elements from the sorbent The same type of interactions discussed before usually occur during the elution step. This time, the type of solvent must be correctly chosen to ensure stronger afﬁnity of the trace element for the solvent, to ensure disruption of its interaction with the sorbent (as illustrated in Fig. 14.2). Thus, if retention on the sorbent is due to chelation, the solvent could contain a chelating reagent that rapidly forms a stronger complex with the trace metal. Elution may also be achieved using an acid that will disrupt the chelate and displace the free trace element. Similarly, if retention is due to ion-exchange, its pH dependence enables the use of eluents with different pH’s to be used, such as acids. Of prime importance is to selectively elute only the target species. So, if they are more strongly retained on the sorbent than the interferent compounds, a washing step with a solvent of moderate elution strength is highly advisable before elution of the target species with the appropriate solvent. 14.2.2 Operation The sorbent may be packaged in different formats: ﬁlled micro-columns, cartridges, syringe barrels, and discs [2,20,21]. The disposable sorbent containers are illustrated in Fig. 14.3.

Fig. 14.3. Disposable sorbent containers.

400

Solid-phase extraction

14.2.2.1 Micro-columns The use of a micro-column is a common procedure for extraction of trace elements from various samples. It affords the opportunity of packing the column with the desired sorbent, so that a broader choice than the commercially disposable containers is available. In addition, the size of the column (i.e. the sorbent weight) may be adapted to the sample volume. In particular, it allows larger sample volumes, thus enabling the preconcentration of metal ions at very low concentration levels. However, such columns must be reused, so that careful blank washings should be conducted to avoid crosscontamination. In addition, columns with a narrow internal diameter limit usable ﬂow-rates to a range 1 – 10 ml/min that necessitates long traceenrichment times for large sample volumes. As an example, the optimum ﬂow-rate for loading several metal ions on quinalizarin-functionalised Amberlite XAD-2 (1 g) packed in a glass column (1 cm £ 10 cm) was found to be between 1.5 and 3.0 ml/min [22]. Above 3 ml/min there was a decrease in adsorption efﬁciency. As will be discussed later, micro-columns are frequently used in systems affording the on-line coupling of SPE to analytical techniques. However, in that case, the size of the column is limited to achieve acceptable analytical performance. 14.2.2.2 Disposable cartridges and syringe barrels Nowadays, the most frequently used design in off-line SPE is the cartridge or the syringe barrel. They are usually made of polypropylene or polyethylene and ﬁlled with packing material having different functional groups. The solid sorbent is contained between two 20-mm polypropylene frits (in some cases they may be made of glass). They afford great selectivity due to the broad types of sorbents contained in commercially available systems, with different column volumes available. In addition, the use of disposable cartridges and syringe barrels prevents possible cross-contamination, which can result when employing a column for multiple uses. Cartridges vary from as little as 100 mg to 1 g or more. Syringe barrels range in size from 1 to 25 ml and packing weights from 50 mg to 10 g. Solvent reservoirs may be used at the top of the syringe barrels to increase the total volume. They are typically 50–100 ml in volume. The barrel of the syringe terminates in a male Luer tip, which is the standard ﬁtting to be used with various SPE vacuum manifolds available. In the case of cartridges, both a female and male Luer tips are available, to enable use of either positive pressure or negative pressure from a vacuum manifold. The major disadvantages of cartridges and syringe barrels are slow sampleprocessing rates and a low tolerance to blockage by particles and adsorbed matrix components, which are caused by the small cross-sectional area of extraction cartridge. Channelling reduces the capacity of the cartridge to retain analytes and results in contamination of the isolated analytes with impurities

401

V. Camel

originating from the manufacturing and packing process. Such contaminants were evident for C18-silica cartridges when using gas chromatography (GC)– electron capture detection (ECD) or GC –mass spectrometry (MS) as the analysis method [23,24]. Comparatively lower contaminants were observed with C18-silica discs [24]. 14.2.2.3 Discs The use of ﬂat discs with high cross-sectional area may largely prevent all the problems encountered with columns, cartridges, and tubes [21]. In the discs, the packing material is frequently embedded in an inert matrix of polytetraﬂuoroethylene (PTFE) microﬁbrils. The typical composition of the discs is 90% w/w sorbent and 10% w/w PTFE ﬁbres [25]. One of the most popular discs is the Emporew disc, which consists of 8– 12 mm particles of packing material embedded in an inert matrix of PTFE ﬁbrils. Other types of discs available use a glass-ﬁbre matrix rather than Teﬂon to hold the sorbent particles, in order to enable higher ﬂow-rates to pass through the discs. The discs are available in different diameters from 4 to 90 mm, the size most frequently used being 47 mm. They are designed to be used in conjunction with a ﬁltration apparatus connected to a water aspirator [25]. In order to remove potential interferences and to ensure optimal extraction of the analytes of interest, disc cleaning and conditioning should be done before its use. Due to a lower void volume and higher surface area associated with small particles as compared to cartridges, partitioning of the analytes is favoured. Hence, a smaller mass of sorbent is required to process a similar volume of sample. Discs thus present the advantage of reducing solvent volumes for both the conditioning and elution steps. Additionally, the decreased backpressure encountered with these devices enables the use of high ﬂow-rates, and their wide bed minimises the chance of plugging. In addition, new technology for embedding the stationary phase prevents channelling and improves mass transfer, especially when 8-mm microparticulates are used. As classical discs are dedicated to the SPE of large-volume samples, new systems have very recently emerged that enable the use of discs for small-volume samples: the extraction disc cartridge (the disc is placed in a syringe-barrel format), and the 96-well microtitre plate conﬁguration [20,21,26]. Such systems are typically dedicated to biological samples. One of the drawbacks of using discs instead of cartridges or syringe barrels is the relatively low capacity, which may lead to breakthrough and incomplete recovery in some cases. As a consequence, discs are recommended when there is a strong interaction between the analyte and the sorbent. As an example, a comparison of C18-silica cartridge and PTFE-supported C18-silica disc for the retention of Cu –bathocuproine complex showed that longer ﬁltration time was required for the C18-silica cartridge (around 20 min as compared to 5 min) [27]. However, discs have lower capacity (e.g. 20 mg) so that for real samples (e.g.

402

Solid-phase extraction

high content of natural organic matter in river water) incomplete retention of the target metal species may result.

14.2.3 Advantages of the technique 14.2.3.1 Comparison to LLE Compared to classical LLE, SPE is an attractive technique as it reduces consumption of, and exposure to, solvents, disposal costs, and extraction time [28]. With an appropriate sorbent and elution solvent, quantitative recoveries may be achieved using SPE, provided samples are appropriately treated when non-labile metal species have to be retained [29,30]. Besides, SPE enables the simultaneous preconcentration of trace elements and removal of interferences, with possible enrichment factors up to 1000. Finally, SPE affords a broad range of applications due to the large choice of solid sorbents. 14.2.3.2 Preservation and storage of the species SPE allows on-site pre-treatment, followed by simple storage and transport of the pre-treated samples [21]. For instance, dithizone-coated C 18-silica cartridges enabled the on-site sampling of water samples, with stability of the retained mercuric complexes (at 48C) for at least 2 weeks [31]. Also, copper was stable on micro-columns packed with 1-(2-pyridylazo)-2-naphthol (PAN) coated Amberlite XAD-4 for at least 2 weeks at 48C [32]. This is a major advantage of SPE as the transport of the sample and its storage until analysis may induce problems, especially changes in speciation. In addition, the space occupied by the solid sorbents is minimal and avoids storage of bulky containers as well as manpower required to handle them. 14.2.3.3 High selectivity SPE offers the opportunity of selectively extracting only the trace elements of interest, thereby avoiding the presence of major ions. This is crucial in some cases, such as with spectrophotometric detection [33]. It may also be possible to selectively retain some particular species of a metal, enabling speciation. For example, Cu2þ was selectively retained on salen I modiﬁed C18-silica discs [34]. Also, the chemical binding of formylsalicylic acid onto amino-silica gel enabled selective Fe(III) extraction [35]. This high selectivity may be used to remove organic substances present in the sample that may hinder metal determination, such as the removal of lipids from biological samples [36]. 14.2.3.4 Automation and on-line coupling to analysis techniques SPE can be easily automated using commercial or home-made systems [26,37]. In addition, it can be interfaced on-line with analytical techniques. This avoids sample manipulation between preconcentration and analysis steps, so that analyte losses and risks of contamination are minimised, allowing higher

403

V. Camel

reproducibility [38]. All the sample volume is analysed, which enables smaller sample volumes. On-line coupling to liquid chromatography On-line SPE–liquid chromatography (LC) systems mainly use an extraction sorbent placed in a precolumn. The same packing should be used in the precolumn and the chromatographic column, to prevent losses in efﬁcacy upon analysis. When two different sorbents are used, retention of the analytes in the precolumn should be lower than in the analytical column, to ensure band refocusing at the head of the chromatographic column. The on-line coupling of SPE to LC with ultraviolet (UV) detection has been reported for the determination of chromium species [39] or Pb, Cd, and Hg in aqueous samples [40]. SPE coupled to an LC –inductively coupled plasma mass spectrometer (ICP-MS) has been applied to the determination of selenium species [41]. Also, SPE has been reported to enable the on-line coupling of supercritical ﬂuid extraction (SFE) with LC [42]. Very recently, the coupling of SPE to LC –cold vapour atomic absorption spectrometry (CV-AAS) via ﬂow injection has been reported and used for the determination of Hg species [43]. On-line coupling to atomic absorption spectrometry Olsen et al. [44] and Fang et al. [45,46] were the ﬁrst to describe an on-line ﬂowinjection (FI) sorbent extraction system for ﬂame AAS (F-AAS). Later, they also proposed a system for on-line FI sorbent extraction with electrothermal vaporisation AAS (ET-AAS) [47]. Since then, numerous papers reported FI with on-line preconcentration followed by AAS, as exempliﬁed by determination of Cu, Cr(VI), or Pb [48– 50]. Selected applications are reported in Table 14.1 (see also Chapter 21). The sorbent should enable rapid sorption and desorption of the analytes [65]. In addition, it should provide for a high selectivity. C18-silica has been widely used because methanol (MeOH) or ethanol (EtOH) can be used as eluting solvents, leading to a better sensitivity in F-AAS. Several complexing agents can be used such as sodium diethyldithiocarbamate (DDTC) [59,60,91]. Other sorbents were found satisfactory for some applications, as shown in Table 14.1, such as polyurethane foam (PUF) [92] or PTFE [48–50,74,75,77, 93]. On-line coupling to ICP The ﬁrst report of FI on-line preconcentration coupled to ICP-atomic emission spectrometry (OES) appeared nearly 20 years ago [94]. Since then, several studies have used this coupling, with different sorbents, as indicated in Table 14.1. Some also reported on-line coupling to ICP-MS [82,84,95]. On-line coupling to spectrophotometry The coupling of spectrophotometry to FI analysis is well suited for monitoring purposes and a few studies present such systems. Several chromogenic

404

TABLE 14.1 Applications of SPE for FI on-line preconcentration systems Trace elements

Chelating agent added

Sorbent

Eluent

Analysis method

Recovery (%)

Loading time (volume)

Preconcentration factor

LOD (ng/l)

Sampling Ref. frequency 21 (h )

Sea and waste waters

Cd

None

DPTHfunctionalisedSiO2

HNO3 – HCl

ICP-OES

97.5 – 104

1 min

86

1100

40

[51]

Sea and waste waters

Cd

None

TSfunctionalisedSiO2

Thiourea 2.5% in HNO3

ICP-OES

95.2 – 103.3 2 min

62

4300

24

[51]

Sea waters

Fe

None

8-HQfunctionalisedSiO2

HCl

Spectrophotometry

106.3

2 min

–

0.016 nM

–

[52]

Geological sample, Cu metal, Pb nitrate

Ag

None

2-MBTfunctionalisedSiO2

Thiourea

F-AAS

93.5 – 101

1 min

–

660

60

[53]

Certiﬁed ore samples, Ni alloy, anode slime, electrolytic solution

Ag, Au, Pd

None

AmidinothioureidoSiO2

Thiourea

F-AAS

98.7 – 101.4 1 min (4.5 ml)

–

1100– 17,000

–

[54]

Fish, human urine

MeHg, EtHg, PhHg, Hg(II)

APDC

C18-silica

MeOH– LC –CV-AAS ACN –water

92– 106

20 min (58.5 ml)

750–950

5.5– 10.4

2.3

[43]

Sea water

MeHg, Hg(II)

DDTC

C18-silica

EtOH

CV-AAS

85– 107.5

(25 ml)

500

16

–

[55]

Certiﬁed sea waters

Cu, Cd

APDC

C18-silica

MeOH

ET-AAS

86.7 – 106.5 104 s (0.5 ml)

25 –100

6.5– 1.26

–

[56]

Matrix

Inorganic sorbents

Solid-phase extraction

405

continued

406

TABLE 14.1 (continuation) Trace elements

Chelating agent added

Sorbent

Eluent

Analysis method

Recovery (%)

Loading time (volume)

Preconcentration factor

LOD (ng/l)

Sampling Ref. frequency 21 (h )

Certiﬁed sea water, mussel, geological samples

Cu, Cd, Co

1,10Phenanthroline

C18-silica

EtOH

F-AAS

88.9 – 100.5

30 s

22 –32

300–6000

90

[57]

Certiﬁed sea waters

Cd

PAR or PADMAP

C18-silica

MeOH

ET-AAS

82.5 – 111.2

110 s (0.5 ml)

25 –50

1.7– 4

9– 12

[58]

Sea water, industrial efﬂuents

Cr(III), Cr(VI), Cr(total)

DDTC

C18-silica

MeOH

F-AAS

95 – 105

60 –300 s

90 –500

20

30

[59]

Sea, river waters

Cr(VI), Cr(total)

DDTC

C18-silica

EtOH

ET-AAS

101– 105

1 min (3 ml)

12

1600–1800

22

[60]

Certiﬁed low alloy steel, mussel, tomato leaves

Co

NN

C18-silica

Acidiﬁed EtOH

F-AAS

98 – 102

30 s (3.25 ml)

17.2

3200

90

[61]

Certiﬁed sea waters

Pb

DDTC

C18-silica

MeOH

ET-AAS

94.9 – 115.8

1 min

26

3

24

[47]

Certiﬁed biological, vegetable samples

Pb

DDTC

C18-silica

IBMK

F-AAS

99.2 – 137.9

2 –10 min

60 –189

3000

24

[62]

Standard solutions

Pb

DDTP

C18-silica

EtOH

F-AAS

–

2.5– 75 min (10– 150 ml)

14 –1000

300–3000

–

[63]

Certiﬁed citrus leaves, marine sediment

MeHg, Hg(II)

DDTP

C18-silica

EtOH

CV-AAS

99.6 – 112.5

4.5 min (23.85 ml)

20

10

12

[64]

Standard solutions

Cu, Pb

DDTC, 8-quinolinol or PAR

C18-silica

MeOH

F-AAS

–

80 s (4 ml)

14 –60

4000–10,000

–

[65]

Sea water

Cu

None

TAN-loadedC18-silica

HCl

ET-AAS

93.2 – 99.6

1 min (3 ml)

33

5

35

[66]

V. Camel

Matrix

TABLE 14.1 (continuation) Trace elements

Chelating agent added

Sorbent

Eluent

Analysis method

Recovery (%)

Loading time (volume)

Preconcentration factor

LOD (ng/l)

Sampling Ref. frequency 21 (h )

Pharmaceutical preparations

Zn

None

TAN-loadedC18-silica

HCl

Spectrophotometry

89.8– 107.8

–

–

10,000

45

[67]

Sea water

Fe(II), Fe(III)

None

FZ-loadedC18-silica

MeOH

Spectrophotometry

–

2 – 20 min (4 – 40 ml)

6 –60

0.1 –0.3 nM

–

[68]

River, tap, rain waters

Al, Bi, Cd, Co, Cr, Cu, Fe, Ga, In, Mn, Mo, Ni, Pb, Tl, V, Sb, Sn, Zn

None

ZrO2

HNO3

ICP-OES

95.4– 99

33 min (100 ml)

100

6 –90

–

[69]

Waters

Cr(VI)

None

Acidic Al2O3

NHþ 4

ICP-OES

–

(2 ml)

10

200

–

[70]

Lake, river, tap waters

Cr(III), Cr(VI)

None

Acidic Al2O3

HNO3 or NHþ 4

F-AAS

90– 106

35 s

25

800– 1000

55

[71]

Sewage waters

Cr(III), Cr(VI)

None

Al2O3

HNO3 or NH4þ

F-AAS

86– 117

–

–

42,000–81,000

–

[72]

Urine

Cr(III)

None

Basic Al2O3

HNO3

ICP-OES

.93

(10 ml)

50

50

–

[73]

Tap, river, coastal waters

Cu

APDC

PTFE turnings

IBMK

F-AAS

94– 102

1 min (12 ml)

340

50

40

[48]

Tap, river, coastal, industrial waste waters

Cr(VI)

APDC

PTFE turnings

IBMK

F-AAS

95.5– 100.5 3 min (37.8 ml)

80

800

18

[49]

Tap, river, and coastal waters, marine sediment, ﬁsh, and mussel tissues

Pb

APDC

PTFE turnings

IBMK

F-AAS

95– 102

330

800

15

[50]

Organic sorbents

3 min (39 ml)

407

continued

Solid-phase extraction

Matrix

408

TABLE 14.1 (continuation) Trace elements

Chelating agent added

Sorbent

Eluent

Analysis method

Recovery (%)

Loading time (volume)

Preconcentration factor

LOD (ng/l)

Sampling Ref. frequency 21 (h )

Drinking, sea waters

Cr(VI)

APDC

PTFE (KR)

EtOH

ET-AAS

105

1 min (5 ml)

19

4.2

21.2

[74]

Certiﬁed natural water, sea water

Cr(VI)

APDC

PTFE (KR)

EtOH

ET-AAS

–

1 min (5 ml)

16.3

16

16.7

[75]

Certiﬁed human hair, pig liver, sea prawn

Cd, Co, Cu, Zn

Dithizone

PTFE (KR)

IBMK

F-AAS

95.3 – 108.4 1 min (5 ml)

23.4– 69.3

1060– 2560

18

[76]

Certiﬁed human hair and rice powder

Cd

DDTC

PTFE (KR)

IBMK

F-AAS

97.9 – 110

50 s (4.2 ml)

30

100

55

[77]

Certiﬁed natural water, sea water

Cr(VI)

APDC

PTFE beads

EtOH

ET-AAS

104– 108

1 min (5 ml)

30.1

8.8

16.7

[75]

Standard solutions

Pb

DDTP

PUF

EtOH

F-AAS

–

2.5– 75 min (10– 150 ml)

14 –1000

300–3000

–

[63]

Certiﬁed sea waters

Cu

None

PAN-coatedXAD-4

HCl – EtOH

F-AAS

99.6 – 103

(2.5 – 25 ml)

29.1– 296.1

60 –600

–

[32]

Standard samples

Cd, Zn

None

BSQ-loadedXAD-7

HCl

Fluorimetry

–

20 s (0.1 ml)

10

1600– 1900

–

[78]

Alloys, ores

Pd, Pt, Rh

ODETA

Highly crosslinked polystyrene

HCl – EtOH

F-AAS

77.8 – 103.6 1 min

–

3000– 8000

30

[79]

Tap water

Cu, Cd, Ni

–

IDA-Novarose

HCl

ICP-OES

–

10 min

500–1000

–

–

[80]

River, ground waters

Cu, Cd, Pb

Sulfasarzene

Lewatit TP807’84

HCl

Spectrophotometry

80 – 120

50 min (100 ml)

50

2000– 5500

–

[33]

River, mineral and tap waters

Cr(III)

None

PAPhA

HCl

F-AAS

97 – 101

90 s (6.6 ml)

35

200

30

[81]

V. Camel

Matrix

TABLE 14.1 (continuation) Eluent

Analysis method

Recovery (%)

Loading time (volume)

Preconcentration factor

LOD (ng/l)

Sampling Ref. frequency 21 (h )

Cu, Cd, Ni, None Zn, Mn

Toyopearl AFChelate 650 M

HNO3

ICP-MS

87 – 110

1 min (1 ml)

–

1.4– 86

–

[82]

Synthetic sea water

Cd, Cu, Pb, Zn

None

Cationexchanger

HNO3

F-AAS

–

–

50– 105

30– 200

60

[45]

Natural waters

Cr(VI), Cr(total)

DPC

Cationexchanger

HNO3 – acetone

Spectro-photometry

93 – 108

–

–

8.9– 15.2

–

[83]

Certiﬁed pig kidney, rye grass, rice ﬂour, tomato leaves

Cd, Ni, Pb

None

Cationexchanger

HCl

ICP-MS

96.7 –103.7

2 min (7.8 ml)

10

1000–4000

90

[84]

Ground water

Cu

Bathocuproine

Anionexchanger

HNO3

Spectrophotometry

–

10 min (8.3 ml)

–

80

–

[85]

Water, vegetable samples

Cd

None

Anionexchanger

HNO3

Spectrophotometry

94 – 104

90 s (5 ml)

–

230

20

[86]

Standard solutions

Pb

DDTP

Activated carbon EtOH

F-AAS

–

2.5– 75 min (10– 150 ml)

14– 1000

300– 3000

–

[63]

Steels, Al solutions Tap, mineral, well, river, swimming pool waters

Bi

DDTP

Activated carbon EtOH

ET-AAS

87 – 104.3

14

48

7

[87]

Cu

APDC

Activated carbon IBMK

F-AAS

88 – 113

4 min (10 ml) 2 min (4.5 ml)

100

600

17

[88]

Silicon, tap water

Fe

None

Activated carbon HCl

MPT-OES

97.4 –105

1 min (1.2 ml)

4.3– 6.4

1000–36,000

–

[89]

Rock, copper ore

Pd

None

Activated carbon Thiourea

F-AAS

103– 107

3 min (15.6 ml)

145

300

15– 20

[90]

Trace elements

Estuarine waters

Chelating agent added

Solid-phase extraction

Sorbent

Matrix

409

V. Camel

reagents have been used, such as sulfasarzene [33], malachite green [86], or ferrozine [68]. Solid-phase spectrophotometry (SPS) has also been reported with FI systems due to its simplicity and low detection limits. The solid sorbent is directly packed in the ﬂow cell, from which retained analytes are periodically removed using an acid or a complexing solution [67,83,85]. The main disadvantages of on-line FI sorbent extraction systems are possible insufﬁcient adsorption of metal species on the sorbent and clogging of the column when insoluble ligands are used [96]. So, in the case of complex samples, off-line SPE should be preferred due to its greater ﬂexibility, and the opportunity to analyse the same extract using various techniques.

14.3

STEP-BY-STEP METHOD DEVELOPMENT GUIDE

Development of an SPE method can be considered as a two-step procedure. First, the most appropriate sorbent for the application should be chosen. Optimisation of the most inﬂuential parameters should then be undertaken. Obviously, optimisation should initially be performed using spiked synthetic solutions, but it must be followed by the use of certiﬁed reference materials or spiked real samples, as matrix components (such as ligands or other ions) may change the trace element retention on the sorbent, thereby decreasing recoveries of the target species. As an example, while Chelex-100 enabled the extraction of metallic species (Cu, Pb, Cd, and Zn) from a synthetic sea water, a fraction of the metals remained unretained in the case of real sea water [30], probably due to chelate formation in the sample when organic matter is present, or to adsorption on colloidal particles not retained due to the low resin pore size. 14.3.1 Selection of solid sorbent Solid sorbents may be hydrophobic or polar. It is common to call reversed-phase sorbents the packing materials that are more hydrophobic than the sample. They are frequently used with aqueous samples. On the other hand, normalphase sorbents refer to materials more polar than the sample. They are used when the sample is an organic solvent containing the target compounds. With hydrophobic supports, retention of ionic metal species requires the formation of hydrophobic complexes. This can be achieved through addition of the proper reagent to the sample, or thorough immobilisation of the reagent on the hydrophobic solid sorbent. Addition of reagent to the sample is appropriate for the immobilisation of unstable metal species to maintain speciation, while immobilisation offers the convenience of having a prepared cartridge or disc before analysis. Immobilisation may also provide a signiﬁcant development in speciation analysis because metal equilibrium in the sample may not be affected by reaction with the sorbent.

410

Solid-phase extraction

The nature and properties of the sorbent are of prime importance for effective retention of metallic species. Careful choice of the sorbent is thus crucial to the development of SPE methodology. In practice, the main requirements for a solid sorbent are: (1) possibility to extract a large number of trace elements over a wide pH range (along with selectivity towards major ions), (2) fast and quantitative sorption and elution, (3) high capacity, (4) regenerability, and (5) accessibility. Sorbents that allow fast reaction rates are preferred to achieve faster extractions as well as higher loading capacities. Hence, sorbents based on hydrophilic macroporous polymers, cellulose, or ﬁbrous materials, provide excellent kinetic properties [97]. The broad variety of sorbents available explains one of the most powerful aspects of SPE, which is selectivity. The selectivity of the modiﬁed solid phases towards certain metal ions is attributed to several well-known factors, such as the size of the organic compound used to modify the sorbent, the activity of the loaded surface groups, and the type of the interacting functional group. However, the selective extraction of a single trace element from other interfering ion(s) represents a direct challenge for ﬁnding a suitable phase capable of exhibiting a sufﬁcient afﬁnity to selectively bind that metal ion. For particular applications, the combination of two sorbents may thus be advisable. As an example, the passage of water samples through two successive chelating resins enabled the determination of trace and major elements [98]. Similarly, the combination of an anion- and a cation-exchange resin enabled the speciation of Cu and Mn in milk samples [99]. Sorbents can be mainly categorised as inorganic- and organic-based ones. Immobilisation of organic compounds on the sorbent surface is usually aimed at modifying the surface with certain target functional groups for a higher selectivity of the extraction. 14.3.1.1 Inorganic-based sorbents Inorganic-based sorbents are mainly made of silica gel, even though other inorganic oxides may be used (cf. Fig. 14.4). Silica gel-based sorbents present the advantages of mechanical, thermal, and chemical stability under various conditions. They frequently offer a high selectivity towards a given metal ion. However, all silica-based sorbents suffer from different chemical limitations, namely the presence of residual surface silanol groups and a narrow pH stability range. Applications of such sorbents to off-line SPE are presented in Tables 14.2 and 14.3. Silica gel Silica gel can be used as a very successful adsorbing agent as it does not swell or strain, has good mechanical strength and can undergo heat treatment. The surface of silica gel is characterised by the presence of silanol groups, which are known to be weak ion-exchangers, causing low interaction, binding, and extraction of ionic species [131]. Thus, several trace elements were found to be

411

V. Camel

Fig. 14.4. Sorbents based on inorganic supports.

retained on this sorbent [132]. Retention is highly dependent on sample pH as, under acidic conditions, silanol groups are protonated and the ion-exchange capacity of the silica gel is greatly reduced, or even reduced to zero, at low pH’s. In addition, silica gel has a very low selectivity and is prone to hydrolysis at basic pH’s. So, modiﬁcation of the surface with speciﬁc organic compounds has been performed, either by physical adsorption (impregnated or loaded sorbent) or chemical immobilisation ( functionalised sorbent). In the ﬁrst case, the organic compound is directly adsorbed on the silanol groups, while in the second approach a chemical bond is formed between the silica gel surface groups and those of the organic compound. Several methods for impregnating reagents (ion-exchangers or chelating compounds) on adsorbents have been used, principally by passing the reagent solution through a column packed with the adsorbent, or soaking the adsorbent in the reagent solution. Numerous reagents have been investigated for impregnation of silica gel as a means of increasing retention capacity and selectivity of the sorbent for trace elements. These include 1-nitroso-2-naphthol (NN) [103], thionalide (2mercapto-N-2-naphthylacetamide) [101,102], 2-mercaptobenzothiazole (2MBT) [133], 8-hydroxyquinoline (8-HQ) [134,135], 3-methyl-1-phenyl-4-stearoyl-5-pyrazolone (MPSP) [100], salicylaldoxime [132], DMG [13], Aliquat 336 (methyl-tricaprylammonium chloride), and Calcon (hydrophobic sodium sulfonate) [136]. The chemical binding of chelating functional groups on silica gel and its application to the determination of trace elements have also been

412

TABLE 14.2 Applications of off-line SPE using inorganic supports to water samples Trace elements

Sorbent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

Tap water

Cu/Co/Ni

MPSPloaded-SiO2

Glass column

Sample pH: 4.5, elution: HCl 1 M

F-AAS

94.6 – 101

43/45/49 mmol/g

40

60/40/70

[100]

Sea water

Pd

Thionalideloaded-SiO2

Glass column (1 cm i.d.)

F-AAS

83 – 99

0.8 mg/g

3200

0.03

[101]

Sea water

As(III)

Thionalideloaded-SiO2

Glass column (1 cm i.d.)

Spectrophotometry

92 – 95

5.6 mmol/g

–

0.12

[102]

River and sea water

Co

NNloaded-SiO2

Glass column

g-Emission

96 – 98

0.03 mmol/g

10– 100

–

[103]

Spiked tap water

Hg(II)

Column

CV-AAS

99 – 99.5

300 mmol/g

200

3.96

[104]

Tap and sea waters

Hg(II)

DithizonefunctionalisedSiO2 DithioacetalsfunctionalisedSiO2

Sample pH: 4, washing, elution: thiourea 0.2 M þ HCl 0.1 M Sample pH: 7, washing, elution: NaBorate/ NaOH/ Iodine (pH 10) Sample pH: 3.5, washing, elution: acetic acid Elution: HCl 10 M

Column

Elution: water

CV-AAS

91 – 100

917–1100 mmol/g

5

–

[105]

Spiked tap and sea waters

Hg(II)

DithiocarbamatesfunctionalisedSiO2

Glass column

Elution: water

CV-AAS

88 – 100

0.6– 0.983 mmol/g

–

–

[106]

Matrix

Silica

Solid-phase extraction

413

continued

414

TABLE 14.2 (continuation) Trace elements

Sorbent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

Sea water

V/Co/Ni/ Ga/Y/Mo/ Cd/Cr/Pr/ Nd/Sm/Eu/ Gd/Tb/Dy/ Mo/Er/Tm/ Yb/Lu/W/U

8-HQfunctionalisedﬂuorinated metal alkoxide glass

Column (6 mm i.d., 30 mm bed height)

Washing, sample pH: 5, washing, elution: HNO3 0.5 M backﬂush

ICP-MS

85 – 116

–

10

0.00037– 2200 ng/l

[107]

Cr(III)/Cr(VI)

TiO2

Glass column (1 cm i.d.)

ET-AAS

78.4 –99.2

8125/6983 mg/g

100

0.030/0.024

[108]

Natural, waste, sea waters

Cd/Co/Cu/Fe/ Mn/Ni/Pb

TiO2

Glass column (1 cm i.d.)

Sample pH: 2 or 8, elution: HNO3 0.5 or 1 M Sample pH: 8, elution: HNO3 1 M and/or EDTA 0.1 M

F-AAS

89 – 100

5000 mg/g

300

0.01– 0.04

[109]

Tap, ground waters

Cr(III)/ Cr(VI)

Neutral Al2O3

Column (1 cm i.d.)

ET-AAS

99 – 100

–

25

0.01

[110]

Tap, ground waters

Se(IV)/Se(VI)

Acidic Al2O3

Teﬂon column (1 cm i.d.)

Sample pH: 6.5 –7, elution: NH3 1 M þ HNO3 4M Sample pH: 2 – 8, elution: NH3 0.1 M and 4 M

ET-AAS

90 – 98

23.2/ 2.0 mg/g

16/100

0.049– 0.80

[111]

Other oxides Rain, river, sea, tap waters

V. Camel

Matrix

TABLE 14.3 Applications of off-line SPE using C18-silica-based supports to water samples Trace elements

Reagent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

Sea water

TBT

None

Cartridge or Empore disc (25 mm)

Conditioning, sample, drying, elution: acidiﬁed ethyl acetate

GC-ECD

93.5– 111.5

–

1000

–

[24]

Spiked sea waters

TPhT

None

Bond Elut cartridge

Conditioning: MeOH þ NaCl, sample, washing, air drying, elution: 1024 M hydroxy ﬂavone in MeOH backﬂush

Fluorimetry

81 –89

–

100

0.07

[112]

Sea water

TPhT

None

Bond Elut cartridge (40 mm)

Washing, sample, washing, air drying, elution: MeOH

Fluorescence (after addition of ﬂavonol to the eluate)

–

–

250

–

[113]

Matrix

No reagent

Sea water

Se/Sb

APDC

Glass column (1.4 cm i.d.)

Sample pH: 1.2, washing, elution: MeOH

ET-AAS

94 –97

–

40 –75

0.007/ 0.05

[114]

Sea water

Cd/Zn/Cu/ Mn/Fe/Ni/Co

8-HQ

Glass column (1.4 cm i.d.)

Sample pH: 8.9, washing (water þ oxine), elution: MeOH

ET-AAS

67 –108

–

50 –100

–

[115]

Neocuproine

Empore discs (47 mm)

Washing, conditioning: MeOH, sample pH: 5.0, drying, elution: isopentyl alcohol

Spectrophotometry (454 nm)

99.7– 102.6

940 mg Cu2þ

50 –100

0.12

[116]

Loaded reagent on the sorbent River water

Cu

415

continued

Solid-phase extraction

Addition of the reagent to the sample

416

TABLE 14.3 (continuation) Trace elements

Reagent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

CRM water (SLRS-3), lake, river, drinking waters

Cu(I)

Bathocuproine

Bond Elut cartridge Empore disc

Conditioning: MeOH þ water, sample pH: 4.3, elution: MeOH –water 90:10 (v/v)

Spectrophotometry (484 nm)

–

–

20 – 40

0.40 –3.8

[27]

Tap waters, well water

Fe

Bathophenanthroline

Empore discs (47 mm)

Washing, activation: MeOH þ water, sample pH: 4 –7, elution: EtOH þ NaClO4

Spectrophotometry (533 nm)

99.2 –100.9

–

–

0.080

[117]

Synthetic sea waters

MeHg/ PhHg/ Hg(II)

Dithizone

Sep Pak cartridge

Sample pH: 4 þ EDTA 0.001 M, washing, elution: MeOH

LC-DAD

95 – 104

–

200

0.14/0.16/ 0.14

[31]

Rain, lake, river waters

MonoBT/DiBT/ TBT/ MonoPhT/ DiPhT/TPhT

Tropolone

Sep Pak cartridge

Conditioning: MeOH, sample pH: 2 –3, elution: diethyl ether

Ethylation – GC –FPD

–

–

–

–

[118]

Tap, well and river waters

Cu

Quinone derivative

Empore discs (47 mm)

Washing, conditioning: buffer, sample pH: 7.0, drying, elution: HNO3 0.1 M

F-AAS

98.4 –102

360 mg Cu2þ

400

0.2

[119]

Tap, rain, snow and sea waters

Cu(II)

Schiff ’s base (salen I)

Empore disc (47 mm)

Washing, sample pH: 5.5– 6, air drying, elution: HNO3 0.1 M

AAS

–

396 mg Cu2þ

. 500

0.004

[34]

Synthetic and spring waters

Pb(II)

Schiff ’s base

Empore disc

Washing, sample pH: 2 –8, air drying, elution: HNO3 0.5 M

F-AAS

97.1 –100.2

700 mg/ disc

50

16.7

[120]

V. Camel

Matrix

TABLE 14.3 (continuation) Trace elements

Reagent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

River water

Pb(II)

BAS

Empore disc (47 mm)

Washing, sample pH: 2– 7, air drying, elution: acetic acid 1 M

F-AAS

–

476 mg Pb2þ

$300

0.050

[121]

Sea waters

Fe(II)

Ferrozine

Sep Pak cartridge

Conditioning: MeOH þ water, sample pH: 6.8 –8.3, washing, elution: MeOH

Spectrophotometry (562 nm)

91

–

40

0.6 nmol/l

[122]

Rain, sea waters

Fe(II)

Ferrozine

Sep Pak cartridge

Conditioning: MeOH þ water, sample, washing, elution: MeOH

LC-UV (254 nm)

92– 99

–

100–500

0.1 nmol/l

[123]

Certiﬁed sea waters (NASS-2 and SLEW-1)

Cu/Cd

APDC

Teﬂon cartridge (0.94 mm i.d.)

Conditioning: MeOH þ water, sample pH: 6– 8, air drying, elution: MeOH

ET-AAS

95.8 – 103.3

–

25– 50

0.0024/ 0.00018

[124]

Tap and spring waters

U(IV)

TOPO

Empore disc (47 mm i.d.)

Washing, conditioning, sample, elution: MeOH

Spectrophotometry

85

4033 mg/disc

8

0.1

[125]

continued

Solid-phase extraction

Matrix

417

418

TABLE 14.3 (continuation) Trace elements

Reagent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

Sea waters

La/Ce/Pr/ Nd/Sm/ Eu/Gd/Tb/ Dy/Ho/ Er/Tm/ Yb/Lu

HDEHP/ H2MEHP

Sep Pak cartridge

Sample pH: 3 –3.5, washing, elution: HCl 6 M

ICP-MS

88.8 – 99.8

–

200–1000

–

[126]

Spiked natural waters

Bi

Cyanex 301

Cartridge (0.5 g)

Conditioning: HCl 0.1 M, sample pH: 1, air drying, elution: HNO3 3 M

ET-AAS

98.5 – 100

–

10 –100

105

[127]

Sea, well and tap waters

Be

Quinalizarine

Sep Pak cartridge

Washing, conditioning, sample pH: 6– 6.6, elution: HNO3 0.5 M

F-AAS

98– 101

200 mg

200

0.2

[128]

Tap, spring waters

Agþ

HT18C6

Empore disc

Conditioning: MeOH þ water, sample, elution: Na2S2O3 0.1 M

AAS

996–100.3

210 mg Ag/disc

200

0.050

[129]

Tap, river, well and spring waters

Hg(II)

HT18C6TO

Empore disc (47 mm i.d.)

Washing, sample pH: , 7, air drying, elution: HBr 1 M

CV-AAS

97.1 – 101.3

241 mg Hg2þ

50

0.006

[130]

V. Camel

Matrix

Solid-phase extraction

reported for more than 20 years. Hence, diamines [137], triamines [13], thioamines [138], dithiocarbamates [137], imino-dithiocarbamates [106], and dithioacetal derivatives [105] were bound to SiO2. Both physical adsorption and chemical immobilisation approaches have been tested for dithizone [104], which is well known to participate in metal binding via nitrogen and sulfur donor atoms [139]. The chemically modiﬁed silica gel was much more stable and less prone to hydrolysis than the physically modiﬁed phase, which was expected due to the higher strength of covalent bonds as compared to hydrogen bonds and van der Waals interactions. In addition, a higher capacity and a faster equilibration were observed with the chemically modiﬁed sorbent [104]. Other chemically immobilised ligands include formylsalicylic acid and purpurogallin (both Fe(III) selective) [35,140], ERT [9], N-propylsalicylaldimine [141], calixarene tetrahydroxamic acid [142], phosphonic acid [143], 2,5dimercapto-1,3,4-thiadiazole [144], 2-MBT [53], 1,5-bis(di-2-pyridyl) methylene dithiocarbohydrazide (DPTH) [51], and methylthiosalicylate (TS) [51]. Silica gel bound macrocycles (SGBMs) have also been synthesised, such as 18-crown6-bound-silica gel [145,146]. It must be kept in mind that, despite chemical bonding of functional groups, free silanols still remain [143]. Their number can be minimised by end-capping the sorbent, but some will still be present. As a consequence, they will participate in the retention of trace elements somewhat, especially at pH’s above their pKa (ionised form). C18-bonded silica gel Numerous applications report the use of C18-silica, as indicated by the studies reported for water samples in Table 14.3. Indeed, this sorbent has currently become the most popular phase used. As acidic Si –OH groups are still present on the surface of the C18-silica, this sorbent can retain ionic species, such as 10% of Cu2þ ions present in water [34]. In addition, organometallic compounds can be retained, such as organic selenium [19], tributyltin (TBT) [24], or triphenyltin (TPhT) [112]. However, due to its hydrophobic character, C18-silica is not well suited for retention of trace element species, as the latter are often polar or ionic. Retention may be improved by addition of a ligand reagent to the sample before its percolation through the sorbent. The hydrophobic part of the ligand will thus have hydrophobic interaction with the C18-silica and be retained on the sorbent, while the functional group of the ligand will ensure chelation of the trace elements. For that purpose, several ligands were used, such as 8-HQ [115], ammonium pyrrolidine dithiocarbamate (APDC) [114], 1,10-phenanthroline [18], or bathocuproine [27]. An alternative approach is to form the complex by passing the sample through a C18-silica already containing the ligand. C18silica loaded with different ligands offers the possibility of concentrating trace amounts of metal ions. In addition, by carefully choosing the ligand, selectivity may be added to the extraction step. Among applications, dithizone was used

419

V. Camel

for inorganic and organic mercury [31], N,N0 -diethyl-N0 -benzoylthiourea (DEBT) for palladium [147], salen I for Cu2þ ions [34], neocuproine for Cuþ ions [116], ferrozine (FZ) for Fe(II) [68,122,123], APDC for Cu and Cd [56,124], DDTC for lead [62]. Organic phosphorus has been also used as they possess highly extractive properties for trace elements, such as tri-n-octylphosphine oxide (TOPO) [125], tri-n-butyl phosphate (TBP) [125], a mixture of bis(2ethylhexyl) hydrogen phosphate (HDEHP) and 2-ethylhexyl dihydrogen phosphate (H2MEHP) [126], or Cyanex 301 [(2,4,4-trimethylpentyl) dithiophosphinic acid] [127]. The loading of quinones is also attractive, such as quinalizarin [128], BAS (bis[1-hydroxy-9,10-anthraquinone-2-methyl]sulﬁde) [121], 11-hydroxy-naphthacene-5,12-quinone [119], and 1-(2-tiazolylazo)2naphthol (TAN) [66,67]. Macrocycles may also be loaded on C18-silica, as recently reported for hexathia-18-crown-6 (HT18C6) [129], hexathia-18-crown6-tetraone [130], and calixarene hydroxamate [142]. For some particular applications, mixed ligand complexes may be used to ensure synergistic adsorption of the metal complex on the solid sorbent. Thus, while Cu(II) ions cannot complex with neutral TBP molecules adsorbed on C18silica contained in a cartridge, the form of Cu(TTA) complex (TTA being 2thenoyltriﬂuoroacetone) was retained at around 80% [148]. Alternatively, in some cases, loading the chelating agent on C8-silica instead of C18-silica gave better results [149]. Yet, the main drawbacks of bonded silica are the limited range of pH that can be used (usually 4–8), with the result that polymeric sorbents may be preferred. Other inorganic oxides Apart from silica, other inorganic oxides have been tested for their retention of trace elements (see Table 14.2). Adsorption of ions is believed to proceed via participation of hydroxyl groups. These groups are negatively charged (deprotonated) under basic conditions, thereby retaining cations, and positively charged (protonated) under acidic conditions, thereby retaining anions. Consequently, acidic SiO2 is expected to adsorb only cations, while basic magnesia (MgO) should adsorb anions. On amphoteric oxides [namely titania (TiO2), alumina (Al2O3), zirconia (ZrO2)], cations are adsorbed under basic conditions (pH above the isoelectric point of the oxide [109]) while anions are adsorbed under acidic conditions (pH below the isoelectric point of the oxide). The adsorption properties of many oxides also strongly depend on the characteristics of the solid: crystal structure, morphology, defects, speciﬁc surface area, hydroxyl coverage, surface impurities, and modiﬁers. Activated alumina may be used in acidic, neutral, or basic form. Its acidic form was reported to allow chromium speciation, as careful adjustment of the sample pH enabled either Cr(VI) or Cr(III) retention [70,71]. To avoid pH change of the sample (which may affect speciation), basic alumina may be used for selectively retaining Cr(III) [73]. More recently, neutral alumina has been shown to retain both Cr(III) and Cr(VI) [110,150]. Acidic alumina also enabled

420

Solid-phase extraction

the retention of both Se(IV) and Se(VI) from water samples without any pH adjustment [111]. Its coating with an anionic surfactant was also investigated for the selective retention of Cr(VI) in very acidic solutions [151]. Several studies reported the efﬁciency of TiO2 in preconcentrating trace elements [108, 109,152,153]. The use of zirconia (ZrO2) has also been reported [69]. 14.3.1.2 Organic-based sorbents Organic-based sorbents may be divided into polymeric and non-polymeric sorbents, as shown in Fig. 14.5. Polymeric sorbents have been, by far, the most used for trace element preconcentration, having the advantage over bonded silica in that they can be used over the entire pH range. Their disadvantage is that the conditioning step is more time consuming. Extensive reviews on polymeric phases have been published, with emphasis on application to organic micropollutants [97,154,155]. The purpose of this section is to summarise the most frequently used organic-based sorbents for trace elements, as well as the more recently reported ones. In most applications, new sorbents have been synthesised by chemically bonding chelating groups to polymeric cross-linked chains. Most of the chelating groups reported have low water solubility to avoid their leaching from the sorbent. At the same time, a very hydrophobic group will hinder wettability of the sorbent by the aqueous sample, resulting in poor retention efﬁciency. A compromise is thus necessary. In addition to the functional group, the efﬁciency of polymeric sorbents depends on various physico-chemical parameters, such as particle size, surface area, pore diameter, pore volume, degree of crosslinking, and particle size distribution.

Fig. 14.5. Sorbents based on organic supports.

421

V. Camel

Polystyrene – divinylbenzene-based sorbents Macroporous hydrophobic resins of the Amberlite XAD series are good supports for developing chelating matrices. Amberlite XAD-1, XAD-2, XAD-4, and XAD16 are polystyrene– divinylbenzene (PS–DVB) resins with a high hydrophobic character and no ion-exchange capacity. In addition to the hydrophobic interaction that also occurs with C18-silica, such sorbents allow p – p interactions with aromatic analytes. Compared with C18-bonded silica gel, these resins offer the advantage of being more stable against acidic or alkaline aqueous solutions and against all organic solvents. However, they require extensive cleaning before use. In addition, as with C18-bonded silica, PS – DVB presents a poor surface contact with aqueous samples due to its hydrophobic surface. Consequently, conditioning the sorbent with an activating solvent, such as methanol, acetone, or ACN, is required to obtain a better surface contact with water samples. The combination of a polydentate ligand of small molecular size and a polymeric support with moderate cross-linking enables the design of chelating resins with high capacity. Among ligands added to the sample for trace element retention on Amberlite XAD-2, APDC [156,157] and 8-HQ [158,159] are very common. Inorganic ligands may also be added, and inorganic complexes retained as observed for iodo complexes on Amberlite XAD-16 [160]. However, organic ligands are preferred as they may offer selectivity. Thus, addition of diphenylcarbazide (DPC) enabled selective retention of Cr(VI) on Amberlite XAD-16 [161]. Ligands may also be attached to the PS –DVB sorbent by physical adsorption, as shown with dithizone [162], 3-(2-pyridyl)-5,6-diphenyl1,2,4-triazine (PDT) [4], tropolone [163,164], or PAN [165] in the case of Amberlite XAD-2. Efﬁcient sorption of ligands on Amberlite XAD-4 was also reported, such as PAN [32], 7-dodecenyl-8-quinolinol (DDQ) [166], ADPC [167], piperidine dithiocarbamate (pipDTC) [167], 2-(5-bromo-2-pyridylazo)-5(diethylamino)-phenol (5-BrPADAP) [168], and calixarene hydroxamate [142]. In practice, as partial leaching of the impregnated ligand may occur, chemically functionalised resins may be preferred for extensive use. Several chemical modiﬁcations of PS–DVB have recently been reviewed [169], but only a few are commercially available. The ligands are generally coupled to a methylene or an azo spacer on the matrix. In particular, Amberlite XAD-2 was functionalised through an azo spacer with several reagents, namely Alizarin Red-S [170], salicylic acid [171], thiosalicylic acid (TSA) [172], Pyrocatechol Violet (PV) [173], chromotropic acid (CA) [174], pyrocatechol (PC) [175,176], Tiron [177], and quinalizarin [22]. Storage of such resins for several months under ambient conditions had no effect on their performance. Amberlite XAD-4 functionalised with several ligands, such as bicine [178] or poly(dithiocarbamate) (PDTC) [179], also enabled the retention of trace elements. Sulfonated PS –DVB resins have been also reported, with an excellent hydrophilicity [154, 155]. Such resins may act as cation-exchangers or, in the case of rapid sulfonation under mild conditions, as mixed-mode sorbents (i.e. adsorption of

422

Solid-phase extraction

neutral compounds on the polymeric resin, and cation-exchange of ionic species on sulfonate groups). As an example, sulfonated Amberlite XAD-4 enabled preconcentration of the cationic cobalt-(5-BrPADAP) chelate [180]. The anionic functional group can also be attached to the polymer through an azo spacer as reported for 2-naphthol-3,6-disulfonic acid (NDSA) [181]. Trimethylammonium functionalised PS–DVB have also been synthesised, with anion-exchange properties. Despite the broad applications of PS–DVB, other polymers have also been used (cf. Table 14.4). Divinylbenzene –vinylpyrrolidone copolymers Sorbents made of divinylbenzene –vinylpyrrolidone (DVB–VP) copolymers have recently been developed [154]. The hydrophilic N-vinylpyrrolidone affords good wettability of the resin, while the hydrophobic divinylbenzene provides reversed-phase retention of analytes. It has been successfully applied to the determination of polar organic compounds in water samples. It is convenient to use, as it can dry out during the extraction procedure without reducing its ability to retain analytes, and it is stable over the entire pH range. However, until now, no application related to the preconcentration of trace elements has been reported. Similarly, no application has been reported for trace elements for a sulfonated DVB–VP [154], which combines the properties of the previous sorbent with those of a strong cation-exchanger. Polyacrylate polymers Amberlite XAD-7 and XAD-8 are ethylene-dimethacrylate resins. They are non-aromatic in character and possess very low ion-exchange capacity. Due to the polarity of acrylates, such resins enable the recovery of polar compounds. Amberlite XAD-7 has been efﬁciently coated with several ligands, such as 8(benzenesulfonamido)-quinoline (BSQ) [78], Xylenol Orange (XO) [176,185], 5BrPADAP [168], or dimethylglyoxal bis(4-phenyl-3-thiosemicarbazone) (DMBS) [196]. A study also reported the chemical bonding of dithizone to poly(ethylene glycol dimethacrylate-hydroxyethyl-methacrylate) microbeads, and its use for retention of inorganic and organic mercury [197]. Polyurethane polymers Due to its sorption capacity for several trace elements, PUF has been tested for use in SPE. Most of the time, it is coated with complexing reagents to enhance the sorption capacity, such as dimethylglyoxime [198], 1-nitroso-2-naphthol [199], DDTC [199], or hexamethylenedithiocarbamate (HMDC) [200]. Alternatively the reagent can be added directly to the sample, and the formed complex further retained on PUF, as shown for thiocyanate complexes [92,201, 202]. PUF has also been used as the solid support for developing an enzymatic test procedure for Pb(II), after its impregnation with alkaline phosphatase immobilised in N-phthalylchitosan [203].

423

424 TABLE 14.4 Applications of off-line SPE using polymeric sorbents to water samples Matrix

Trace elements

Sorbent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

Tap water

Cd/Cu/Mn/ Ni/Pb/Zn þ 8-HQ

XAD-2

Polypropylene column

Conditioning, sample pH: 8 –9, elution: HCl 2 M

ICP-AES

82.3 – 97.2

–

100

–

[159]

Tap water

Cr(VI), total Cr þ DPC

XAD-16

Glass column (1 cm i.d.)

Washing, sample pH: 1, elution: H2SO4 0.05 M /MeOH

F-AAS

97.3 – 99.0

0.4 mg/g

5.25

45

[161]

Drinking and sea waters

Bi/Cd/Co/ Cu/Fe/Ni/ Pb þ APDC

Chromosorb102

Glass column (0.9 cm i.d.)

Washing, conditioning, sample pH: 6, elution: acetone

F-AAS

95 – 110

–

300

0.10 –11

[157]

Tap, mineral, river waters

Co þ 8-HQ

Chromosorb105

Column (4 mm i.d.)

Washing, sample pH: 8, elution: EtOH/ HNO3 2 M

ET-AAS

95.2 – 99.2

–

80

0.0134

[182]

Tape, lake, waste waters

Cr(III)

Cellulose

Syringe barrel

Puriﬁcation, sample pH: 11, elution: HCl 2 M

ET-AAS

98 – 99.3

–

100

0.0018

[183]

TBT

Tropoloneloaded-XAD-2

Glass column (1.5 cm i.d.)

Sample þ 0.8% H2SO4, washing, elution: IBMK

ET-AAS

104

–

80

0.0144

[164]

Adsorptive resins

Waters

V. Camel

Chelating resins

TABLE 14.4 (continuation) Trace elements

Sorbent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

Sea water

Co/Cu/Fe/ Ni/Zn

PDT-loadedXAD-2

Column (0.9 cm i.d.)

Washing, elution: MeOH in Soxhlet apparatus

F-AAS

96.3 – 103.5

–

–

–

[4]

Well water, river water

Cu/Cd/Co /Pb/Zn/Mn

QuinalizarinefunctionalisedXAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 5– 7, elution: HNO3 4 M

F-AAS

91 – 98

3.15/1.70/1.62/ 5.28/1.42/0.94/ 2.19 mg/g

100/50/40/50 /100/65/65

2.0/1.3/5.0 /15.0/1.0/1.6

[22]

Well waters

Zn/Cd/Pb/Ni

PV-functionalised-XAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 3– 7, elution: HNO3 4 M

F-AAS

98

1410/1270/ 620/1360 mg/g

60/50/23/18

–

[173]

Well waters

Zn/Pb

Salicylic acid-functionalised-XAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 5.0, elution: HCl 1 M—2–4 M

F-AAS

98 – 100

1146/461 mg/g

180/140

–

[171]

Well waters

Zn/Cd/Pb/Ni

Alizarin RedS-functionalised-XAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 4– 6, elution: HNO3 1–4 M or HCl 4 M

F-AAS

95 – 100

511/124/306/ 124 mg/g

40

10

[170]

River waters

Cu/Cd/ Co/Ni/Pb/Zn/ Mn/Fe

Tiron-functionalised-XAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 4– 7.5, elution: HNO3 4 M

F-AAS

91 – 99

–

25 – 200

0.5– 24

[177]

River waters

Cd/Co/Cu/ Ni/Fe/Zn

CA-functionalised-XAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 4 – 7, elution: HNO3 or HCl 2 M

F-AAS

95 – 100

9.35/3.84/8.50 /3.24/6.07/ 9.65 mg/g

100– 200

–

[174]

continued

Solid-phase extraction

Matrix

425

426

TABLE 14.4 (continuation) Trace elements

Sorbent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

River and tap waters

Pb

CA-functionalised-XAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 3 – 8, elution: HNO3 2 –10 M

F-AAS

97

186.3 mmol/g

200

4.06

[176]

River and tap waters

Cd/Co/Cu/ Fe/Ni/Zn

PC-functionalised-XAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 3 –6.5, elution: HNO3 2 M

F-AAS

–

0.023 –0.092 mmol/g

80 –200

–

[175]

River and tap waters

Pb

PC-functionalised-XAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 5 – 7.5, elution: HNO3 1M

F-AAS

94

104.7 mmol/g

100

3.80

[176]

River and tap waters

Pb

TSA-functionalised-XAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 4, elution: HNO3 0.5– 2 M

F-AAS

93

89.3 mmol/g

100

4.87

[176]

Tap, river waters

Cd/Co/Cu/ Fe/Ni/Zn

TSA-functionalised-XAD-2

Glass column (1 cm i.d.)

Washing, sample pH: 3.5 –7, washing, elution: HNO3 2 M

F-AAS

92 –98

197.5/106.9/214.0/ 66.2/309.9/ 47.4 mmol/g

180– 400

0.48/0.20/4.05/ 0.98/1.28/3.94

[172]

Artiﬁcial sea water, natural waters

Cd/Cu/Mn/ Ni/Pb/Zn

APDC-loadedXAD-4

Glass column (0.9 cm i.d.)

Sample pH: 5.0, washing, elution: HNO3 4 M

ICP-OES

98.2– 99.6

9.47/11.08/8.62/ 7.21/10.25/ 10.62 mg/g

180/230/120/ 130/160/215

0.1/0.4/0.3/ 0.4/0.6/0.5

[167]

Artiﬁcial sea water, natural waters

Cd/Cu/Mn/ Ni/Pb/Zn

pipDTCloadedXAD-4

Glass column (0.9 cm i.d.)

Sample pH: 5.0, washing, elution: HNO3 4 M

ICP-OES

97.6– 99.1

9.18/10.76/8.17/ 7.46/9.86/ 10.28 mg/g

150/200/140/ 120/150/200

0.7/1.0/0.8/ 0.9/1.7/1.2

[167]

V. Camel

Matrix

TABLE 14.4 (continuation) Trace elements

Sorbent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

Sea water

Ag/Al/Bi/Cd/ Cu/Fe/Ga/ Mn/Ni/Pb/Ti

DDQ-loadedXAD-4

Teﬂon column (8 mm i.d.)

Washing, sample pH: 8, washing, elution: HCl 2 M backﬂush

F-AAS or ET-AAS

73 – 107

0.55 mmol/g

62.5

0.00016– 0.3

[166]

Tap water

Cu/Mn/Zn

Calixarene tetrahydroxamate-loadedXAD-4

Plastic cartridge (0.9 cm i.d.)

Sample pH: 8.5, elution: acidiﬁed water (pH 2.0)

F-AAS

–

–

25

–

[142]

Tap and mineral waters

Mn

PDTC-functionalisedXAD-4

Glass column (1 cm i.d.)

Conditioning, sample pH: 10, elution: HNO3 8 M

F-AAS

97.2

9.1 mmol/g

20

0.5

[179]

Spiked solutions

Co/Cu/Fe/ Hg/Ni/Pb/Zn

Bicine-functionalisedXAD-4

Glass column

Conditioning, sample pH: 5.5– 7, elution: HCl 1 M

ET-AAS

97.6 –99.1

0.32 – 0.44 mmol/g

40 – 50

–

[178]

Waste waters

Cr(III)/ Cr(VI)

NDSA-functionalisedPS – DVB

Glass column (1 cm i.d.)

Sample pH: 1.5 or 6, elution: HCl 4 M

F-AAS

85.9 –96.1

0.40/1.18 mmol/g

20

–

[181]

River water

Hg(II)

PAAM-functionalisedPS – DVB

Glass column (1 cm i.d.)

Conditioning, sample pH: 5.4, elution: H2SO4 2M

Spectrophotometry

96

0.6 mmol/g

30

–

[184]

River and tap waters

Pb

XO-loadedXAD-7

Glass column (1 cm i.d.)

Washing, sample pH: 5, elution: HNO3 1 M

F-AAS

91

16.9 mmol/g

100

2.44

[176]

River waters

Cd/Co/Cu/Fe /Ni/Zn

XO-loadedXAD-7

Glass column

Sample pH: 4 – 5, elution: HCl 1 or 2 M

F-AAS

96 – 100

1.6 –2.6 mg/g

10 – 200

9/24/6/6/3/21

[185]

continued

Solid-phase extraction

Matrix

427

428

TABLE 14.4 (continuation) Trace elements

Sorbent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

River and reservoir waters

Cr(III)

8-HQ-functionalisedpolyacrylonitrile

Glass column (4 mm i.d.)

Washing, sample pH: 6, washing, elution: HCl 2 M/HNO3 0.1 M

ICP-MS

98– 105

41.7 mmol/g

5

0.06

[186]

Aqueous sample from a non-ferrous metal smelter

Au/Pt/Pd/Ir

Aminothiourea-functionalisedpolyacrylonitrile

Glass column (0.5 cm i.d.)

Washing, sample pH: 2, washing, elution: HCl 4 M/ CS(NH2)2 3%

ICP-OES

97– 99

2.80/1.75/ 1.56/1.15 mmol/g

6– 65

–

[187]

Sea water

Be/Bi/Co/Ga/ Ag/Pb/Cd/Cu /Mn/In

Aminophosphonicdithiocarbamate-functionalisedpolyacrylonitrile

Glass column (4 mm i.d.)

Sample pH: 6, washing, elution: HCl 2 M

ICP-MS

93– 104

0.83– 74.1 mmol/g

200

0.002 – 0.601 ng/l

[188]

River, lake, and rain waters

Hg(II)/MeHg

Dithiocarbamatefunctionalisedpolyvinyle

Column (1.5 cm i.d.)

Washing, sample pH: 1 –11, elution: thiourea 5% in HCl

CV-AAS

91– 95

–

667

0.0002

[189]

Sea water

Cd/Cu/Mn/ Ni/Pb/Zn

Chelamine

Column

Washing, sample pH: 6.5, washing, elution: HNO3 2 M

ET-AAS

91– 102

1 mmol/g

200

0.0023– 0.033

[190]

Sea water

Cd/Co/Cu/ Mn/Ni/Pb/Zn

Chelex-100

Column

Sample pH: 6.5, washing, elution: HNO3 2 M

ET-AAS or F-AAS

.98

–

50

0.001 – 0.1

[191]

River water, sea water

Cd/Pb/Zn

Amberlite IRC-718

Glass column (0.5 cm i.d.)

Washing, conditioning, sample, washing, elution: HNO3

F-AAS

63– 104

1.06/0.096/1.77 mmol/g

10

–

[192]

V. Camel

Matrix

TABLE 14.4 (continuation) Matrix

Trace elements

Sorbent

Operation

Experimental conditions

Analysis method

Recovery (%)

Adsorptive capacity

Preconcentration factor

LOD (mg/l)

Ref.

Ground and fresh waters

Se(IV)/ Se(VI)/ SeCyst

Amberlite IRA-743

Glass column (1 cm i.d.)

Washing, conditioning, sample, elution: HClO4 1 M þ water

LC –ICPMS

93 – 97

–

55

0.010

[193]

Wastewater

Cr

Anion-exchanger (SAX)

Cartridge (0.5 g)

Activation: MeOH þ buffer, sample pH: 4.5, elution: Na2SO4 0.5 M

Spectrophotometry (544 nm) after reaction DPC

80 – 98.8

–

–

8

[194]

Sea, river, tap waters

Se(IV)/ Se(VI)

Anion-exchanger (SAX)

Cartridge

Conditioning, sample pH: 7, elution: HCOOH 1 M þ HCl 3 M

GC –MS

91 – 99

–

40 – 500

1600/1400

[19]

River, spring, waste waters

Be

Anion-exchanger

Column (1 cm i.d.)

Conditioning: HCl þ H2O þ NaOH, sample pH: 6– 8, elution: HCl 1.5 M

F-AAS

95 – 102.5

–

125

0.045

[195]

Ion-exchangers

Solid-phase extraction

429

V. Camel

Polyethylene polymers Polyethylene is also attractive for SPE of trace elements, as this support adsorbs several metals complexed with hydrophobic ligands. The adsorbed complexes can be eluted with a small volume of organic solvents, allowing for high enrichment factors. Polyethylene can also be used in strongly acidic and basic media. Hence, in the presence of DPC, it enabled the retention of chromium in an acidic medium [204]. PTFE polymers Metals complexed with selected ligands may be retained on PTFE. Hence, after addition of APDC to the sample, metal –PDC complexes could be preconcentrated on PTFE, as observed for Cu(II), Cr(VI), and Pb(II) [48–50,74,75]. Dithizone was also found efﬁcient as a chelator [76]. PTFE may also be coated with a suitable ligand, as observed with 2-methyl-8-hydroxyquinoline for Co retention [93]. Polystyrene polymers The use of a hyper cross-linked polystyrene has been reported to be more efﬁcient than other sorbents (Amberlites XAD-2 and XAD-8, C18-silica) for retaining Rh, Pd, and Pt complexed with 4-(n-octyl)-diethylenetriamine [79]. Polyamide polymers The use of polyamide membranes for the batch retention of complexes of rare earth elements with Thorin has been reported [205]. Retention of the complexes was assumed due to both electrostatic forces and non-hydrophobic interaction. Iminodiacetate-type chelating resins Polymeric resins containing iminodiacetate groups [ –CH2 –N(CH2COO2)2] as active sites (IDA resins) have been widely used. They can be synthesised by bonding iminodiacetate functional groups to several polymeric sorbents [12]. Due to the weak acid character of the functional group, the degree of protonation will critically affect the ability of the resin to retain metal cations. Among the commercially available products, the cross-linked polystyrenebased Chelex-100 is one of the most well characterised. It has been used for years for extracting trace elements [10,29,30,191,206 –209]. In this resin, protonation of the carboxylates and the donor N atom are reported to be complete at pH 2.2, while a completely deprotonated form is reached at pH 12.3. Retention of some metal ions (e.g. Cu, Ni) may be predicted by the complexing properties of a monomer having a structure similar to that of the active groups. For other ions (e.g. Ca) sorption mechanisms are somewhat different. In any case, Chelex-100 suffers from a lack of selectivity, which limits its application to real samples as noted for sea water [30] or milk samples [99]. This often leads to a decrease in trace element retention, caused by retention of other species

430

Solid-phase extraction

(e.g. Ca, Mg) or complexation with ligands present in the sample such as organic matter [210]. So, other IDA resins have been used. For example, the macroporous Amberlite IRC-718 allowed preconcentration of Cd, Pb, and Zn from water samples, with a suspected negative steric effect on coordination with the iminodiacetate groups [192]. Another IDA resin, based on a highly crosslinked agarose gel (IDA-Novarose), enabled fast retention of Cu, Cd, Ni, and Ca [80]. However, in tap water the presence of dissolved organic matter reduced the retention of Cu and Ni by complexing these metals. Indeed, such complexes are in many cases more stable than the associations of the metals with iminodiacetate groups of the resin, which greatly limits the use of IDA resins for trace element retention from real samples. Propylenediaminetetraacetate-type chelating resins The synthesis of a ﬁne-particle macroporous polymer-based propylenediaminetetraacetic acid (PDATA) type resin has recently been reported [211]. Its structure is very similar to that of EDTA, with a spacer arm. The retention behaviour of several rare earth elements was evaluated and selectivity of the resin very close to that of free PDATA, indicating a great ﬂexibility of the ligand on the resin due to the spacer arm. Polyacrylonitrile-based resins Several functional groups have been chemically immobilised on a polyacrylonitrile ﬁbre, and the obtained resins successfully applied to the preconcentration of trace elements, such as aminophosphonic and dithiocarbamate groups [188], 8-HQ [186,212], or aminothiourea [187]. Ring-opening metathesis polymerisation-based polymers New polymers have also been synthesised using ring-opening metathesis polymerisation (ROMP) [213,214]. Due to the polymerisation sequence, most of the functional groups were located at the surface of the polymer beads, making them easily accessible. Hence, a carboxylic acid-functionalised resin enabled the retention of rare earth elements from rock digests [213], while dipyridyl amide-functionalised resin allowed the extraction of “soft” metals such as Pd(II) and Hg(II) [214]. Carbon sorbents Classical activated carbons are prepared by low-temperature oxidation of vegetable charcoals. Due to their large surface areas, these sorbents are well recognised for their strong sorption, both for trace organic compounds and trace elements. There is evidence of two types of adsorption sites: (1) graphite-like basal planes, that enable adsorption through van der Waals forces, especially p-electron interactions; (2) polar groups like carbonyls, hydroxyls, and carboxyls that may interact via ionic interaction or hydrogen bonding [215]. Consequently, trace elements may be directly adsorbed on activated carbon as

431

V. Camel

observed for selenium species [19,216]. Complexing reagents may also be added to the sample for further retention of metal chelates, like amino acids [217], dithizone [218], APDC [88,219], PAN [220], 8-HQ [221], cupferron [221], Bismuthiol II [222], or O,O-diethyl-dithiophosphate (DDTP) [63,87]. However, in some cases, destruction of the chelates may occur on the sorbent, probably due to the strong adsorption of the ligand on the active surface of the carbon and its reduced efﬁciency as a chelant [215]. The main drawback of classical activated carbons lies in their heterogeneous surface containing active functional groups that often leads to low reproducibility. In addition, they are very reactive and can act as catalysts for oxidation and other chemical reactions. Fortunately, a new generation of carbon sorbents appeared in the 1970s and 1980s, with a more reproducible and more homogeneous structure than the classical activated carbons. These are graphitised carbon blacks (GCBs), obtained from heating carbon blacks at 2700 –30008C in an inert atmosphere [155] and considered as both reversedphase sorbents and anion-exchangers, and porous graphitised carbons (PGCs), obtained after immobilisation of graphite on a silica substrate. Hence, a GCB (Carbopack B) was shown to selectively retain tributyltin and triphenyltin from water samples [223]. Cellulose Cellulose alone may retain trace elements, as observed for Cr(III) [183]. Trace elements may also be retained as complexes, as observed upon addition of the chelating agent calmagite in the sample [224]. Finally, cellulose may also be chemically functionalised. Hence, an anion-exchanger could be obtained by functionalising cellulose with a quaternary amine, with possible retention of both Se(IV) and Se(VI) [225]. Naphthalene-based sorbents Trace elements can be retained on microcrystalline naphthalene after formation of a neutral ion-pair as observed with Cu –TAN complex in the presence of sodium tetraphenylborate (TPB) [226]. Functionalised naphthalene may also be synthesised, such as 2,4,6-tri-2-pyridinyl-1,3,5-triazine (TPTZ)– TPB – naphthalene [227] or tetradecyldimethyl-benzyl-ammonium iodide (TDBAþI2)-naphthalene [228]. 14.3.2 Inﬂuential parameters The main experimental variables that affect analyte recovery by SPE have been extensively reported by Poole et al. [2,229]. They are brieﬂy discussed below and illustrated with reported applications.

432

Solid-phase extraction

14.3.2.1 Conditioning parameters Washing step A washing step is highly recommended, especially when ultratrace concentrations of elements are to be determined. Thus, blank extracts containing trace levels of Zn, Cu, and Fe were suspected to be due to contaminants from C18-silica [115]. As another example, the use of C18-silica Sep Pak cartridge for the preconcentration of rare earth elements required pre-washing of the solid sorbent with 6 M HCl to remove picogram amounts of the analytes [126]. Conditioning solvent Even though some sorbents have been used without a conditioning step, this is not recommended. This step will at least remove possible remaining contaminants and air from the sorbent bed. Additionally, in some cases, this step is crucial for successful retention of the analytes. As an example with hydrophobic supports, such as C18-silica, quite polar organic solvents such as methanol should be used, as, for example, in the case of C18-silica loaded with 11-hydroxynaphthacene-5,12-quinone [119]. The nature of the conditioning solvent must be appropriate to the nature of the solid sorbent to ensure good wettability of the functional groups. The sorbent should further be conditioned by a solvent whose nature is similar to that of the sample. Thus, for aqueous samples, the solvent will be water with a pH and ionic strength similar to that of the sample. 14.3.2.2 Loading parameters Sample volume to be percolated An important parameter to control in SPE is the breakthrough volume, which is the maximum sample volume that should be percolated through a given mass of sorbent after which analytes start to elute from the sorbent, resulting in nonquantitative recoveries (Fig. 14.6). The breakthrough volume is strongly correlated to the chromatographic retention of the analyte on the same sorbent, and depends on the nature of both the sorbent and the trace element, as well as on the mass of sorbent considered and the analyte concentration in the sample [3]. In addition, it depends on the sorbent containers, as discs usually offer higher breakthrough volumes than cartridges. As an example, above 600 ml of sample, non-quantitative recoveries were observed for Fe and Bi retained on Chromosorb-102 as their APDC-complexes [157]. The breakthrough volume of salen I modiﬁed C18-silica discs was reported to be higher than 2.5 l for retention of Cu2þ from water [34]. In the case of BAS modiﬁed C18-silica discs, its value was greater than 1.5 l for Pb2þ [121]. Naturally, the breakthrough volume is also strongly dependent on the nature of the sample, in particular, the possible presence of ligands. As an example, the breakthrough volume of Cu(II) onto Amberlite CG50 decreased from 236 l in the absence of ligands, to 429 ml in the presence of glycine [230]. The breakthrough volume may be

433

V. Camel

Fig. 14.6. Typical representation of the breakthrough curve (i.e. concentration of the analyte at the outlet of the SPE system versus sample volume percolated through the system). VB is the breakthrough volume, VR is the chromatographic elution volume, VC is the sample volume corresponding to the retention of the maximum amount of analyte, and C0 is the initial analyte concentration in the sample.

determined experimentally or estimated using several methods, as already detailed in Ref. [229]. Sample ﬂow-rate The sample ﬂow-rate should be optimised to ensure quantitative retention along with minimisation of the time required for sample processing. The value of the ﬂow-rate may have a direct effect on the breakthrough volume, and elevated ﬂow-rates may reduce the breakthrough volume, as reported [4,229]. As a rule, cartridges and columns require lower maximum ﬂow-rates than discs. Hence, quantitative retention of Cr(III) on 8-HQ-immobilised-polyacronitrile ﬁbre packed in a glass column required ﬂow-rates below 0.6 ml/min [186]. The maximum value was reported to be 3 ml/min for uptake of trace elements on CA-functionalised-Amberlite XAD-2 resin packed in a microcolumn [174]. This value may be increased by a factor of 10 using discs. For example, in the range of 5–70 ml/min, the retention of Hg(II) on modiﬁed C18silica membrane disc was quantitative [130], while in the range of 5 –50 ml/min Agþ was quantitatively retained on HT18C6-loaded-C18-silica discs [129]. Retention of iron ions by C18-silica discs was not signiﬁcantly affected by ﬂowrates of 8–75 ml/min for the sample solution [117]. In the range 5–25 ml/min, Cu(II) was retained by 11-hydroxynaphthacene-5,12-quinone modiﬁed C18silica discs [119]. Sample pH Sample pH is of prime importance for efﬁcient retention of trace elements on the sorbent. Its inﬂuence strongly depends on the nature of the sorbent used.

434

Solid-phase extraction

Careful optimisation of this parameter is thus important to ensure quantitative retention of the trace elements and, in some cases, selective retention. In the case of cation-exchangers, low pH usually results in poor extraction due to competition between protons and cationic species for retention on the sorbent. As an example, the optimum pH range for adsorption of several metals [Cu(II), Cd(II), Co(II), Pb(II), Zn(II), and Mn(II)] on quinalizarin-loaded Amberlite XAD-2 was found to be 5–7 [22]. The pH has also been shown crucial for retention of trace elements on Chelex-100 [29,209]. A pH near 6.5 was recommended for sea water samples, as compared to an optimum value around 5 for synthetic samples. By contrast, it was found that for all metal ions tested, retention on Lewatit TP807’84 resin increased as the pH increased. Moreover, for sample pH adjusted to 3.0 –3.5, quantitative retention of Zn(II) and Fe(II) was achieved, whereas there was no signiﬁcant adsorption of Cd(II), Cu(II), Pb(II), and Ni(II) [33]. Passing the ﬁrst aliquot of sample at pH 3.2 through such a resin enabled the removal of most of the Zn(II) and Fe(II); subsequent adjustment to pH 5.5 targeted species such as Cd(II), Cu(II), and Pb(II) without any Ni(II) retention. Adjustment of pH could also be used to control selectivity of trace element retention on salicylaldoxime-loaded-silica gel [132]. Thus, Cu(II) may be separated from other metals by varying the pH of the solution. In the same manner, Ni(II) and Co(II) can be separated from Zn(II) and Fe(III), while Zn(II) can be separated from Fe(III). With regard to the retention of inorganic selenium on an anion-exchanger (SAX), optimum pH was found to be near 7–8 [19]. When retention of trace elements is based on chelation (either in the sample or on the solid sorbent), the sample pH is also a very important factor, as most chelating ligands are conjugated bases of weak acid groups and, accordingly, they have a very strong afﬁnity for hydrogen ions. The pH will determine the values of the conditional stability constants of the metal complexes. For example, in the case of 11-hydroxynaphthacene-5,12-quinone, due to the presence of a hydroxy group in the ligand structure, the retention of Cu(II) increases with pH until about 6.7 is reached, where quantitative retention occurs [119]. In the case of C18-silica modiﬁed with the Schiff ’s base salen I, retention of copper was incomplete below pH 5 [34]. Retention of copper– neocuproine complex onto C18-silica discs was found to be quantitative in the pH range 4–8.5 [116]. Above these pH’s, the recoveries decreased, probably due to competition of NH3 with neocuproine and formation of a non-retained Cu(NH3)4 2þ complex. Adjustment of the sample pH also enabled the speciation of chromium using a NDSA-functionalised-PS –DVB [181]: in acidic media Cr(VI) was the species retained, whereas in neutral media only Cr(III) was absorbed, both being eluted with HCl. By contrast, pH may have no inﬂuence with some organic ligands, as evidenced by C18-silica impregnated with hexathia-18-crown-6-tetraone and its retention capacity towards Hg2þ in the 1–7 pH range [130].

435

V. Camel

For inorganic oxides, pH is also of prime importance. In particular, on amphoteric oxides such as TiO2 or Al2O3, cations are adsorbed at elevated pH’s due to the deprotonation of functional groups, whereas anion retention requires acidic conditions for the protonation of functional groups. Hence, retention onto TiO2 was found to be optimum for most trace elements at pH above 7–8 [109]. In the case of acidic alumina, retention of inorganic selenium species (SeO3 22 and SeO4 22) was quantitative for pH values up to 8 [111]. Above this pH, retention decreased due to the presence of negative charges on the surface of alumina. Sample matrix The presence of other ions or ligands in the sample matrix may reduce the retention of the target species onto the sorbent. The ionic strength may also have an effect. Presence of other ligands. The presence of ligands in the sample matrix may affect trace element retention when stable complexes are formed in the sample with these ligands, as the trace elements are less available for further retention. Thus, if metals are present in the sample as strong complexes, they may not dissociate, resulting in no retention of the free metal on the sorbent. As an example, the addition of EDTA at concentrations over 1027 M was found to reduce the recovery of several trace elements from spiked artiﬁcial sea water using DDQ-impregnated-XAD-4 [166]. Reduction in the retention of copper onto Amberlite CG50 occurs in the presence of ligands such as glycine [230]. The effect of adding iminodiacetic acid to synthetic sea water had a direct effect on Cu(II) and Cd(II) adsorption on Chelex-100, shifting the sorption curves to higher pH values [208]. In the case of real samples, the presence of natural organic matter is of great concern, as it may complex trace elements. In particular, Cu(II) has been shown to form stable water-soluble complexes with organic materials [231,232]. In some cases, the presence of ligands may be a valuable tool for adding selectivity to the SPE step. Hence, the buffering of water samples to pH 4 and the addition of EDTA avoided the retention of metal ions other than mercury species on C18-silica coated with dithizone [31]. Presence of other ions. The presence of ions other than those of interest in the samples may cause problems during the SPE step. In particular, due to their usually high levels [e.g. Ca(II)], they may hinder the preconcentration step by overloading the sorbent, or cause interferences during spectrophotometric analysis. Therefore, their inﬂuence should be studied before validating an SPE method. The effect of several ions on the sorption of trace elements by Amberlite XAD-4 resin coated with APDC or pipDTC showed that various cations and anions present in water samples do not interfere, as maximum tolerance limits were 40 and 50 g/l, respectively, for Ca2þ and Mg2þ, 50 and 70 g/l for Kþ and Naþ, and 40 g/l for Cl2, NO3 2, SO4 22, and CH3COO2 [167]. The effect of other metal ions in the sample has also been studied to test the

436

Solid-phase extraction

selectivity of different modiﬁed silica gels [104 –106]. A very high selectivity for Hg(II) was obtained when the silica gel was modiﬁed with sulfur-containing organic compounds (distribution coefﬁcient values Kd of 50,000 for a dithizone sorbent). A low retention of Cu(II) was also reported (Kd of 2750 for the same sorbent). When other ions lead to interferences, the addition of proper masking agents may be useful. Thus, for Be uptake by quinalizarine-loaded-C18-silica, interferences by other ions present in natural waters could be eliminated by addition of the following masking agents: 0.3 M EDTA for ZrO2þ, 0.16 M thiourea for Fe3þ and 0.05 M triethanolamine for Al3þ [128]. Ionic strength. The maximum sample volume that could be passed through a C18-silica cartridge loaded with ferrozine for the preconcentration of Fe(II) from sea water samples was found to be dependent on the ionic strength of the sample [122]. For sea water with salinities greater than 25‰, complete retention of iron was observed up to 500 ml. When salinities were in the range 2–25‰, the maximum volume was reduced to 200 ml. In addition, using a 60 ml sample, recovery was found to decrease when salinity was reduced (i.e. 91% recovery with 36‰ salinity, versus 80% at 5‰ salinity). Therefore, the addition of NaCl to sea water sample of low salinity would be advisable for quantitative recovery of iron. The sample ionic strength was also found crucial for the retention of beryllium on an OH-form strong anion-exchange resin [195]. Sorption decreased for ionic strengths over 0.2 due to elution of OH2 from the column. 14.3.2.3 Elution parameters Nature of the solvent The nature of the elution solvent is of prime importance and should optimally meet three criteria: efﬁciency, selectivity, and compatibility, as discussed below. In addition, it may be desirable to recover the analytes in a small volume of solvent to ensure a signiﬁcant enrichment factor. The eluent may be an organic solvent (when reversed-phase sorbents are used), an acid (usually with ionexchangers), or a complexing agent. Efﬁciency. The eluting solvent should be carefully chosen to ensure efﬁcient recovery of the retained target species. As an example, several solvents were tested for the elution of triphenyltin retained on a C18-silica cartridge [112]. No elution was achieved with aqueous solutions of the surfactant Triton X-100. ACN gave low recoveries, while tetrahydrofuran (THF) gave irreproducible results (probably due to the low stability of TPhT in THF solutions and its conversion to diphenyltin). Only 5% recovery was obtained with a methanol – water mixture (80:20). In fact, pure methanol was required to achieve acceptable recoveries (around 85%). In another study, several acids were tested for elution of Pb2þ retained on C18-silica discs modiﬁed with BAS [121]; acetic acid gave the best recovery. Indeed, acetate ion forms a highly stable complex with Pb2þ ion. The higher the acetic acid concentration used, the lower

437

V. Camel

the volume necessary for quantitative elution of the lead. Similar results were found with HBr for eluting Hg(II) from a modiﬁed C18-silica (with hexathia-18crown-6-tetraone) [130]. However, for elevated HBr concentrations (.1 M), leaching of the crown ether from the disc occurs. Selectivity. A further characteristic of the elution solvent arises with the possibility of introducing selectivity. The selectivity of C18-silica modiﬁed with Cyanex 301 towards Bi was evaluated [127] and it appears that Pb(II) and Cu(II) were also retained. However, washing the sorbent with 1 M HCl removed Pb(II), while Cu(II) was eluted using 1.5 M HCl. Subsequent elution of Bi(III) was ensured using 3 M HNO3. These results show that the complex formed between Cyanex 301 and bismuth had the highest stability constant, followed by copper and lead. As another example of selectivity during the elution step, 1 M HCOOH removed only Se(IV) from an anion-exchange resin, leaving Se(VI) retained on the sorbent [19]. The latter species could be eluted using 2 M HCl. Compatibility. The elution solvent should also be compatible with the analysis technique, as exempliﬁed by the recovery of a Cr(III) –DPCO complex from C18-silica, wherein the nature of the elution solvent was found crucial for the following chromatographic analysis [204]. With ACN, weak and wide peaks were obtained, possibly due to the low stability of the complex in this solvent. By contrast, sharp and symmetric peaks were obtained when methanol was used, due to the high stability of the complex in this solvent because of hydrogen bonding. Although several acids (HNO3, H2SO4, HCl) were found efﬁcient for eluting Be from quinalizarine-loaded-C18-silica, HNO3 was selected as nitrate ion is a more acceptable matrix for both ﬂame and electrothermal AAS [128]. Solvent pH As retention of trace elements on solid sorbents is usually pH dependent, careful choice of the elution solvent pH may enhance selectivity in the SPE procedure. As an example, once retained on ERT-functionalised silica gel, Mg(II) could be eluted ﬁrst at a pH around 4, while increasing the pH to 5–6 was required for eluting Zn(II) [9]. Elution mode Most of the time, for practical reasons, sample loading and elution steps are performed in a similar manner. However, to avoid irreversible adsorption and ensure quantitative recoveries, elution in the backﬂush mode is recommended in some cases. This means that the eluent is pumped through the sorbent in the opposite direction to that of the sample during the preconcentration step. This is especially crucial when carbon-based sorbents have to be used due to possible irreversible adsorption of the analytes.

438

Solid-phase extraction

Solvent ﬂow-rate The elution solvent ﬂow-rate should be high enough to avoid excessive duration, and low enough to ensure quantitative recovery of the target species. As an example, quantitative elution of iron ions from C18-silica discs was achieved using a ﬂow-rate in the range of 1.0 –3.5 ml/min [117]. Elution of Cu(II) from 11hydroxynaphthacene-5,12-quinone modiﬁed C18-silica discs was achieved in a ﬂow-rate range of 0.5 –5 ml/min using 3 ml of 0.1 M HNO3 [119]. At higher ﬂowrates, larger volumes of nitric acid solution were needed. Similar results were obtained in the case of lead retained on C18-silica modiﬁed with BAS when eluted with acetic acid [121], as well as of Hg(II) retained on crown-ether-modiﬁed C18silica discs for elution with HBr [130]. In the case of salen I modiﬁed C18-silica discs, Cu2þ elution could be achieved using ﬂow-rates ranging from 1 to 20 ml/ min [34]. Flow-rates of 1–10 ml/min enabled the quantitative elution of Agþ from HT18C6-loaded C18-silica discs with 10–15 ml of 0.1 M S2O3 22 solution [129]. Higher ﬂow-rates required larger volumes for complete elution. Solvent volume Similar to breakthrough volume, the elution volume may be determined either experimentally or estimated theoretically [229]. Minimum elution volume for a cartridge is deﬁned as two bed volumes of elution solvent. Bed volume is typically 120 ml per 100 mg of sorbent. For classical discs, the minimum solvent volume required is approximately 10 ml/mg of sorbent [20]. Consequently, larger elution volumes would be required for discs, even though the use of micro-sized discs may allow reduced solvent volumes. The elution step should enable sufﬁcient time and sorbent volume to permit the metallic species to diffuse out of the solid sorbent pores. As an illustration, a volume of at least 8 ml of 90:10 (v/v) MeOH– water was required to completely elute Cu – bathocuproine complex from C18-silica discs [27]. Furthermore, two elution cycles were found to be better than a single step (e.g., a single 10 ml elution volume gave a 15% lower recovery compared to 2 £ 5 ml elutions). Soaking time was also critical and a 2 min soak was allowed before each elution. The elution volume can usually be reduced by increasing the concentration of the eluting solvent (e.g. acid). However, in this case, problems with subsequent analysis may be encountered (e.g. F-AAS). 14.4

APPLICATIONS OF SPE TO THE DETERMINATION OF SOME TRACE ELEMENTS

14.4.1 Chromium Most SPE methods are based on the high reactivity of Cr(VI), as indicated in Table 14.5a. Several FI on-line preconcentration procedures have been reviewed up to 1992 by Sperling et al. [71], and later (until 1998) by Prasada Rao et al. [59]. Determination of chromium is frequently achieved by

439

440

TABLE 14.5A Applications of SPE to chromium Matrix

Reagent

Sorbent

Eluent

Operation

Analysis method

LOD (ng/l)

Ref.

Cr(VI)

Geological samples

DPC

Polyethylene

MeOH

Off-line SPE

LC-UV

2000

[204]

Tap water

DPC

XAD-16

H2SO4 – MeOH

Off-line SPE

F-AAS

45,000

[161]

Tap, mineral, rain, surface, and waste waters, soil and airborne particulate matter extracts

DPC

Cationexchanger

None

Off-line SPE

Spectrophotometry

10

[233]

Natural waters

DPC

Cationexchanger

HNO3 – acetone

FI system

Spectrophotometry

8.9– 15.2

[83]

Wastewater

None

Anionexchanger (SAX)

Na2SO4

Off-line SPE

Spectrophotometry after reaction DPC

8000

[194]

Sea, river waters

DDTC

C18-silica

EtOH

FI system

ET-AAS

1600

[60]

Tap, river, coastal, industrial waste waters

APDC

PTFE turnings

IBMK

FI system

F-AAS

800

[49]

Drinking, sea waters

APDC

PTFE (KR)

EtOH

FI system

ET-AAS

4.2

[74]

V. Camel

Metal species

TABLE 14.5A (continuation) Metal species

Cr(VI)/ Cr(III)

Reagent

Sorbent

Eluent

Operation

Analysis method

LOD (ng/l)

Ref.

Certiﬁed natural water, sea water

APDC

PTFE (KR)

EtOH

FI system

ET-AAS

16

[75]

Certiﬁed natural water, sea water

APDC

PTFE beads

EtOH

FI system

ET-AAS

8.8

[75]

Waters

None

Acidic Al2O3

NHþ 4

FI system

ICP-OES

200

[70]

Urine

None

Basic Al2O3

HNO3

FI system

ICP-OES

50

[73]

River, mineral, and tap waters

None

PAPhA

HCl

FI system

F-AAS

200

[81]

Tape, lake, waste waters

None

Cellulose

HCl

Off-line SPE

ET-AAS

1.8

[183]

Sea water

8-HQ

PS –DVB

HCl – MeOH

Off-line SPE

ET-AAS

–

[158]

River and reservoir waters

None

8-HQfunctionalisedpolyacrylonitrile

HCl – HNO3

Off-line SPE

ICP-MS

60

[186]

Sea water, industrial efﬂuents

DDTC

C18-silica

MeOH

FI system

F-AAS

20

[59]

Water

APDC

C18-silica

H2O– ACN

On-line SPE

LC-UV

60– 200

[39]

Lake, river, tap waters

None

Acidic Al2O3

HNO3 or NHþ 4

FI system

F-AAS

800 – 1000

[71]

441

continued

Solid-phase extraction

Cr(III)

Matrix

442

TABLE 14.5A (continuation) Metal species

Reagent

Sorbent

Eluent

Operation

Analysis method

LOD (ng/l)

Ref.

Synthetic aqueous solutions

None

Acidic Al2O3

HNO3 or NHþ 4

On-line SPE

ICP-OES

–

[234]

Tap, ground waters

None

Neutral Al2O3

HNO3 or NHþ 4

Off-line SPE

ET-AAS

10

[110]

Sewage waters

None

Al2O3

HNO3 or NHþ 4

FI system

F-AAS

42,000 – 81,000

[72]

Rain, river, sea, tap waters

None

TiO2

HNO3

Off-line SPE

ET-AAS

24–30

[108]

Waste waters

None

NDSAfunctionalisedPS– DVB

HCl

Off-line SPE

F-AAS

–

[181]

River water

None

Aminocarboxylicfunctionalised-PS

HCl

Off-line SPE

ET-AAS

30

[235]

Industrial wastewater

EDTA

Anion-exchanger (SAX)

NaCl

Off-line SPE

AAS

400 –1100

[236]

V. Camel

Matrix

Solid-phase extraction

spectrophotometry after derivatisation with a reagent such as DPC, which is very selective for Cr(VI). The chromate oxidises DPC to DPCO to form a soluble strongly red-violet compound with Cr(III). This complex can be retained on polyethylene [204], Amberlite XAD-16 [161], or on a cation-exchanger [83,233]. Cr(VI) may also be chelated by DDTC and subsequently retained on C18-silica [60], or by APDC with further retention on PTFE [49,74,75]. Acidic alumina enables selective retention of Cr(VI) [70], even though it was later reported that between pH 3 and 6 Cr(III) could also be retained [234]. In all these methods, determination of total chromium may be achieved after preliminary oxidation of Cr(III). With correct adjustment of sample pH, selective retention of Cr(III) may occur on some sorbents such as basic alumina [73], cellulose [183], or poly(aminophosphonic acid) (PaPhA) resin [81]. Complexation of Cr(III) with 8HQ may also be performed [158,186]. In that case, determination of total chromium is achieved after preliminary reduction of Cr(VI). A few studies report chromium speciation based on complex formation with APDC [39] or DDTC [59], and subsequent retention on C18-silica. Inorganic oxides have also been used, such as acidic Al2O3 [71,234] or TiO2 [108], with correct adjustment of the sample pH to achieve selective retention of either Cr(III) or Cr(VI). This latter step is avoided with neutral alumina Al2O3, speciation being obtained upon selective elution of the retained species [110]. Some polymeric sorbents have also been reported to enable selective retention of chromium species upon sample pH adjustment [181,235,236]. 14.4.2 Iron Several methods have been proposed for the determination of Fe(III) and Fe(II) species. Examples are given in Table 14.5b. Solid phases have been synthesised with high selectivity towards Fe(III), such as silica gel functionalised with formylsalicylic acid [35] or purpurogallin [140]. Dissolved iron in sea water could be determined by preconcentrating both dissolved Fe(III) and Fe(II) on 8-HQfunctionalised silica gel [52]. Alternatively, trace amounts of iron can be converted into Fe(II), by adding hydroxylamine to the sample, which forms a highly stable coloured complex with bathophenanthroline, retained on C18-silica [117]. Similarly, ferrozine selectively forms a stable coloured complex with Fe(II) in a pH range of 4–10 [237]. This complex, along with the excess reagent, could be retained on C18-silica [123]. Separation of both eluted species was subsequently performed by LC. Alternatively, Fe(II) could be preconcentrated on ferrozine-loaded C18-silica [68,122]. Fe(III) determination was also possible after its initial reduction with ascorbic acid. However, this can cause a shift in the Fe(II)/Fe(III) equilibrium in the solution. Consequently, a procedure has been reported that allows simultaneous determination of labile Fe(II) and Fe(III). Addition of 1,10-phenanthroline and ferron enables selective complexation of Fe(II) and Fe(III), respectively, followed by retention of these complexes on selective solid sorbents [18].

443

444

TABLE 14.5B Applications of SPE to iron Matrix

Reagent

Sorbent

Eluent

Operation

Analysis method

LOD (ng/l)

Ref.

Fe(II)

Aerosols, rain water, sea water

FZ

C18-silica

MeOH

Off-line SPE

LC-UV

0.1 nmol/l

[123]

Sea waters

None

FZ-loaded -C18-silica

MeOH

Off-line SPE

Spectrophotometry

0.6 nmol/l

[122]

Sea water

None

FZ-loaded -C18-silica

MeOH

FI system

Spectrophotometry

0.1 – 0.3 nM

[68]

Sea waters

None

8-HQfunctionalised -SiO2

HCl

FI system

Spectrophotometry

0.016 nM

[52]

Tap waters, well water (þ NH2OH)

Bathophenanthroline

C18-silica

EtOH þ NaClO4

Off-line SPE

Spectrophotometry

80

[117]

Wines

Phenanthroline þ ferron

Anion-exchanger þ C18-silica

HCl þ MeOH –HCl

Off-line SPE

F-AAS

–

[18]

Beverages, water (þ ascorbic acid)

None

TPTZ –TPB –naphthalene

DMF dissolution

Off-line SPE

Spectrophotometry

–

[227]

Fe(III) /Fe(II)

V. Camel

Metal species

Solid-phase extraction

14.4.3 Mercury Inorganic mercury Hg(II) can be retained on silica gel functionalised with dithizone [104], dithioacetal derivatives [105] or iminodithiocarbamates [106]. New chelating sorbents have also been synthesised for the preconcentration of inorganic mercury, such as a resin containing histidine covalently bound to its carboxyl groups [238], or a PS–DVB-based resin with picolinic acid amide as the functional groups [184]. Alternatively, impregnation of C18-silica with selective ligands may be efﬁcient, such a hexathia-18-crown-6-tetraone [130]. For the preconcentration of both inorganic mercury and organic mercuric species (e.g. methylmercury, ethylmercury), chelating resins containing dithiocarbamate groups have been widely used [189,239,240]. Another study reported the use of dithizone-loaded C18-silica [31]. Chelation may also be performed directly in the sample upon addition of the reagent, and further retention of the chelates on C18-silica. Among reagents, APDC [43,55], DDTP [64], DDTC [55], and dithizone (DZ) [55] were found efﬁcient (see Table 14.5c). 14.4.4 Selenium An overview of SPE procedures developed for selenium has been recently given [193]. Most methods were dedicated to the determination of inorganic selenium. Some of them use selective reagents for complexation of Se(IV), such as APDC [114] or Bismuthiol II [222]. The use of chelating resins has also been reported, such as Chelex-100 [241]. A reduction of Se(VI) to Se(IV) prior to chelation enables the determination of total inorganic Se. Other methods involve ion-exchangers for the retention of anionic Se(IV) and Se(VI) [40,225, 242 –244], or acidic alumina [111] (see Table 14.5d). Speciation can be obtained further by selective elution of both inorganic species or by LC separation. Alternatively, ion-pairs can be formed in the sample with inorganic selenium, and then retained on C18-silica [40]. Organoselenium species can be selectively retained on Amberlite XAD-2 [244]. However, a partial retention of Se(IV) on that sorbent has been noted [225]. Simultaneous preconcentration of inorganic and organic selenium species is a more difﬁcult task. An anion-exchanger cartridge placed on the top of a C18-silica cartridge retained inorganic selenium, while organic species were retained on the reversed-phase sorbent [19]. Determination of organic and inorganic selenium was also achieved using preconcentration on a mixedmode sorbent (i.e. anionic and chelating resin), Amberlite IRA-743, followed by separation of the species by LC [193]. 14.4.5 Tin Several studies report the use of SPE for organotin determination as already overviewed [245 –247]. Examples are given in Table 14.5e. C18-silica has been

445

446

TABLE 14.5C Applications of SPE to mercury Matrix

Reagent

Sorbent

Eluent

Operation

Analysis method

LOD (ng/l)

Ref.

Hg(II)

Spiked tap water

None

DithizonefunctionalisedSiO2

HCl

Off-line SPE

CV-AAS

3960

[104]

Spiked tap and sea waters

None

DithioacetalsfunctionalisedSiO2

H 2O

Off-line SPE

CV-AAS

–

[105]

Spiked tap and sea waters

None

IminodithiocarbamatesfunctionalisedSiO2

H 2O

Off-line SPE

CV-AAS

–

[106]

River water

None

PAAMfunctionalisedPS –DVB

H2SO4

Off-line SPE

Spectrophotometry

–

[184]

Tap, river, well, and spring waters

None

HT18C6TOloadedC18-silica

HBr

Off-line SPE

CV-AAS

6

[130]

Fish, human urine

APDC

C18-silica

MeOH – ACN – water

FI system

LC –CV-AAS

5.5 –10.4

[43]

Sea water

DDTC

C18-silica

EtOH

FI system

CV-AAS

16

[55]

Hg(II)/ organic Hg

V. Camel

Metal species

TABLE 14.5C (continuation) Metal species

Reagent

Sorbent

Eluent

Operation

Analysis method

LOD (ng/l)

Ref.

Certiﬁed citrus leaves, marine sediment, dogﬁsh liver

DDTP

C18-silica

EtOH

FI system

CV-AAS

10

[64]

Synthetic sea waters

None

Dithizone -loadedC18-silica

MeOH

Off-line SPE

LC-DAD

140

[31]

River, lake, and rain waters

None

Dithiocarba matefunctionalisedpolyvinyl

Thiourea – HCl

Off-line SPE

CV-AAS

0.2

[189]

Sea and fresh waters

None

Dithiocarbamate resin

Thiourea –HCl

FI system

GC-MPD

0.05 –0.15

[240]

Solid-phase extraction

Matrix

447

448

TABLE 14.5D Applications of SPE to selenium Metal species

Matrix

Reagent

Sorbent

Eluent

Operation

Analysis method

LOD (ng/l)

Ref.

Se(IV)

Sea waters

APDC

C18-silica

MeOH

Off-line SPE

ET-AAS

7

[114]

Off-line SPE

ET-AAS

49 –800

[111]

Se(IV)/Se(VI)

None

Acidic Al2O3

Spiked tap and ground waters

None

Anionexchanger (Cellex T)

HNO3

Off-line SPE

ET-AAS

2400 –2.1 £ 106

[225]

Drinking and fresh waters

None

Anionexchanger (Dowex 1X8)

HCl

FI system

HG-AAS

5

[242]

Spiked river, drinking and well waters

None

Anionexchanger (PRP-X100)

NH4HCO3 – (NH4)2CO3

On-line SPE

LC – ICP-MS

8 –420

[40]

Ground and fresh waters

None

Amberlite IRA-743

HClO4

Off-line SPE

LC – ICP-MS

10

[193]

Sea, river, tap waters

None

Anionexchanger (SAX) þ C18-silica

HCOOH þ HCl þ CS2

Off-line SPE

GC –MS (after derivatisation)

0.6 – 903

[19]

V. Camel

Se(IV)/Se(VI)/ organic Se

Tap, ground waters

NHþ 4

TABLE 14.5E Applications of SPE to tin Matrix

Reagent

Sorbent

Eluent

Operation

Analysis method

LOD (ng/l)

Ref.

TBT

Sea water

None

C18-silica

Acidiﬁed ethyl acetate

Off-line SPE

GC-ECD

–

[24]

Fresh waters

None

Tropoloneloaded-XAD-2

IBMK

Off-line SPE

ET-AAS

14.4

[164]

Spiked sea waters

None

C18-silica

Hydroxyﬂavone in MeOH

Off-line SPE

Fluorimetry

70

[112]

Sea water

None

C18-silica

MeOH

Off-line SPE

Fluorimetry

–

[113]

TBT/TPhT

Sea water

None

GCB

MeOH –CH2Cl2

Off-line SPE

LC –ET-AAS

0.7 – 0.8

[223]

DiBT/TBT/ DiPhT/TPhT

Oyster and mussel extracts

None

Cationexchanger (SCX)

NH4Cl in MeOH –acetic acid –H2O

Off-line SPE

LC-FLD

3 –140

[36]

MonoBT/ DiBT/TBT/ MonoPhT/ DiPhT/TPhT

Rain, lake, river waters

None

Tropoloneloaded-C18silica

Diethyl ether

Off-line SPE

Ethylation – GC –FPD

–

[118]

River waters

None

C18-silica

MeOH

FI system

GC-AES

0.10 – 0.17

[247]

Harbour and river waters

NaBEt4

C18-silica

Supercritical CO2

Off-line SPE

GC –FPD

–

[248]

TPhT

MonoBT/DiBT/ TBT/MonoPhT/ DiPhT/TPhT/ DiCyT/TCyT

Solid-phase extraction

Metal species

449

V. Camel

found effective in preconcentrating several organotin compounds, mainly tributyltin (TBT) [24,247] and triphenyltin (TPhT) [112,113,247] from aqueous solutions, with possible storage of TBT for at least 1 month [24]. Derivatisation of the compounds may also be performed prior to the SPE step, and the ethylated organotins retained on C18-silica [248]. Tropolone-loaded C18-silica was efﬁcient for several organotins [118], as well as tropolone-loaded Amberlite XAD-2 [163], with possible selective retention of TBT on the latter sorbent upon addition of 0.8% sulfuric acid to the sample [164]. GCB may also be used for the preconcentration of organotin compounds [223]. These cationic species can also be efﬁciently retained on a strong cation-exchanger [36].

14.5

CONCLUSION

The use of SPE procedures has been growing in the past few years due to its advantages for trace element determination, namely conservation of species, good preconcentration factors (thus enabling the achievement of very low limits of detection), ease of automation, and possible on-line coupling to analysis techniques. Although the numerous steps and parameters used enable an efﬁcient extraction and recovery of the target analytes, the choice of the solid sorbent is the most critical step. Among the numerous sorbents that have been used, it clearly appears that the initial use of ion-exchangers is being replaced by more selective supports containing chelating functional groups. Such sorbents are frequently based on hydrophobic supports, namely, C18-silica or PS –DVB copolymer, the latter affording a broader pH range. A suitable way of proceeding is to load the reagent on the solid sorbent. The coated sorbent thus obtained may usually be used several times, but partial leaching of the chelating agent may occur over time during elution of the analytes, especially if the eluting solvent is too concentrated. Alternatively, the reagent may also be chemically bound to the sorbent, leading to the synthesis of new phases, and thus avoiding any leaching of the reagent. Promising results have also been recently obtained with inorganic oxides, such as titania and alumina, as retention of some trace elements could be achieved with the raw sample (i.e. neither reagent addition nor pH adjustment), thereby avoiding possible speciation changes in the sample. Examples given in this chapter for several trace elements show the high potential of SPE, especially its potentially high selectivity (by choosing the nature of the sorbent and/or the chelating agent, as well as the nature of the eluent). In fact, in some cases, differentiation of species may be achieved, thereby offering new opportunities for speciation. There is thus no doubt that this technique will face a growing interest for trace element determination and speciation in the future, as has already been seen for organic micropollutant determinations in recent years.

450

Solid-phase extraction

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35

I. Liska, J. Chromatogr. A, 885 (2000) 3. C.F. Poole, Encyclopedia of Separation Science, Vol. 3. Academic Press, London, (UK). 2000, p. 1405. M.C. Hennion, Trends Anal. Chem., 10 (1991) 317. J.L. Lundgren and A.A. Schilt, Anal. Chem., 49 (1977) 974. M. Chikuma, M. Nakayama, T. Itoh, H. Tanaka and K. Itoh, Talanta, 27 (1980) 807. H. Akaiwa, H. Kawamoto and K. Ogura, Talanta, 28 (1981) 337. K.S. Lee, W. Lee and D.W. Lee, Anal. Chem., 50 (1978) 255. K. Kilian and K. Pyrzynska, Fresenius J. Anal. Chem., 371 (2001) 1076. M.E. Mahmoud, Talanta, 45 (1997) 309. M. Pesavento, R. Biesuz, M. Gallorini and A. Profumo, Anal. Chem., 65 (1993) 2522. M. Pesavento, R. Biesuz and J.L. Cortina, Anal. Chim. Acta, 298 (1994) 225. H. Kumagai, Y. Inoue, T. Yokoyama, T.M. Suzuki and T. Suzuki, Anal. Chem., 70 (1998) 4070. J. Seneviratne and J.A. Cox, Talanta, 52 (2000) 801. M.C. Carson, J. Chromatogr. A, 885 (2000) 343. V. Porta, E. Mentasti, C. Sarzanini and M.C. Gennaro, Talanta, 35 (1988) 167. P. Janos, K. Stulik and V. Pacakova, Talanta, 39 (1992) 29. R. Van Grieken, Anal. Chim. Acta, 143 (1982) 3. A.B. Tawali and G. Schwedt, Fresenius J. Anal. Chem., 357 (1997) 50. J.L. Gomez-Ariza, J.A. Pozas, I. Giraldez and E. Morales, Analyst, 124 (1999) 75. E.M. Thurman and K. Snavely, Trends Anal. Chem., 19 (2000) 18. C.F. Poole, Encyclopedia of Separation Science, Vol. 9. Academic Press, London (UK). 2000, p. 4141. M. Kumar, D.P.S. Rathore and A.K. Singh, Fresenius J. Anal. Chem., 370 (2001) 377. G.A. Junk, M.J. Avery and J.J. Richard, Anal. Chem., 60 (1988) 1347. O. Evans, B.J. Jacobs and A.L. Cohen, Analyst, 116 (1991) 15. D.F. Hagen, C.G. Markell, G.A. Schmitt and D.D. Blevins, Anal. Chim. Acta, 236 (1990) 157. D.T. Rossi and N. Zhang, J. Chromatogr. A, 885 (2000) 97. L.B. Bjo¨rklund and G.M. Morrison, Anal. Chim. Acta, 343 (1997) 259. R.E. Majors, LC–GC, 4 (1989) 272. R.E. Sturgeon, S.S. Berman, A. Desaulniers and D.S. Russell, Talanta, 27 (1980) 85. T.M. Florence and G.E. Batley, Talanta, 23 (1976) 179. D.M. Sanchez, R. Martin, R. Morante, J. Marin and M.L. Munuera, Talanta, 52 (2000) 671. M.C. Yebra, N. Carro, M.F. Enriquez, A. Moreno-Cid and A. Garcia, Analyst, 126 (2001) 933. E. Castillo, J.-L. Cortina, J.-L. Beltran, M.-D. Prat and M. Granados, Analyst, 126 (2001) 1149. M. Shamsipur, A.R. Ghiasvand, H. Sharghi and H. Naeimi, Anal. Chim. Acta, 408 (2000) 271. M.E. Mahmoud and E.M. Soliman, Talanta, 44 (1997) 15.

451

V. Camel 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71

452

E. Gonzalez-Toledo, M. Benzi, R. Compano, M. Granados and M.D. Prat, Anal. Chim. Acta, 443 (2001) 183. O. Keil, J. Dahmen and D.A. Volmer, Fresenius J. Anal. Chem., 364 (1999) 694. J.F. Tyson, Spectrochim. Acta Rev., 14 (1991) 169. M. Bittner and J.A.C. Broekaert, Anal. Chim. Acta, 364 (1998) 31. Q. Hu, G. Yang, J. Yin and Y. Yao, Talanta, 57 (2002) 751. Y. Cai, M. Cabanas, J.L. Fernandez-Turiel, M. Abalos and J.M. Bayona, Anal. Chim. Acta, 314 (1995) 183. B.W. Wenclawiak, T. Hees, C.E. Zo¨ller and H.-P. Kabus, Fresenius J. Anal. Chem., 358 (1997) 471. X. Yin, W. Frech, E. Hoffmann, C. Lu¨dke and J. Skole, Fresenius J. Anal. Chem., 361 (1998) 761. S. Olsen, L.C.R. Pessenda, J. Ruzicka and E.H. Hansen, Analyst, 108 (1983) 905. Z. Fang, J. Ruzicka and E.H. Hansen, Anal. Chim. Acta, 164 (1984) 23. Z. Fang, Z. Zhu, S. Zhang, S. Xu, L. Guo and L. Sun, Anal. Chim. Acta, 214 (1988) 41. Z. Fang, M. Sperling and B. Welz, J. Anal. At. Spectrom., 5 (1990) 639. A.N. Anthemidis, G.A. Zachariadis and J.A. Stratis, Talanta, 54 (2001) 935. A.N. Anthemidis, G.A. Zachariadis, J.-S. Kougoulis and J.A. Stratis, Talanta, 57 (2002) 15. G.A. Zachariadis, A.N. Anthemidis, P.G. Bettas and J.A. Stratis, Talanta, 57 (2002) 919. M. Zougagh, A. Garcia de Torres and J.M. Cano Pavon, Talanta, 56 (2002) 753. D.A. Weeks and K.W. Bruland, Anal. Chim. Acta, 453 (2002) 21. Q. Pu, Q. Sun, Z. Hu and S. Su, Analyst, 123 (1998) 239. S. Zhang, Q. Pu, P. Liu, Q. Sun and Z. Su, Anal. Chim. Acta, 452 (2002) 223. M. Fernandez Garcia, R. Pereiro Garcia, N. Bordel Garcia and A. Sanz-Medel, Talanta, 41 (1994) 1833. Z.-S. Liu and S.-D. Huang, Anal. Chim. Acta, 281 (1993) 185. A. Ali, X. Yin, H. Shen, Y. Ye and X. Gu, Anal. Chim. Acta, 392 (1999) 283. P.-G. Su and S.-D. Huang, Anal. Chim. Acta, 376 (1998) 305. T. Prasada Rao, S. Karthikeyan, B. Vijayalekshmy and C.S.P. Iyer, Anal. Chim. Acta, 369 (1998) 69. M. Sperling, X. Yin and B. Welz, Analyst, 117 (1992) 629. Y. Ye, A. Ali and X. Yin, Talanta, 57 (2002) 945. R. Lima, K.C. Leandro and R.E. Santelli, Talanta, 43 (1996) 977. S. Quinaia, J.B.B. Da Silva, M.C.E. Rollemberg and A.J. Curtius, Talanta, 54 (2001) 687. A.C.P. Monteiro, L.S.N. Andrade and R.C. Campos, Fresenius J. Anal. Chem., 371 (2001) 353. J. Ruzicka and A. Arndal, Anal. Chim. Acta, 216 (1989) 243. Z.F. Queiroz, F.R.P. Rocha, G. Knapp and F.J. Krug, Anal. Chim. Acta, 463 (2002) 275. L.S.G. Teixeira, F.R.P. Rocha, M. Korn, B.F. Reis, S.L.C. Ferreira and A.C.S. Costa, Anal. Chim. Acta, 383 (1999) 309. S. Blain and P. Tre´guer, Anal. Chim. Acta, 308 (1995) 425. E. Vassileva and N. Furuta, Fresenius J. Anal. Chem., 370 (2001) 52. A.G. Cox, I.G. Cook and C.W. McLeod, Analyst, 110 (1985) 331. M. Sperling, S. Xu and B. Welz, Anal. Chem., 64 (1992) 3101.

Solid-phase extraction 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107

108

M.J. Marques, A. Morales-Rubio, A. Salvador and M. de la Guardia, Talanta, 53 (2001) 1229. A.G. Cox and C.W. McLeod, Anal. Chim. Acta, 179 (1986) 487. S. Nielsen and E.H. Hansen, Anal. Chim. Acta, 366 (1998) 163. W. Som-Aum, S. Liawruangrath and E.H. Hansen, Anal. Chim. Acta, 463 (2002) 99. C. Shuyu, Z. Zhifeng and Y. Huaming, Anal. Chim. Acta, 451 (2002) 305. Z. Fang, S. Xu, L. Dong and W. Li, Talanta, 41 (1994) 2165. R. Compano, R. Ferrer, J. Guiteras and M.D. Prat, Analyst, 119 (1994) 1225. I.A. Kovalev, L.V. Bogacheva, G.I. Tsysin, A.A. Formanovsky and Y.A. Zolotov, Talanta, 52 (2000) 39. P. Hashemi and A. Olin, Talanta, 44 (1997) 1037. R.M. Cespon-Romero, M.C. Yebra-Biurrun and M.P. Bermejo-Barrera, Anal. Chim. Acta, 327 (1996) 37. N.G. Beck, R.P. Francks and K.W. Bruland, Anal. Chim. Acta, 455 (2002) 11. S. Matsuoka, Y. Tennichi, K. Takehara and K. Yoshimura, Analyst, 124 (1999) 787. C.E.S. Miranda, B.F. Reis, N. Baccan, A.P. Packer and M.F. Gine´, Anal. Chim. Acta, 453 (2002) 301. K. Yoshimura, S. Matsuoka, Y. Inokura and U. Hase, Anal. Chim. Acta, 268 (1992) 225. J.A. Gomes Neto, A.P. Oliveira, G.P.G. Feshi, C.S. Dakuzaku and M. Moraes, Talanta, 53 (2000) 497. J.B.B. Silva, M.B.O. Giacomelli and A.J. Curtius, Analyst, 124 (1999) 1249. R.E. Santelli, M. Gallego and M. Valcarcel, Talanta, 41 (1994) 817. H. Zhang, X. Yuan, X. Zhao and Q. Jin, Talanta, 44 (1997) 1615. S. Lin, C. Zheng and G. Yun, Talanta, 42 (1995) 921. S.K. Xu, M. Sperling and B. Welz, Fresenius J. Anal. Chem., 344 (1992) 535. C.S.L. Ferreira, D.S. Jesus, R.J. Cassella, A.C.S. Costa, M.S. Carvalho and R.E. Santelli, Anal. Chim. Acta, 378 (1999) 287. S. Tsakovski, K. Benkhedda, E. Ivanova and F.C. Adams, Anal. Chim. Acta, 453 (2002) 143. S.D. Hartenstein, J. Ruzicka and G.D. Christian, Anal. Chem., 57 (1985) 21. K. Benkhedda, H.G. Infante, F.C. Adams and E. Ivanova, Trends Anal. Chem., 21 (2002) 332. J.N. King and J.S. Fritz, Anal. Chem., 57 (1985) 1016. C. Kantipuly, S. Katragadda, A. Chow and H.D. Gesser, Talanta, 37 (1990) 491. C. Gueguen, J. Dominck and D. Perret, Fresenius J. Anal. Chem., 370 (2001) 909. O. Abollino, M. Aceto, M.C. Bruzzoniti, E. Mentasti and C. Sarzanini, Anal. Chim. Acta, 375 (1998) 299. A. Tong, Y. Akama and S. Tanak, Analyst, 115 (1990) 947. K. Terada, K. Matsumoto and Y. Taniguchi, Anal. Chim. Acta, 147 (1983) 411. K. Terada, K. Matsumoto and T. Inaba, Anal. Chim. Acta, 158 (1984) 207. K. Terada and K. Nakamura, Talanta, 28 (1981) 123. M.E. Mahmoud, M.M. Osman and M.E. Amer, Anal. Chim. Acta, 415 (2000) 33. M.E. Mahmoud and G.A. Gohar, Talanta, 51 (2000) 77. M.E. Mahmoud, Anal. Chim. Acta, 398 (1999) 297. Y. Sohrin, S.-I. Iwamoto, S. Akiyama, T. Fujita, T. Kugii, H. Obata, E. Nakayama, S. Goda, Y. Fujishima, H. Hasegawa, K. Ueda and M. Matsui, Anal. Chim. Acta, 363 (1998) 11. E. Vassileva, K. Hadjiinov, T. Stoychev and C. Daiev, Analyst, 125 (2000) 693.

453

V. Camel 109 110 111 112 113 114 115 116 117 118 119 120 121 122 123 124 125 126 127 128 129 130 131 132 133 134 135 136 137 138 139 140 141 142 143 144 145

454

E. Vassileva, I. Proinova and K. Hadjiivanov, Analyst, 121 (1996) 607. A.C. Sahayam, Anal. Bioanal. Chem., 372 (2002) 840. K. Pyrzynska, P. Drzewicz and M. Trojanowicz, Anal. Chim. Acta, 363 (1998) 141. R. Compano, M. Granados, C. Leal and M.D. Prat, Anal. Chim. Acta, 283 (1993) 272. J. Saurina, C. Leal, R. Compano, M. Granados, R. Tauler and M.D. Prat, Anal. Chim. Acta, 409 (2000) 237. R.E. Sturgeon, S.N. Willie and S.S. Berman, Anal. Chem., 57 (1985) 6. R.E. Sturgeon, S.S. Berman and S.N. Willie, Talanta, 29 (1982) 167. Y. Yamini and A. Tamaddon, Talanta, 49 (1999) 119. Y. Yamini and N. Amiri, J. AOAC Int., 84 (2001) 713. M.D. Mu¨ller, Anal. Chem., 59 (1987) 617. M. Shamsipur, A. Avanes, M.K. Rofouei, H. Sharghi and G. Aghapour, Talanta, 54 (2001) 863. O.R. Hashemi, M.R. Kargar, F. Raouﬁ, A. Moghimi, H. Aghabozorg and M.R. Ganjali, Microchem. J., 69 (2001) 1. M. Shamsipur, F. Raouﬁ and H. Sharghi, Talanta, 52 (2000) 637. D.W. King, J. Lin and D.R. Kester, Anal. Chim. Acta, 247 (1991) 125. Z. Yi, G. Zhuang, P.R. Brown and R.A. Duce, Anal. Chem., 64 (1992) 2826. Z.-S. Liu and S.-D. Huang, Anal. Chim. Acta, 267 (1992) 31. M. Shamsipur, A.R. Ghiasvand and Y. Yamini, Anal. Chem., 71 (1999) 4892. M.B. Shabani, T. Akagi and A. Masuda, Anal. Chem., 64 (1992) 737. Y. Yamini, M. Chaloosi and H. Ebrahimzadeh, Talanta, 56 (2002) 797. Y. Yamini, J. Hassan, R. Mohandesi and N. Bahramifar, Talanta, 56 (2002) 375. M. Shamsipur and M.H. Mashhadizadeh, Fresenius J. Anal. Chem., 367 (2000) 246. Y. Yamini, N. Alizadeh and M. Shamsipur, Anal. Chim. Acta, 355 (1997) 69. R.J. Kvitek, J.F. Evans and P.W. Carr, Anal. Chim. Acta, 144 (1982) 93. A.R. Sarkar, P.K. Datta and M. Sarkar, Talanta, 43 (1996) 1857. K. Terada, K. Matsumoto and H. Kimura, Anal. Chim. Acta, 153 (1983) 237. R.E. Sturgeon, S.S. Berman, S.N. Willie and J.A.H. Desaulniers, Anal. Chem., 53 (1981) 2337. J.W. McLaren, A.P. Mykytiuk, S.N. Willie and S.S. Berman, Anal. Chem., 57 (1985) 2907. R. Kocjan and S. Przeszlakowski, Talanta, 39 (1992) 63. D.E. Leyden, G.H. Luttrell, A.E. Sloan and N.J. DeAngelis, Anal. Chim. Acta, 84 (1976) 97. T. Seshadri and H.-J. Haupt, Anal. Chem., 60 (1988) 47. P. Jones, P.J. Hobbs and L. Ebdon, Anal. Chim. Acta, 149 (1983) 39. M.E. Mahmoud and M.S.M. Al Saadi, Anal. Chim. Acta, 450 (2001) 239. K.S. Abou-El-Sherbini, I.M.M. Kenawy, M.A. Hamed, R.M. Issa and R. Elmorsi, Talanta, 58 (2002) 289. S. Hutchinson, G.A. Kearney, E. Horne, B. Lynch, J.D. Glennon, M.A. McKervey and S.J. Harris, Anal. Chim. Acta, 291 (1994) 269. R. Garcia-Valls, A. Hrdlicka, J. Perutka, J. Havel, N.V. Deorkard, L.L. Tavlarides, M. Munoz and M. Valiente, Anal. Chim. Acta, 439 (2001) 247. P. Lessi, N.L. Dias Filho, J.C. Moreira and J.T.S. Campos, Anal. Chim. Acta, 327 (1996) 183. R.M. Izatt, R.L. Bruening, M.L. Bruening, B.J. Tarbet, K.E. Krakowiak, J.S. Bradshaw and J.J. Christensen, Anal. Chem., 60 (1988) 1825.

Solid-phase extraction 146 147 148 149 150 151 152 153 154 155 156 157 158 159 160 161 162 163 164 165 166 167 168 169 170 171 172 173 174 175 176 177 178 179 180 181 182 183 184 185

M.L. Bruening, D.M. Mitchell, J.S. Bradshaw, R.M. Izatt and R.L. Bruening, Anal. Chem., 63 (1991) 21. G. Philippeit and J. Angerer, J. Chromatogr. B, 760 (2001) 237. V. Cuculic, M. Mlakar and M. Branica, Anal. Chim. Acta, 339 (1997) 181. M.H. Pournaghi-Azar, Dj. Djozan and H. Abdolmohammad Zadeh, Anal. Chim. Acta, 437 (2001) 217. S.J. Kumar, P. Ostapczuk and H. Emons, At. Spectrosc., 20 (1999) 194. J.L. Manzoori, M.H. Sorouraddin and F. Shemirani, Talanta, 42 (1995) 1151. E. Vassileva and K. Hadjiinov, Fresenius J. Anal. Chem., 357 (1997) 881. P. Liang, Y. Qin, B. Hu, C. Li, T. Peng and Z. Jiang, Fresenius J. Anal. Chem., 368 (2000) 638. C.W. Huck and G.K. Bonn, J. Chromatogr. A, 885 (2000) 51. N. Masque´, R.M. Marce´ and F. Borrull, Trends Anal. Chem., 17 (1998) 384. O. Abollino, M. Aceto, M.C. Bruzzoniti, E. Mentasti and C. Sarzanini, Anal. Chim. Acta, 375 (1998) 293. S. Sarac¸oglu and L. Elc¸i, Anal. Chim. Acta, 452 (2002) 77. K. Isshiki, Y. Sohrin, H. Karatani and E. Nakayama, Anal. Chim. Acta, 224 (1989) 55. O. Abollino, E. Mentasti, V. Porta and C. Sarzanini, Anal. Chem., 62 (1990) 21. L. Elci, M. Soylak, A. Uzun, E. Bu¨yu¨kpatir and M. Dogan, Fresenius J. Anal. Chem., 368 (2000) 358. A. Tunc¸eli and A.R. Tu¨rker, Talanta, 57 (2002) 1199. A.G. Howard and M.H. Arbab-Zavar, Talanta, 26 (1979) 895. P. Bermejo-Barrera, G. Gonzalez-Campos, M. Ferron-Novais and A. BermejoBarrera, Talanta, 46 (1998) 1479. P. Bermejo-Barrera, R.M. Anllo-Sendin, M.J. Cantelar-Barbazan and A. BermejoBarrera, Anal. Bioanal. Chem., 372 (2002) 837. C.S.L. Ferreira, C.F. de Brito, A.F. Dantas, N.M. Lopo de Araujo and A.C. Spinola Costa, Talanta, 48 (1999) 1173. K. Isshiki, F. Tsuji, T. Kuwamoto and E. Nakayama, Anal. Chem., 59 (1987) 2491. A. Ramesh, K.R. Mohan and K. Seshaiah, Talanta, 57 (2002) 243. A.N. Masi and R.A. Olsina, Fresenius J. Anal. Chem., 357 (1997) 65. M.E. Leon-Gonzalez and L.V. Perez-Arribas, J. Chromatogr. A, 902 (2000) 3. R. Saxena, A.K. Singh and S.S. Sambi, Anal. Chim. Acta, 295 (1994) 199. R. Saxena, A.K. Singh and D.P.S. Rathore, Analyst, 120 (1995) 403. P.K. Tewari and A.K. Singh, Analyst, 125 (2000) 2350. R. Saxena and A.K. Singh, Anal. Chim. Acta, 340 (1997) 285. P.K. Tewari and A.K. Singh, Analyst, 124 (1999) 1847. P.K. Tewari and A.K. Singh, Talanta, 53 (2001) 823. P.K. Tewari and A.K. Singh, Talanta, 56 (2002) 735. M. Kumar, D.P.S. Rathore and A.K. Singh, Analyst, 125 (2000) 1221. K. Dev and G.N. Rao, Talanta, 42 (1995) 591. M.C. Yebra-Biurrun, M.C. Garcia-Dopazo, A. Bermejo-Barrera and M.P. BermejoBarrera, Talanta, 39 (1992) 671. N. Uehara, A. Katamine and Y. Shijo, Analyst, 119 (1994) 1333. B.C. Mondal, D. Das and A.K. Das, Talanta, 56 (2002) 145. Y. Cai, G. Jiang and J. Liu, Talanta, 57 (2002) 1173. F. Shemirani and M. Rajabi, Fresenius J. Anal. Chem., 371 (2001) 1037. B. Sengupta and J. Das, Anal. Chim. Acta, 219 (1989) 339. P.K. Tewari and A.K. Singh, Fresenius J. Anal. Chem., 367 (2000) 562.

455

V. Camel 186 187 188 189 190 191 192 193 194 195 196 197 198 199 200 201 202 203 204 205 206 207 208 209 210 211 212 213 214 215 216 217 218 219 220 221 222 223 224

456

B. Wen, X.-Q. Shuan and J. Lian, Talanta, 56 (2002) 681. B. Gong and Y. Wang, Anal. Bioanal. Chem., 372 (2002) 597. B. Wen, X.-Q. Shuan, R.-X. Liu and H.-X. Tang, Fresenius J. Anal. Chem., 363 (1999) 251. K. Minagawa, Y. Takizawa and I. Kifune, Anal. Chim. Acta, 115 (1980) 103. S. Blain, P. Appriou and H. Handel, Anal. Chim. Acta, 272 (1993) 91. S.-C. Pai, P.-Y. Whung and R.-L. Lai, Anal. Chim. Acta, 211 (1988) 257. M.E. Malla, M.B. Alvarez and D.A. Batistoni, Talanta, 57 (2002) 277. M. Bueno and M. Potin-Gautier, J. Chromatogr. A, 963 (2002) 185. M. Llobat-Estelles, A.R. Mauri-Aucejo and M.D. Lopez-Catalan, Fresenius J. Anal. Chem., 371 (2001) 358. A. Afkhami, T. Madrakian, A.A. Assl and A.A. Sehhat, Anal. Chim. Acta, 437 (2001) 17. S. Hoshi, H. Fujisawa, K. Nakamura, S. Nakata, M. Uto and K. Akatsuka, Talanta, 41 (1994) 503. B. Salih, R. Say, A. Denizli, O. Genc and E. Piskin, Anal. Chim. Acta, 371 (1998) 177. D.W. Lee and M. Halmann, Anal. Chem., 48 (1976) 2214. T. Braun and A.B. Farag, Anal. Chim. Acta, 76 (1975) 107. A. Alexandrova and S. Arpadjan, Analyst, 118 (1993) 1309. D.S. Jesus, R.J. Cassella, S.L.C. Ferreira, A.C.S. Costa, M.S. Carvalho and R.E. Santelli, Anal. Chim. Acta, 366 (1998) 263. C.S.L. Ferreira, H.C. Santos and D.S. Jesus, Fresenius J. Anal. Chem., 369 (2001) 187. I.A. Veselova and T.N. Shekhovtsova, Anal. Chim. Acta, 413 (2000) 95. S. Tian and G. Schwedt, Fresenius J. Anal. Chem., 354 (1996) 447. I.E. De Vito, R.A. Olsina and A.N. Masi, Fresenius J. Anal. Chem., 368 (2000) 392. J.P. Riley and D. Taylor, Anal. Chim. Acta, 40 (1968) 479. R. Boniforti, R. Ferraroli, P. Frigieri, D. Heltai and G. Queirazza, Anal. Chim. Acta, 162 (1984) 33. M. Pesavento and R. Biesuz, Anal. Chim. Acta, 346 (1997) 381. S.-C. Pai, Anal. Chim. Acta, 211 (1988) 271. C. Gueguen, C. Belin, B.A. Thomas, F. Monna, P.-Y. Favarger and J. Dominik, Anal. Chim. Acta, 386 (1999) 155. H. Kumagai, T. Yokoyama, T.M. Suzuki and T. Suzuki, Analyst, 124 (1999) 1595. B. Wen, X.-Q. Shuan and S.-G. Xu, Analyst, 124 (1999) 621. M.R. Buchmeiser, R. Tessadri, G. Seeber and G.K. Bonn, Anal. Chem., 70 (1998) 2130. F. Sinner, M.R. Buchmeiser, R. Tessadri, M. Mupa, K. Wurst and G.K. Bonn, J. Am. Chem. Soc., 120 (1998) 2790. D.J. Hutchinson and A.A. Schilt, Anal. Chim. Acta, 154 (1983) 159. A. Afkhami and T. Madrakian, Talanta, 58 (2002) 311. E. Piperaki, H. Berndt and E. Jackwerth, Anal. Chim. Acta, 100 (1978) 589. Y. Petit de Pena, M. Gallego and M. Varcarcel, J. Anal. At. Spectrom., 9 (1994) 691. T. Aydemir and S. Gucer, Anal. Lett., 29 (1996) 351. M. Soylak, I. Narin, L. Elci and M. Dogan, Trace Elem. Electrolytes, 16 (1999) 131. M. Yaman and S. Gu¨cer, Analyst, 120 (1995) 101. T. Kubota, K. Suzuki and T. Okutani, Talanta, 42 (1995) 949. T. Ferri, E. Cardarelli and B.M. Petronio, Talanta, 36 (1989) 513. M. Soylak, U. Divrikli, L. Elci and M. Dogan, Talanta, 56 (2002) 565.

Solid-phase extraction 225 226 227 228 229 230 231 232 233 234 235 236 237 238 239 240 241 242 243 244 245 246 247 248

K. Pyrzynska, Analyst, 120 (1995) 1933. A. Bhalotra and B.K. Puri, J. AOAC Int., 84 (2001) 47. B.K. Puri, M. Satake, G. Kano and S. Usami, Anal. Chem., 59 (1987) 1850. B.K. Puri and S. Balani, Talanta, 42 (1995) 337. C.F. Poole, A.D. Gunatilleka and R. Sethuraman, J. Chromatogr. A, 885 (2000) 17. M. Pesavento and E. Baldini, Anal. Chim. Acta, 389 (1999) 59. K. Hirose, Y. Dokiya and Y. Sugimura, Mar. Chem., 11 (1982) 343. P.J.M. Buckley and C.M.G. van den Berg, Mar. Chem., 19 (1986) 281. W. Frenzel, Fresenius J. Anal. Chem., 361 (1998) 774. S. Ahmad, R.C. Murthy and S.V. Chandra, Analyst, 115 (1990) 287. I. Kubrakova, T. Kudinova, A. Formanovsky, N. Kuz’min, G. Tsysin and Y. Zolotov, Analyst, 119 (1994) 2477. D.M. Adria-Cerezo, M. Llobat-Estelles and A.R. Mauri-Aucejo, Talanta, 51 (2000) 531. M.M. Gibbs, Water Res., 13 (1979) 295. C.-Y. Liu, Anal. Chim. Acta, 192 (1987) 85. E. Yamagami, S. Tateishi and A. Hashimoto, Analyst, 105 (1980) 491. H. Emteborg, D.C. Baxter and W. Frech, Analyst, 118 (1993) 1007. T. Ferri and P. Sangiorgio, Anal. Chim. Acta, 321 (1996) 185. ¨ rnemark and A. Olin, Talanta, 41 (1994) 67. U. O T. Kubota and T. Okutani, Anal. Chim. Acta, 351 (1997) 319. D. Tanzer and K.G. Heumann, Anal. Chem., 63 (1991) 1984. W.M.R. Dirkx, R. Lobinski and F.C. Adams, Anal. Chim. Acta, 286 (1994) 309. J.L. Gomez-Ariza, E. Morales, I. Giraldez, D. Sanchez-Rodas and A. Velasco, J. Chromatogr. A, 938 (2001) 211. J. Szpunar-Lobinska, M. Ceulemans, R. Lobinski and F.C. Adams, Anal. Chim. Acta, 278 (1993) 99. Y. Cai and J.M. Bayona, J. Chromatogr. Sci., 33 (1995) 89.

457

This page intentionally left blank

Chapter 15

Chelation solvent extraction for separation of metal ions Hideyuki Itabashi and Taketoshi Nakahara

15.1

INTRODUCTION

Solvent extraction (IUPAC recommends the term “liquid –liquid extraction”) can provide practical methods for separation and preconcentration of metal ions in aqueous samples. Because most of the extraction methods do not need any special apparatus or skills, sample preparation using the solvent extraction has widely been used in the analytical ﬁeld. In addition, a large number of thermodynamic constants characterizing the extraction reaction of metal ions have been measured and enable selection of some proper extractants and appropriate experimental conditions. In particular, chelate extraction, in which chelation between metal ions and chelating extractants plays an important role, has many associated practical data, such as extraction constants [1]. These data permit the design of chelate extraction systems for sample preparation in trace metal analysis without undertaking preliminary experiments. In this chapter, chelate extraction of metal ions from aqueous samples is described as the pretreatment technique prior to instrumental analyses such as atomic absorption spectrometry (AAS), inductively coupled plasma-optical emission spectrometry (ICP-OES), and inductively coupled plasma-mass spectrometry (ICP-MS). In the ﬁrst part of this chapter, theoretical considerations relating to extraction methods for preconcentration, mutual separation, and speciation of metal ions are discussed, focusing on the selection of chelating agents and setting of extraction conditions. The separation of metal ions using chelating resins is also brieﬂy described, because the mechanism of adsorption of metal ions onto such resins is similar to that of the chelate extraction of metal ions. Kinetically controlled separation, which may be signiﬁcant for the separation of labile metal ions from inert ones [2,3], is not discussed here because the actual sample preparation using this extraction procedure is usually done in the equilibrium state. In the second part of this chapter, some experimental procedures using the chelation-extraction and chelating resins Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

459

H. Itabashi and T. Nakahara

are described. This section provides some practical procedures and concrete recipes for the extraction of metal ions from various samples. It is hoped that this approach will provide some practical background for sample preparation for trace metal analysis using chelation.

15.2

THEORETICAL CONSIDERATIONS

15.2.1 General principles In order to extract metal ions from an aqueous phase into an organic phase, it is necessary to neutralize their ionic charge. In chelate extraction, the neutralization is achieved by the complexation between the metal ions and the chelating agent anions. The anions, which act as a Lewis base, are formed by deprotonation of the chelating agents that are weak organic (lipophilic) acids. Thus, the anions not only neutralize the ionic charge of the metal ions, but also coordinate to the metal ions and then substitute for the water molecules in the coordination sphere of the metal ions. Consequently, the resultant metal chelates have a hydrophobic character and transfer to the organic phase. In this section, the extraction reaction is characterized using some equilibrium constants, and an approach to the selection of proper extraction systems for sample preparation is presented. Hereafter, “HR” is used to represent a generic chelating agent to simplify the description of equations, although some chelating agents may act as two or more protic acids, i.e., H2R, H3R, etc. 15.2.1.1 Equilibrium constants for extraction The chelate extraction reaction is conventionally shown as a series of four stepwise processes (Fig. 15.1): the distribution of the chelating agent, HR, between the organic and aqueous phases; the acid dissociation of the agent in the aqueous phase; the complex formation reaction between the n-valent metal ion, Mnþ , and the chelating agent anion; and the distribution of the formed metal chelate, MRn , between the two phases. HR O HRorg ;

Kdr ¼ ½HRorg ½HR21

Fig. 15.1. Extraction of Mnþ with HR.

460

ð15:1Þ

Chelation solvent extraction for separation of metal ions

HR O R2 þ Hþ ; M

nþ

2

þ nR O MRn ;

MRn O MRn;org ;

Ka ¼ ½R2 ½Hþ ½HR21 Kf ¼ ½MRn ½M

nþ 21

ð15:2Þ 2 2n

½R

Kdc ¼ ½MRn org ½MRn 21

ð15:3Þ ð15:4Þ

The subscript org denotes species in the organic phase and Kdr , Ka , Kf , and Kdc represent the distribution constant for the chelating agent, the acid dissociation constant of the agent, the formation constant of the metal chelate, and the distribution constant of the metal chelate, respectively. The overall extraction reaction is expressed as: Mnþ þ nHRorg O MRn;org þ nHþ

ð15:5Þ

which gives an equilibrium constant as follows: Kex ¼ ½MRn org ½Hþ n ½Mnþ 21 ½HR2n org

ð15:6Þ

where Kex is deﬁned as the “extraction constant”. Equations (15.1)–(15.4) are substituted into Eq. (15.6), and the following relationship is obtained: 2n Kex ¼ Kan Kf Kdc Kdr

ð15:7Þ

Equation (15.7) indicates that Kex is the product of the constants for each stepwise reaction and that each constant must be evaluated to completely understand the overall extraction. However, in practice, it is more important to understand the relationship between the extraction constant and the degree of extraction of metal ions because reliable Kex values, which have been reported for various chelate extraction systems, allow the design of suitable extraction systems. In subsequent sections, parameters relevant to the selection of proper chelating agents and the optimization of experimental conditions for sample preparation are described. 15.2.1.2 Relationship between the extraction constant and distribution ratio Consider the extraction of a þ n-valent metal ion, Mnþ , in an aqueous phase with a chelating agent, HR, in an organic phase. Assume that the only species of M in the aqueous phase is Mnþ , and that in the organic phase it is MRn , other M species do not exist in either phase. In this case, the distribution ratio of the metal species, D { ¼ (total concentration of M in the organic phase)/(total concentration of M in the aqueous phase)}, is expressed as: D ¼ ½MRn org ½Mnþ 21

ð15:8Þ

Equations (15.6) and (15.8) give the following relationship: D ¼ Kex ½HRnorg ½Hþ 2n log D ¼ log Kex þ nlog½HRorg þ npH

ð15:9Þ ð15:10Þ

461

H. Itabashi and T. Nakahara

Equation (15.10) shows that a plot of log D against pH (or log[HR]org) gives a straight line with slope n and that the distribution ratio is determined by Kex and experimental conditions ([HR]org and pH). Namely, if Kex is known, we can easily calculate the D value from [HR]org and pH. Thus, if a large D is necessary for the extraction of the metal ion of interest, we should select a chelating agent with a large Kex and use experimental conditions of higher concentration of [HR]org and higher pH. However, it is necessary to pay attention to the solubility of the chelating agent in the organic phase and the formation of the hydroxo complex of the metal ion. For example, the solubility of 3-mercapto-1,5-diphenylformazan (dithizone), which is usually used to extract soft (as Lewis acid) metal ions, is about 2 £ 1023 mol dm23 in carbon tetrachloride at 208C. In the case of the extraction of some trivalent metal ions, such as Fe(III), whose hydroxo complex, Fe(OH)3, has extremely small solubility in water, the pH of the solution should be adjusted to below 3. Therefore, it may be difﬁcult to obtain the optimum D value by changing the experimental conditions if the chelating agent has low solubility in the organic solvent and the metal ion forms a stable complex with hydroxide ion. On the other hand, even if the above problem can be bypassed, the maximum value of D for metal ions depends on the Kdc value as follows: for large D, the concentration of Mnþ in the aqueous phase becomes extremely small, then the assumption of Eq. (15.8), in which predominant species of M in the aqueous phase is only Mnþ, will not be satisﬁed, since the relative concentration of MRn to Mnþ in the aqueous phase increases with increase in the D value. Thus, it is necessary to consider the existence of MRn in the aqueous phase. In this case, the distribution ratio is deﬁned as: D ¼ ½MRn org ð½Mnþ þ ½MRn Þ21

ð15:11Þ

Equations (15.3), (15.4) and (15.11) give: log D ¼ log Kdc þ log{Kf ½R2 n ð1 þ Kf ½R2 n Þ21 }

ð15:12Þ 2

Equation (15.12) indicates that a plot of log D against log[R ] gives a smooth curve that is of the normalized form log Y ¼ log{Xð1 þ XÞ21 }, in which Y ¼ 21 DKdc and X ¼ Kf ½R2 n : A typical plot of log D against log[R2] is shown in Fig. 15.2. When [R2] is very small, i.e., Kf ½R2 ,, 1, Eq. (15.12) is rewritten as: log D ¼ log Kdc þ log Kf þ nlog½R2

15:13Þ

2

A plot of log D against log[R ] gives a straight line with a slope of n, whereas if [R2] becomes very large, i.e., Kf ½R2 .. 1, Eq. (15.12) gives: log D ¼ log Kdc

ð15:14Þ

The horizontal line displayed in Fig. 15.2 means that the maximum value of D, Dmax , corresponds to Kdc , i.e., an extraction system with a large Kdc will result in a large Dmax value. The Kdc value depends on the solubility of MRn in the

462

Chelation solvent extraction for separation of metal ions

Fig. 15.2. Typical log D vs. log½R2 curve in the extraction of Mnþ with HR.

organic phase: the greater hydrophobicity of MRn will give the larger Kdc value. Therefore, to design the extraction system to yield a large Dmax , a chelating agent possessing a more lipophilic character should be selected. The use of synergism, by which hydrophobic extracts are formed will also be useful for this purpose, as described later. 15.2.1.3 Effect of HSAB nature of chelating agents on extraction constants Complexation reactions are usually described by the hard-soft acid –base (HSAB) principle [4], which states that hard acids prefer hard bases, and soft acids prefer soft bases. From this principle, it is expected that the chelating agents having a hard base nature will form stable complexes with hard metal ions and that those with a soft one will form stable complexes with soft metal ions. The stability of the resulting metal chelate strongly affects the Kex value, since the stability constant, Kf , is directly proportional to Kex , as shown in Eq. (15.7). Therefore, hard chelating agents will provide a large Kex value for the extraction of hard metal ions, whereas soft ones will give the extraction of soft metal ions a large Kex value. Table 15.1 shows the Kex values of the extraction systems in which some bidentate chelating agents, such as 4,4,4-triﬂuoro-1-(2-thienyl)-1,3-butanedione (TTA), 8-quinolinol, dithizone, and diethyldithiocarbamate (DDTC) are used. The coordination atoms of these chelating agents are as follows: two oxygens for TTA, a nitrogen and oxygen for 8-quinolinol, a nitrogen and sulfur for dithizone, and two sulfurs for DDTC. In general, the hardness of coordination atoms, as Lewis base, increases in the order: S ,, N , O. This order shows that the hardness of these chelating agents will increase in the following order: DDTC , dithizone ,, 8-quinolinol , TTA. Therefore, in

463

H. Itabashi and T. Nakahara TABLE 15.1 Extraction constants (log Kex) of metal chelates Metal ions

Ag Al Am(III) Ba Be Bi Ca Cd Ce(III) Co(II) Cu(II) Eu Fe(III) Ga Gd Hf Hg Ho In La Lu Mg Mn(II) Mo(VI) Nd Ni Pa(IV) Pb Pd Pm(III) Pr Pu(III) Pu(IV) Pu(VI) Sc

TTA (benzene)

25.25 27.45 214.4 23.2 23.3 212 29.45 26.7 21.3 27.65 23.3

8-Quinolinol (chloroform)

Dithizone (carbon tetrachloride)

DDTC (carbon tetrachloride)

7.18

11.9

9.75

16.8

2.15

5.4

1.59 10.5

2.3 13.7

25.2 221b 21.2 217.9a 25.9b 22.15b 1.75 4.1 3.7

1.2 ,2 1.30

27.6 7.8– 8.2 ,2 3 27.25 24.35 210.5 26.75

20.9 216.35

26.8 4.85

215.15 29.3 9.9

32 10.35

24.4

28.55 22.2 6.7 25.2 28.05 28.5, 28.85 24.45 6.85, 6.35 21.8 20.75

28.05 ,15

21.19 0.38

7.8 .32

26.65 continued

464

Chelation solvent extraction for separation of metal ions

TABLE 15.1 (continuation) Metal ions

TTA (benzene)

Sm Sn Sr Tb Th Ti Tl(I) Tl(III) Tm U(IV) U(VI) V(V) Y Yb Zn Zr

27.7

8-Quinolinol (chloroform)

Dithizone (carbon tetrachloride)

DDTC (carbon tetrachloride)

22 214 27.5 20.7, 21.4 25.2 25.7 26.95 5.3 22.25, 22.0

219.7b 27.2 0.9 28.8 ,5

23.3

20.5

2.3

3.0

21.6a 1.65

27.4 26.7 9.2– 9.6

22.4b 2.7

TTA: thenoyltriﬂuoroacetone, DDTC: diethyldithiocarbamate, the solvents in parentheses represent the organic phase in each extraction system. Formed complex: a, MRn(HR); b, MRn(HR)2. Source: Ref. [5].

the extraction systems using these chelating agents, the Kex values of hard metal ions will increase in the above order, whereas that of soft ones will increase in the opposite order. In practice, hard metal ions such as Al(III) and Be(II) will be quantitatively extracted into the organic phase by using TTA or 8-quinolinol, whereas DDTC and dithizone generally will not extract such metal ions. In contrast, for the extraction of soft metal ions, such as Ag(I) and Hg(II), a larger distribution ratio will be obtained by use of DDTC or dithizone than by use of TTA or 8-quinolinol under the same experimental conditions. Therefore, in the case of separation of hard metal ions from soft ones, hard extractants should be used, while soft extractants should be selected for the separation of soft metal ions from hard ones. 15.2.2 Preconcentration of metal ions 15.2.2.1 Enrichment factor For solvent extraction, preconcentration of metal ions in an aqueous sample is usually performed by extracting the metal ions from a large volume of the

465

H. Itabashi and T. Nakahara

sample solution into a small volume of an organic phase. For example, if a metal ion is quantitatively extracted from 1000 cm3 of an aqueous phase into 10 cm3 of an organic phase, the concentration of the metal ion in the organic phase after the extraction becomes 100-fold larger than it was in the aqueous phase before the extraction. Enrichment factor (EF) is deﬁned as the ratio of the concentration of the metal ion in the organic phase after the extraction to that in the aqueous phase before the extraction. When the extraction of the metal ion into the organic phase is quantitative, EF is equal to the volume ratio of the aqueous to organic phases. In this case, the metal concentration in the sample solution [M]s is expressed as follows: ½Ms ¼ ½Morg Vorg V 21 ¼ ½Morg ðEFÞ21

ð15:15Þ

where V and Vorg denote the volume of the aqueous and organic phases, respectively, and [M]org is the concentration of species M in the organic phase after extraction. Thus, from Eq. (15.15), [M]s can be obtained easily through the measurement of [M]org. On the other hand, if the extraction of the metal ion is not quantitative, the [M]s value cannot be obtained only through the measurement of [M]org, because 21 Þ: In this case, EF in the extraction system is not equal to the volume ratio ðVVorg the EF values must be determined for each experimental condition. The measurement of EF would be a rather tedious task for many bench chemists, so it is preferable to select a chelating agent that can quantitatively extract the metal ion of interest into the organic phase. The degree of extraction can be evaluated by the extractability of metal ions. The extractability E (%) is deﬁned as: 21 21 Eð%Þ ¼ 100½Morg Vorg ð½Morg Vorg þ ½MVÞ21 ¼ 100DðD þ VVorg Þ

ð15:16Þ

where [M] represents the concentration of species M in the aqueous phase after the extraction. Equation (15.16) shows that the extractability decreases with 21 an increase in the volume ratio of the two phases ðVVorg Þ: For example, if 21 ¼ 1, the extraction system in which D ¼ 100 gives E . 99%, whereas VVorg 21 if VVorg ¼ 100, E decreases to 50%. Therefore, to quantitatively extract the metal ion into the organic phase ðE . 99%Þ, the distribution ratio achieved 21 under the experimental conditions should be larger than 99VVorg : 15.2.2.2 Synergism As described in the above section, a large D value is necessary for the practical preconcentration of metal ions. To obtain a large D value, the addition of a neutral hydrophobic (lipophilic) Lewis base to the organic phase in the chelate extraction system is known to be effective. The phenomenon in which D is increased by the addition of a neutral base is called “synergism” [6]. The primary cause for synergism is generally explained by the formation of a hydrophobic adduct.

466

Chelation solvent extraction for separation of metal ions

For synergism, suppose that the following reaction occurs when a neutral base, B, is added to the reaction shown in Eq. (15.5): Mnþ þ nHRorg þ mBorg O MRn Bm;org þ nHþ

ð15:17Þ

The extraction constant involving the adduct formation is deﬁned as: 2m K 0ex ¼ ½MRn Bm org ½Hþ n ½Mnþ 21 ½HR2n org ½Borg

ð15:18Þ

Equations (15.6) and (15.18) give: 21 2m ¼ ½MRn Bm org ½MRn 21 K 0ex Kex org ½Borg ¼ Kad

ð15:19Þ

where Kad is the adduct formation constant. For reaction (15.17), the distribution ratio, D0 , is deﬁned as: nþ 21 D0 ¼ ½MRn Bm org ½Mnþ 21 ¼ ½MRn org Kad ½Bm ¼ DKad ½Bm org ½M org

ð15:20Þ

Equation (15.20) indicates that the addition of B with a large Kad value advantageously acts on the extraction of M into the organic phase. Several Kad values for TTA extraction are shown in Table 15.2, and the effect of the addition of tributyl phosphate (TBP) on the degree of extraction of U(VI) with TTA is shown in Fig. 15.3. These results illustrate that chelate extraction, when accompanied by synergism is a useful technique for the preconcentration of metal ions. On the other hand, when the extraction is conducted under large EF conditions, the solubility of the organic solvent in the aqueous phase must be considered. In this case, an organic solvent having low solubility in water must be selected; otherwise the organic phase may disappear by dissolution into the aqueous phase. Table 15.3 lists the solubility of representative organic solvents in water. If the solubility of the chelating agent in the organic solvent is large enough, an organic solvent with low solubility in water, such as n-hexane, should be selected for this purpose. 15.2.3 Mutual separation of metal ions 15.2.3.1 Separation factor Consider the mutual separation of two metal ions, Mnþ and Nnþ , by the extraction shown in Eq. (15.5), (assume V ¼ Vorg ). In the separation of Mnþ from Nnþ , the separation factor (SF) is deﬁned as: 21 SF ¼ DðMÞ D21 ðNÞ ¼ KexðMÞ KexðNÞ

ð15:21Þ

where DðMÞ , DðNÞ , KexðMÞ , and KexðNÞ represent the distribution ratios and the extraction constants for M and N, respectively. Assuming that DðMÞ . 100 and DðNÞ , 0:01, i.e., EðMÞ . 99% and EðNÞ , 1%, quantitative separation of M from N can be achieved, and the following SF value as the condition for quantitative separation can be obtained: SF ¼ KexðMÞ =KexðNÞ . 104 : In other words, for the

467

H. Itabashi and T. Nakahara TABLE 15.2 Adduct ðMRn Bm Þ formation constants of metal TTA chelates ðMRn Þ with some neutral ligands (B) in carbon tetrachloride MRn

AcR3 AmR3 BaR2

CaR2

CoR2 CuR2 EuR3

LaR3 LuR3 ScR3 SrR2

ThR4 ZnR2

B

TBP MIBK TBP MIBK TBP MIBK TOPO TBP MIBK TOPO TBP TOPO TBP MIBK TBP MIBK TOPO TBP MIBK TBP MIBK TBP TBP MIBK TOPO TBP TBP MIBK

log Kad m¼1

m¼2

4.2 1.3 5.06 1.8 2.62 1.7 5.68 4.11 1.8 5.64 2.38 6.13 2.27 0.47 5.15 1.8 7.49 4.83 2.0 5.69 1.7 3.44 3.76 1.8 5.39 5.18 4.34 1.22

8.9 3.1 8.89 2.4 5.84 2.3 9.98 8.22 2.7 10.68 3.68 9.30 – – 8.89 2.5 12.26 9.33 2.9 6.67 – – 7.52 2.6 9.78 – – –

TBP: tributyl phosphonate, TOPO: trioctylphosphineoxide, MIBK: methyl isobutyl ketone. Source: Ref. [7].

separation of M from N, it is sufﬁcient to select a chelating agent whose Kex satisﬁes the SF . 104 : From the practical standpoint, however, it is necessary to pay attention to experimental conditions. For example, dithizone satisﬁes the condition of

468

Chelation solvent extraction for separation of metal ions

Fig. 15.3. Synergistic extraction: distribution of U(VI) between 0.01 M HNO3 and mixture of TTA – TBP at constant total molarity ([TTA]org þ [TBP]org ¼ 0.02 mol dm23) in cyclohexane. Source: Ref. [8].

SF . 104 for the separation of Hg(II) from Cu(II), since KexðHgÞ ¼ 1032 and KexðCuÞ ¼ 1013:7 : However, the condition for quantitative separation of Hg(II) from Cu(II), i.e., log DðHgÞ ¼ log KexðHgÞ þ 2log½HRorg þ 2pH . 2 and log DðCuÞ ¼ log KexðCuÞ þ 2log½HRorg þ 2pH , 22 leads to the following relationship: 2 2 log KexðHgÞ , 2log½HRorg þ 2pH , 22 2 log KexðCuÞ

ð15:22Þ

Equation (15.22) suggests that the pH of the aqueous phase should be adjusted to 22.9 from 210 even when ½HRorg ¼ 1025 mol dm23 : It is impossible to prepare a solution where the pH ¼ 22:9, thus, separation of Hg(II) from Cu(II) under the above-mentioned conditions is not possible using a dithizone extraction. To separate the Hg(II) from Cu(II), a masking agent, described later, will have to be used. In the following example, consider the separation of a trace amount of metal ion M from a major amount of metal ion N; i.e., ½N=½M ¼ 10x ðx . 2Þ: If the condition for the quantitative separation of M from N is EðMÞ . 99% and ½Morg =½Norg . 102 , log D for M and N can be expressed as follows: log DðMÞ ¼ log KexðMÞ þ nlog½HRorg þ npH . 2

ð15:23Þ

log DðNÞ ¼ log KexðNÞ þ nlog½HRorg þ npH , 22 2 x

ð15:24Þ

469

H. Itabashi and T. Nakahara TABLE 15.3 Solubility of some organic solvents in water at 258C Solvents

Solubility (wt%)

c-Hexane n-Hexane Benzene Toluene p-Xylene Chloroform Carbon tetrachloride 1,1-Dichloroethane 1,2-Dichloroethane Chlorobenzene 1,2-Dichlorobenzene 1-Butanol Isobutyl alcohol 1-Pentanol Isoamyl alcohol Diethyl ether Diisopropyl ether Bis(2-chloroethyl) ether Methyl isobutyl ketone Propylene carbonate Nitrobenzene

0.0055 0.00123 0.179 0.0515 0.0156 0.815a 0.077 5.03a 0.81a 0.0488b 0.0156 7.45 10 2.19 2.97 6.04 1.2a 1.02a 1.7 17.5 0.19a

a

At 208C. At 308C.

b

Equations (15.23) and (15.24) give the following relationship: ð2 2 log KexðMÞ Þn21 , log½HRorg þ pH , ð22 2 x 2 log KexðNÞ Þn21

ð15:25Þ

To satisfy Eq. (15.25), the ratio of the extraction constant of M and N, KexðMÞ =KexðNÞ , should be larger than 104þx : Thus, in the separation of trace metal ions from some matrices, it is necessary to consider the ratio of the concentration of trace metal ions to that of principal ones in the sample solution. 15.2.3.2 Masking The use of a masking agent is often effective for the mutual separation of metal ions. A water-soluble complexing agent is usually used for the masking agent in the solvent extraction. In this case, consider that a masking agent, L, is added to the aqueous phase containing a metal ion, Mnþ , and water soluble

470

Chelation solvent extraction for separation of metal ions

(not extracted) complexes ML; ML2 ; …; MLp are formed (charges of L and ML complexes are omitted to keep the equations simpler), i.e., Mnþ þ L O ML; M

nþ

þ 2L O ML2 ; .. .

b1ðMÞ ¼ ½ML½Mnþ 21 ½L21

ð15:26Þ

22

ð15:27Þ

bpðMÞ ¼ ½MLp ½Mnþ 21 ½L2p

ð15:28Þ

b2ðMÞ ¼ ½ML2 ½M

nþ 21

½L

.. .

Mnþ þ pL O MLp ;

where b represents the stability constant of the formed complex. If the extraction reaction of Mnþ with HR occurs according to Eq. (15.5), the distribution ratio, D0ðMÞ , involving ML complexes can be written as: D0ðMÞ ¼ ½MRn org ð½Mnþ þ ½ML þ ½ML2 þ · · · þ ½MLp Þ21 ¼ ½MRn org ½Mnþ 21 ð1 þ b1ðMÞ ½L þ b2ðMÞ ½L2 þ · · · þ bpðMÞ ½Lp Þ21 ¼ DðMÞ a21 LðMÞ

ð15:29Þ

where aLðMÞ represents the side reaction coefﬁcient of the extraction reaction of Mnþ : Similarly, for the extraction of Nnþ , the distribution ratio, D0ðNÞ , is expressed as follows: D0ðNÞ ¼ DðNÞ a21 LðNÞ

ð15:30Þ

The separation factor, SF0, for the separation of Mnþ from Nnþ in the presence of L is deﬁned as: 0

21 21 SF0 ¼ D0ðMÞ DðNÞ ¼ KexðMÞ KexðNÞ aLðNÞ a21 LðMÞ

ð15:31Þ nþ

nþ

Equation (15.31) shows that quantitative separation of M from N would be 21 , 104 if the value of aLðNÞ a21 attained even when KexðMÞ KexðNÞ LðMÞ is large enough. For example, in the extraction of U(VI) with TTA, ethylenediaminetetraacetic acid (EDTA) acts as a useful masking agent for diverse metal ions, because the stability constant for U(VI) –EDTA complex, b1ðUÞ ¼ 1010:4 , is signiﬁcantly smaller than that for other metal complexes such as Al(III)-, Fe(III)-, and Cu(II)-EDTA, etc. ðb1ðA1Þ ¼ 1016:1 , b1ðFeÞ ¼ 1025:1 , b1ðCuÞ ¼ 1018:8 Þ: In this case, such diverse metal ions are masked with EDTA, whereas U(VI) can be quantitatively extracted into the organic phase as the TTA complex. In the foregoing reaction, i.e., the separation of Hg(II) from Cu(II) using dithizone, EDTA also acts as an effective reagent: Hg(II) is extracted into the organic phase while Cu(II) is masked with EDTA. Thus, the use of a masking agent often provides a desirable result for mutual separation of metal ions. However, when the masking agent forms a stable complex with the metal ion of interest, it is necessary to pay attention to the extent of complex formation between the metal ion and the masking agent.

471

H. Itabashi and T. Nakahara

This occurs because the distribution ratio of the metal ion of interest also decreases by adding the masking agent. In this case, log D0 achieved under the experimental conditions should be estimated from Eq. (15.32) before the experiment. log D0 ¼ log Kex þ nlog½HRorg þ npH 2 log aL

ð15:32Þ

The aL values of several masking agents at various pH are summarized in Ref. [9]. 15.2.4 Speciation of metal ions in natural water 15.2.4.1 Estimation of free metal ion concentration in natural water In the study of the speciation of metal ions in natural water, the fractional determination of free metal ions (aquo complexes) and complexed metal ions is one of the most important tasks, because the toxicity of the metal ions to aquatic organisms depends heavily on their chemical forms [10]. Chelate extraction can provide a method for the speciation of metal ions in natural water if the complex formation reaction between free metal ions and naturally occurring ligands is considered to be the side reaction of the extraction [11]. In this section, consider the 1:1 (metal:ligand) complex formation between a free metal ion, Mnþ , and a naturally occurring ligand, L, in a water sample (charges of L and ML are omitted to keep the equations simpler). The complexation reaction can be written as: Mnþ þ L O ML

ð15:33Þ

for which the stability constant, b, is deﬁned by:

b ¼ ½ML½Mnþ 21 ½L21

ð15:34Þ

The total concentration of M’s species in the water sample, [M]w, can be expressed as: ½Mw ¼ ½Mnþ þ ½ML ¼ ½Mnþ ð1 þ b½LÞ ¼ ½Mnþ aL

a21 L

ð15:35Þ

where is the mole fraction of M in the water sample: ¼ ½M ½M21 w : nþ On the other hand, assuming that the extraction of M with HRorg occurs according to Eq. (15.5), we can obtain the following equations. The absolute amount (mole) of species M in the extraction system, M(mol), is expressed as follows: nþ

a21 L

MðmolÞ ¼ V½Mw þ Vorg ½MRn org

nþ

ð15:36Þ

From Eqs. (15.6), (15.35) and (15.36), Eq. (15.37) is derived: 21 þ n 21 21 2n ½MRn 21 org ¼ Vorg MðmolÞ þ aL Vorg ½H Kex MðmolÞ ½HRorg

472

ð15:37Þ

Chelation solvent extraction for separation of metal ions þ n 2n According to Eq. (15.37), a plot of ½MRn 21 org against ½H ½HRorg should give a 21 21 straight line with a slope of aL Vorg Kex MðmolÞ and an intercept Vorg M21 ðmolÞ : If Kex is known, the aL and MðmolÞ values can be calculated from the slope and the intercept. From these values, the concentration of the free metal ion, ½Mnþ , and that of the complexed metal ion, [ML], can be estimated. The method has been applied to the speciation of Cu(II) in river water samples [11]. The results indicated that most of the Cu(II) in the samples existed as complexes with naturally occurring ligands; [Cu2þ] was estimated to be 10213 mol dm23.

15.2.4.2 Estimation of ligand concentrations and stability constants of formed metal complexes The information concerning the concentrations of naturally occurring ligands and the stability constants of formed metal complexes makes possible the speciation of metal ions in natural water, because the equilibrium calculations using the information can provide their chemical forms in the water. The chelate extraction in which the equilibrium calculations are involved permits the simultaneous determination of the ligand concentrations and the stability constants of the formed metal complexes [12]. In this method, consider the back-extraction in which a metal ion moves from the organic phase to the aqueous phase. The back-extraction of Mnþ is expressed as: MRn;org O Mnþ þ nR2

ð15:38Þ

which gives a mass action constant Kbex : Kbex ¼ ½Mnþ ½R2 n ½MRn 21 org

ð15:39Þ

On the other hand, assuming that i types of ligands, L1 ; L2 ; …; Li , exist in a water sample and complexations of Mnþ with the ligands proceeds as given by Eq. (15.40), we can introduce the following equations (charges of L and ML are omitted for simplicity): Mnþ þ Li O MLi

ð15:40Þ

the stability constant ðbLi Þ for ML is deﬁned as: i

bLi ¼ ½MLi ½Mnþ 21 ½Li 21

ð15:41Þ

The total concentration of M’s species in the aqueous phase is expressed as: ½Mtot ¼ ½Mnþ þ ½ML1 þ ½ML2 þ · · · þ ½MLi

15:42Þ

If the concentration of free species of M in the aqueous phase is negligible, the total concentration of species M in the aqueous phase is equal to ½MLtot

473

H. Itabashi and T. Nakahara

ð½MLtot ¼ ½ML1 þ ½ML2 þ · · · þ ½MLi Þ: By considering the mass balances, we obtain: 2½MLtot þ ½HRorg;init ¼ ½R2 þ ½HR þ ½HRorg ½Li tot ¼ ½Li þ ½MLi

ð15:43Þ 15:44Þ

where ½HRorg;init is the concentration of HR added to the organic phase before shaking and ½Li tot is the total concentration of Li : From the above equations, the following relationship is obtained: ½MLtot ¼

i X

{½Li tot ð1 þ 10log x2log bLi Þ21 }

ð15:45Þ

i¼1 21 21 þ 21 where x ¼ ð2½MLtot þ ½HRorg;init Þ2 a22 R Kbex ½MRn org and aR ¼ 1 þ ½H Ka þ Kdr ½Hþ Ka21 : According to Eq. (15.45), [ML]tot is plotted as a function of log x. The application of a nonlinear least-squares method to the resulting plots gives ½Li tot and log bLi : The data obtained can provide the mole fractions of each species by using the following equations: 1 nþ 21 ½Mnþ ½M21 þ bL2 ½L2 tot tot ¼ {1 þ bL1 ½L tot ð1 þ ½M bL1 Þ

ð1 þ ½Mnþ bL2 Þ21 þ · · · þ bLi ½Li tot ð1 þ ½Mnþ bLi Þ21 }21 ¼ a21 L

ð15:46Þ

1 1 21 ½ML1 ½M21 tot ¼ bL1 ð½L tot 2 ½ML ÞaL

ð15:47Þ

2 2 21 ½ML2 ½M21 tot ¼ bL2 ð½L tot 2 ½ML ÞaL

ð15:48Þ

.. . i i 21 ½MLi ½M21 tot ¼ bLi ð½L tot 2 ½ML ÞaL

ð15:49Þ

The method has been applied to the speciation of Cu(II) in natural waters [12]. The results indicated that most of the Cu(II) in natural waters existed as complexes combined with ligands derived from humic substances [13].

15.3

ADSORPTION OF METAL IONS USING CHELATING RESINS

The principle of adsorption of metal ions on chelating resins can be considered as well as that of chelate extraction. In this section, general principles are brieﬂy described, followed by the features of resins used in the analytical ﬁeld.

474

Chelation solvent extraction for separation of metal ions

15.3.1 General principles Chelating resins have functional groups capable of forming chelates with metal ions. These functional groups are usually weak organic acids containing oxygen, nitrogen, or sulfur as coordinating atoms. When the chelating resins are treated with an acid solution, their functional groups combine with hydrogen ion and, as such, the functional groups can be expressed as Hm R (R represents the functional groups of the resin). The adsorption of Mnþ on the chelating resin is expressed as: Mnþ þ Hm Rr O MRrðn2mÞþ þ mHþ

ð15:50Þ

which gives a mass action constant KadðMÞ : KadðMÞ ¼ ½MRðn2mÞþ r ½Hþ m ½Mnþ 21 ½Hm R21 r

ð15:51Þ

where r represents the resin phase. In this reaction, the distribution ratio of M can be expressed as follows: DðMÞ ¼ ½MRðn2mÞþ r ½Mnþ 21

ð15:52Þ

Equations (15.51) and (15.52) give the following relationship, which is analogous to Eq. (15.10) obtained in the chelate extraction system: log DðMÞ ¼ log KadðMÞ þ log½Hm Rr þ mpH

ð15:53Þ

Equation (15.53) shows that a resin having a large KadðMÞ and ½Hm Rr provides large DðMÞ and adsorption of M on the resin becomes advantageous with an increase in pH. Similarly, the distribution ratio of Nnþ is expressed as: DðNÞ ¼ ½NRðn2mÞþ r ½Nnþ 21

ð15:54Þ

The separation factor, which is usually called “selectivity coefﬁcient”, is deﬁned as 21 SF ¼ DðMÞ D21 ðNÞ ¼ KadðMÞ KadðNÞ

ð15:55Þ

In this case, almost the same considerations as described for chelate extraction are applicable. The principles of separation of metal ions using chelating resins are quite similar to those already discussed in the above section and are not further described here. Further information can be found in the literature [14]. 15.3.2 Features of some chelating resins Chelating resins having different functional groups, such as carboxyl, hydroxyl, amino, thiol, etc., can be used for the separation and preconcentration of metal ions in water samples. Various chelating resins having iminodiacetate, dithiocarbamate, amidoxime, etc., functional groups have been synthesized.

475

H. Itabashi and T. Nakahara

Fig. 15.4. Structural formulas of several functional groups in chelating resins.

Among them, chelating resins containing iminodiacetate functional groups are the most widely used in the analytical ﬁeld. Figure 15.4 (a) shows the structure of the iminodiacetate functional group. The complexation behavior of the resin is analogous to that of EDTA, thus many metal ions form stable complexes with the functional group of the resin. Although several commercial resins with this functional group are available, Chelex-100 (Bio-Rad Laboratories) is the most widely used. The following afﬁnity series of metal ions to Chelex-100 has been reported [15] for divalent cations: Hg . U(VI) . Cu . V(V) . Pb . Ni . Cd . Zn . Co . Fe(II) . Mn(II) . Be . Ca . Mg . Ba . Sr. For trivalent metals: Cr . In . Fe . Ce . Al . La. For alkali metals: Li . Na . K . Rb . Cs. Metal ions such as Cd, Co, Cu, Mn, Ni, Pb, and Zn have been separated from sea water using Chelex-100 resin. Pai [16] reported that these metal ions were quantitatively collected using the Mg form of the resin from sea water whose pH was preliminary adjusted to 6.5. Haraguchi and co-workers [17–26]

476

Chelation solvent extraction for separation of metal ions

preconcentrated many metal ions, including rare earth elements, from natural waters or biological samples using Chelex-100 resin, and determined their concentrations by ICP-MS or ICP-OES. Other chelating resins containing nitrogen and oxygen as donor atoms have also been developed. Blasius and Brozio [27,28] prepared a resin based on pyridine-2,6-dicarboxylic acid, and reported on its use for the separation of the alkaline-earth elements. Inoue et al. [29] synthesized a resin having nitrilotriacetate as the functional group [shown in Fig. 15.4 (b)], and applied the resin to the separation of rare earth elements. Colella et al. [30,31] synthesized a poly(acrylamidoxime) resin (Fig. 15.4 (c)) which was used to concentrate Fe(III), Cu(II), Cd(II), Pb(II), and Zn(II) from sea water and pond water. Chelating resins containing heterocyclic amine and amidoxime groups, called POLYORG were developed by Myasoedova et al. [32]. These resins were applied to the preconcentration of noble metals from rocks, ores, and industrial products [33]. Chelating resins containing sulfur as donor atoms have also been reported. Figure 15.4 (d) shows the structure of the dithiocarbamate functional group, which has two sulfur atoms as coordinating sites; thus soft metal ions are adsorbed well by the resin. Yamagami et al. [34] preconcentrated Hg from natural water whose pH was adjusted to 2–3 using this resin. Su et al. [35] synthesized a poly(vinylthiopropionamide) resin (Fig. 15.4 (e)), and reported that Au, Pt, Pb, and Ir were retained with up to 90% efﬁciency by this resin. Other chelating resins with different functional groups have been developed; their information can be found in the literature [14,15].

15.4

APPLICATION OF CHELATION TO SAMPLE PREPARATION FOR TRACE METAL ANALYSIS

Various methods of sample preparation for trace metal analysis based on chelation have been reported. Several examples are listed in Tables 15.4 –15.9. In this section, some examples with concrete experimental procedures for the separation and preconcentration of metal ions using chelate extraction and chelating resin techniques are presented. 15.4.1 Procedure for the extraction of metal ions from natural waters 15.4.1.1 Preconcentration and separation of Cd, Cu, Fe, Mo, Ni, V, and Zn from sea water by ammonium pyrrolidine dithiocarbamate (APDC) – diethylammonium diethyldithiocarbamate (DDDC) extraction and determination by ICP-OES [43] Pretreatment of sea water: sea water is acidiﬁed on collection to pH ca. 2 by the addition of 1 cm3 of nitric acid to 1 dm3 of sea water.

477

478 TABLE 15.4 Chelate extraction of trace elements from water samples Trace elements

Chelating agents

Organic solvents

Determination techniques

References

Sea water

Mn Cd, Co, Cu, Ni, Zn Au, Cu, Hg Cr Ag, Cd, Cr, Cu, Fe, Ni, Pb, Zn Au, Pt Cd, Co, Cu, Fe, Mo, Ni, Pb, V, Zn Cr Cd, Cu, Fe, Mn, Pb, Zn Cd, Co, Cr, Cu, Fe, Mo, Ni, Pb, V, Zn Cd, Co, Cu, Mn, Pb, U, Zn Cd, Co, Cu, Fe, Mn, Ni, Zn

8-Quinolinol Dithizone DDTC APDC APDC

Chloroform Chloroform Chloroform MIBK MIBK

GF-AAS AAS NAA AAS GFAAS

[36] [37,38] [39] [40] [41]

APDC APDC þ DDDC

MIBK Freon TF, chloroform

GFAAS, ICP-MS AAS, ICP-OES

[42] [43,44]

Diphenylcarbazide DDTC APDC

Isoamyl alcohol Isoamyl alcohol DIBK

GFAAS AAS AAS

[45] [46] [47]

DDTC

Chloroform

AAS

[48]

APDC

MIBK

AAS

[49]

River water

Drinking water

DDTC: diethyldithiocarbamate, APDC: ammonium pyrrolidine dithiocarbamate, DDDC: diethylammonium diethyldithiocarbamate, MIBK: methyl isobutyl ketone, DIBK: diisobutyl ketone, Freon TF: 1,1,2-trichloro-1,2,2-triﬂuoroethane, AAS: ﬂame atomic absorption spectrometry, NAA: neutron activation analysis, GFAAS: graphite furnace atomic absorption spectrometry, ICP-MS: inductively coupled plasma-mass spectrometry, ICP-OES: inductively coupled plasma-optical emission spectrometry.

H. Itabashi and T. Nakahara

Samples

TABLE 15.5 Extraction of trace elements from water samples by chelating resins Trace elements

Functional groups

Determination techniques

References

Sea water

Al, Cd, Co, Cr, Cu, Fe, Mn, Mo, Ni, Pb, Ti, V, Zn Cd, Cu, Fe, Pb, Zn Cd, Co, Cr, Cu, Fe, Mn, Ni, Pb, Zn Hg Al, Cd, Co, Cu, Fe, Mn, Mo, Ni, Pb, Ti, V, Y, Zn Al, As, Bi, Ce, Cd, Co, Cr, Cu, Dy, Er, Eu, Fe, Gd, Ho, La, Lu, Mn, Mo, Nd, Ni, Pb, Pr, Sb, Se, Sm, Sn, Tb, Ti, Tm, U, V, W, Yb, Zn Ce, Dy, Er, Eu, Gd, Ho, La, Lu, Nd, Pr, Sm, Tb, Tm, Y, Yb Cd, Co, Cu, Pb Ga, In, Ti Cd, Cu, Mn, Ni, Pb Al, Bi, Cd, Co, Cr, Cu, Mn, Ni, Pb, U, V, Zn Cd, Cu, Pb, Zn

Iminodiacetate

ICP-OES

[17]

Acrylamidoxime Iminodiacetate

[31] [50,51]

Dithiocarbamate Iminodiacetate

AAS GF-AAS, ICP-OES, ID-SS-MS AAS ICP-OES

Iminodiacetate

ICP-MS, ICP-OES

[22,25,26]

Iminodiacetate

ICP-MS

[19]

Iminodiacetate, isocyanate 8-Hydroxyquinolinate Iminodiacetate Iminodiacetate

ICP-OES ICP-MS ICP-MS ICP-MS

[53] [54] [55] [56]

Iminodiacetate

AAS

[31]

Sea water

Pond water

[34,52] [18]

479

continued

Chelation solvent extraction for separation of metal ions

Samples

480 TABLE 15.5 (continuation) Samples

River water

Functional groups

Determination techniques

References

Al, Cd, Ce, Cu, Fe, La, Mn, Mo, Ni, Pb, Pr, Sn, Ti, U, V, Y, Zn Al, Ba, Ce, Cd, Co, Cu, Dy, Fe, Er, Eu, Gd, La, Lu, Ho, Mn, Mo, Nd, Ni, Pb, Pr, Rb, Sb, Sm, Sn, Sr, Tb, Ti, Tm, U, V, Y, Yb, Zn, Zr Hg Sb Ag, Al, As, Ba, Cd, Co, Cr, Cs, Cu, Fe, Ga, Li, Mo, Mn, Ni, Pb, Pd, Pr, Rb, Sb, Se, Sn, Sr, Te, Ti, V, W, U, Zn, Zr

Iminodiacetate

ICP-MS

[20]

Iminodiacetate

ICP-MS

[21]

Dithiocarbamate Iminodiacetate Iminodiacetate

AAS AAS ICP-MS

[34,52] [57] [58]

ICP-OES: inductively coupled plasma-optical emission spectrometry, AAS: ﬂame atomic absorption spectrometry, GFAAS: graphite furnace atomic absorption spectrometry, IDSSMS: isotope dilution spark source mass spectrometry, ICP-MS: inductively coupled plasma-mass spectrometry.

H. Itabashi and T. Nakahara

Lake water

Trace elements

TABLE 15.6 Chelate extraction of trace elements from high-purity materials and other inorganic solid samples Trace elements

Chelating agents

Organic solvents

Determination techniques

References

(NH4)2HPO4 Al metal U compounds W and its oxide

Ca Ga Ag, Cu, Hg Co, Cu, Ni, Pb, Zn Co, Cr, Cu, Fe, Mn, Ni Co, Cu, Ni Li La U Ti

8-Quinolinol 8-Quinolinethiol Dithizone Dithizone

3-Methyl-1-butanol MIBK Carbon tetrachloride Chloroform

AAS Fluor. Phot. XRF

[59] [60] [61] [62]

DDTC

MIBK

AAS

[63]

DDTC DPM NHDTAHA HPBI Cupferron

MIBK Ethyl ether Chloroform DIBK MIBK

AAS Phot. ICP-OES ICP-OES AAS

[64] [65] [66] [67] [68]

In Cu, Ni, Zn

DDTC APDC

n-Butyl acetate MIBK

AAS, OES AAS

[69] [70]

Glasses, Na2CO3, CaCO3 Rocks Be and its oxide Monazite sand Apatite minerals Ti in ferric oxides, Ni–Cr steel, Ni alloy, steels Al alloy Pd compounds

481

MIBK: methyl isobutyl ketone, DIBK: diisobutyl ketone, AAS: ﬂame atomic absorption spectrometry, Fluor.: ﬂuorometric analysis, XRF: X-ray ﬂuorescence analysis, Phot.: photometric analysis, ICP-OES: inductively coupled plasma-optical emission spectrometry, DDDC: diethylammonium diethyldithiocarbamate, DDTC: diethyldithiocarbamate, DPM: dipivaloylmethane, NHDTAHA: 5,14N,N0 -hydroxyphenyl-4,15-dioxo-1,5,14,18-tetraaza hexacosane, HPBI: 3-phenyl-4-benzoyl-5-isoxazolone, APDC: ammonium pyrrolidine dithiocarbamate.

Chelation solvent extraction for separation of metal ions

Matrices

482 TABLE 15.7

Samples

Trace elements

Functional groups

Determination techniques

References

Cu alloy and Ni sponge Graphite Ni alloy Rock, soil Rock Pond sediment

Se Al, Cu, Fe, Si, Ti, V Au, Ru, Ti, V U Cd, Cu, Ni, Zn Cd, Cu

Iminodiacetate Dithiocarbamate Epoxy-melamine 4-(2-Thiazolylazo)resorcinol Iminodiacetate Iminodiacetate

AAS ICP-OES ICP-OES ICP-OES AAS AAS

[71] [72] [73] [74] [75] [76]

AAS: ﬂame atomic absorption spectrometry, ICP-OES: inductively coupled plasma-optical emission spectrometry.

H. Itabashi and T. Nakahara

Extraction of trace elements from high-purity materials and other inorganic solid samples by chelating resins

TABLE 15.8 Chelate extraction of trace elements from biological samples Trace elements

Chelating agents

Organic solvents

Determination techniques

References

Barley, grass, straw, rice ﬂour Pine needles, birch leaves Orchard Leaves

Co

2-Nitroso-1-naphthol

Heptan-2-one

GF-AAS

[77]

Mo

MIBK, DIBK, isoamyl alcohol Xylene

AAS, GF-AAS

[78]

Cd, Co, Cu, Fe, Mn, Mo, Ni, Pb, V, Zn Se

8-Quinolinol, toluene-3,4-dithiol APDC, HMAHMDC

ICP-OES

[79]

DDDC

Chloroform

GF-AAS

[80]

Gd

TTA, PMBP

MIBK

GF-AAS

[81]

Cd

DPTH

MIBK

GF-AAS

[82]

Alfalfa, hevea leaves, tuna ﬁsh, mussel homogenate, ﬁsh ﬂesh homogenate, animal muscle, bovine liver, wheat ﬂour Kidney, muscle, liver, lung, spleen, brain, serum, urine, heart Olive leaves, pig kidney, bovine muscle, dogﬁsh liver, lobster hepatopancreas, dogﬁsh muscle, bovine liver, human urine

continued

Chelation solvent extraction for separation of metal ions

Samples

483

484 TABLE 15.8 (continuation) Trace elements

Chelating agents

Organic solvents

Determination techniques

References

Orchard leaves, citrus leaves, tomato leaves, serum, urine Blood

Bi, Cd, Cr, Fe, Hg, Ni, Pb, Se, Zn

APDC

AAS, NAA

[84–86], [85]

Cd, Ni, Pb, Tl Bi, Cd, Co, Cu, Fe, Li, Mn, Ni, Pb, Sr, Zn

DDTC, APDC Cupferron, DDDC, APDC, 8-quinolinol, TTA Dithizone

Chloroform, MIBK, methyl-namyl ketone MIBK MIBK

AAS AAS

[83] [87]

Chloroform

AAS

[37]

Kelp, whale heart, whale fat, trout

Ag, Cd, Co, Cu, Ni, Pb, Zn

APDC: ammonium pyrrolidine dithiocarbamate, HMAHMDC: hexamethyleneammonium hexamethyldithiocarbamate, DDDC: diethylammonium diethyldithiocarbamate, TTA: thenoyltriﬂuoroacetone, TBP: tributyl phosphate, PMBP: 4-benzoyl-3-methyl-1phenyl-2-pyrazolin-5-one, DPTH: 1,5-bis(di-2-pyridylmethylene)thiocarbohydrazine, DDTC: diethyldithiocarbamate, MIBK: methyl isobutyl ketone, DIBK: diisobutyl ketone, GFAAS: graphite furnace atomic absorption spectrometry, AAS: ﬂame atomic absorption spectrometry, ICP-OES: inductively coupled plasma-optical emission spectrometry, NAA: neutron activation analysis.

H. Itabashi and T. Nakahara

Samples

Extraction of trace elements from biological samples by chelating resins Samples

Trace elements

Functional groups

Determination techniques

References

Pepperbush, kidney Pepperbush, mussel, tomato leaves Mussels Blood serum

Cu, Pb Cd, Cu

Iminodiacetate Iminodiacetate

ICP-OES AAS

[88] [76]

Cd Ce, Cu, Dy, Er, Eu, Gd, Ho, La, Lu, Nd, Pr, Sm, Tb, Tm, Yb, Zn Cu, Fe, Ni, Pb, Zn Bi, Cd, Cu, Hg, Ni, Pb, Se, Sn, Te, U Cd, Co, Cu, Pb

Aminophosphonate Iminodiacetate

AAS ICP-MS

[89] [23,24]

Iminodiacetate Dithiocarbamate

ICP-MS ICP-OES

[90] [91]

Iminodiacetate, isocyanate

ICP-MS

[53]

Urine

ICP-OES: inductively coupled plasma-optical emission spectrometry, AAS: ﬂame atomic absorption spectrometry, ICP-MS: inductively coupled plasma-mass spectrometry.

Chelation solvent extraction for separation of metal ions

TABLE 15.9

485

H. Itabashi and T. Nakahara

Preparation of APDC – DDTC solution: APDC (0.5 g) and DDDC (0.5 g) are taken in a 50 cm3 measuring cylinder and dissolved in 50 cm3 of water. The solution is then transferred to a 250 cm3 separatory funnel and extracted using 10 cm3 of puriﬁed chloroform. The procedure is repeated a further ﬁve times using fresh chloroform. The puriﬁed extracting reagent is stored in a 100 cm3 PTFE bottle. Extraction procedure: 500 g of the acidiﬁed sea water sample (pH ca. 2) is taken in a 500 cm3 separatory funnel, and 1.4 cm3 of acetate buffer, 2 cm3 of the APDC –DDTC solution, and 10 cm3 of chloroform are added. The mixture is shaken for 3 min and the phases are allowed to separate for 5 min. The organic phase is transferred to a 125 cm3 separatory funnel. An additional 10 cm3 of chloroform is added into the 500 cm3 separatory funnel, and the solution is shaken for 3 min. The phases are allowed to stand for 5 min and the organic phase is then transferred to the 125 cm3 separatory funnel. Nitric acid (0.20 cm3) is added to the organic phase, the mixture is shaken for 1 min and allowed to stand for 5 min. High-purity water (about 2 cm3) is added to the 125 cm3 separatory funnel and the mixture is shaken for 1 min. After the phases are allowed to separate for 5 min, the organic phase is discarded and the aqueous phase is transferred to a 4 cm3 PTFE beaker. The separatory funnel is rinsed with about 2 cm3 of high-purity water, the washing solution is added to the PTFE beaker. The extract is evaporated to dryness using an infrared lamp in a clean-air bench and the residue is dissolved in 0.3 cm3 of nitric acid. The solution is heated under the infrared lamp to a small volume (about 0.05 cm3), and water is added (exactly 1.0 or 2.0 cm3). The solution is then supplied directly to ICP-OES from the PTFE beaker. Other information: detection limits (ng dm23): 2 for Cd, Cu, and Zn; 6 for Fe and V; 12 for Mo; 16 for Ni. Extraction pH 4 is recommended for the extraction of V and Mo. Element recoveries are almost 100%, except for Mo (92%). 15.4.1.2 Preconcentration and separation of Cr(VI) from river, estuarine, sea, and drinking water using diphenylcarbazide and determination by graphite furnace atomic absorption spectrometry (GF-AAS) [45] Extraction procedure: 25 cm3 of a water sample are taken into a 50 cm3 polypropylene screw capped tube. Sulfuric acid (5 mol dm23), 0.25 cm3, and 0.5 cm3 acetone containing 1% (w/v) diphenylcarbazide are added to the tube. The sample is swirled to mix and left for 10 min. Then 20 cm3 of a saturated sodium chloride solution is added, followed by 2.5 cm3 of isoamyl alcohol. The tube is capped and shaken for 4 min. After the phases are allowed to separate, the upper alcohol phase is analyzed by GFAAS. Other information: detection limit: 0.024 mg dm23, analytical range: 0–2 mg dm23. The use of standard additions calibration is recommended for the measurement of the samples which contain an abundance of humic substances.

486

Chelation solvent extraction for separation of metal ions

15.4.1.3 Preconcentration and separation of 28 trace metals from sea water using poly(iminodiacetate) resin and determination by ICP-MS [26] Pretreatment of sea water: a seawater sample is ﬁltered with a membrane ﬁlter (pore size of 0.45 mm) immediately after sampling. The ﬁltered sample is acidiﬁed to pH ca. 1 by adding nitric acid. Preconcentration procedure: 250 cm3 of the acidiﬁed sample solution is taken in a glass beaker and the pH of the sample is adjusted to 6 using acetic acid and ammonium hydroxide solutions. After 0.2 g (dry weight) of the chelating resin (Chelex-100) is added to the sample solution, the mixture is stirred for 2 h with a magnetic stirrer. The chelating resin is then collected on a glass ﬁlter (G4) by ﬁltration, and rinsed with 8 cm3 of 1 mol dm23 ammonium acetate solution to release Mg and Ca partly adsorbed on the resin. Finally, analyte metals (Al, V, Mn, Fe, Co, Ni, Cu, Zn, Y, Mo, Cd, La, Ce, Pr, Nd, Sm, Eu, Gd, Tb, Dy, Ho, Er, Tm, Yb, Lu, W, Pb, U) adsorbed on the chelating resin are eluted with 6 cm3 of a 2 mol dm23 nitric acid solution, into which 0.5 cm3 of a mixed solution of Ga, In, Re, and Tl (100 mg dm23 each) is added as internalstandard elements to correct for matrix effects. The solution containing the analyte metals is analyzed by ICP-MS. Other information: detection limit (ng dm23): over the range from 20 for Ti to 0.006 for Ho (see Ref. [26] for each element). Fe is quantitated by ICP-OES. 15.4.1.4 Preconcentration and separation of Hg from river water using poly(dithiocarbamate) resin and determination by cold-vapor AAS [52] Pretreatment of river water: river water is collected in a 20 dm3 high-density polyethylene bottle. The sample is adjusted to pH 2 with nitric acid, and 1 mg of HAuCl4 is added as a preservative. Preconcentration and measurement procedure: the wet dithiocarbamate resin (Sumichelate Q-10, 20 –50 mesh, Sumitomo Chemicals) is packed into a column (15 mm i.d., 5 cm long). The sample solution is passed through the column at ca. 30 cm3 min21 to collect (adsorb) Hg onto the resin. After the collection, 30 cm3 of an acidic 5% thiourea solution is passed through the column to elute the adsorbed Hg. Inorganic and total mercury are determined separately in two 10-cm3 aliquots of this efﬂuent. For the determination of inorganic mercury, 10 cm3 aliquot of the efﬂuent is placed in a reaction vessel in a vapor generation system, 10 cm3 of 30% (w/v) potassium hydroxide is added followed by 2 cm3 of a tin(II) chloride solution (10% w/v), and a carrier air ﬂow is immediately started. The mixture is allowed to react for 30 s, during which time the mercury vapor generated is passed through a quartz gas cell of an AAS spectrometer. The peak height response is used for measurement. For the determination of total mercury, the same procedure is used, except that a tin(II) chloride –cadmium chloride mixture (10–1%), instead of the tin(II) chloride alone, is used as the reductant.

487

H. Itabashi and T. Nakahara

Other information: the range of determination: 0.2 –5000 ng dm23. Recovery: more than 95%. 15.4.2 Procedure for the extraction of metal ions from high-purity materials and inorganic solid samples 15.4.2.1 Separation of In in Al alloys by DDTC extraction and determination by AAS and OES [69] Dissolution procedure for Al alloy: a 0.2 –0.5 g sample is placed in a covered 400 cm3 beaker and heated with 20 cm3 of 6 mol dm23 hydrochloric acid until completely dissolved. After cooling, the pH of the solution is adjusted to 3–3.5 using a sodium hydroxide solution. Extraction procedure: the sample solution (pH 3–3.5) is placed in a 250 cm3 separatory funnel, and 5 cm3 of a 1% (w/v) sodium–DDTC solution is added. After dilution to 50 cm3 with pure water, 20 cm3 of methyl isobutyl ketone (MIBK) or n-butyl acetate is added to the funnel. The mixture is shaken for 1 min. After the phases are allowed to separate, the organic phase is analyzed by the AAS or OES. Other information: detection limit (mg dm23): 0.10 for AAS, 0.08 for OES. Minimum time for which complex is stable: 12 h for MIBK as solvent, 72 h for nbutyl acetate as solvent. Extractability: 95% for MIBK as solvent, 99% for nbutyl acetate as solvent. 15.4.2.2 Separation of Co, Cr, Cu, Fe, Mn, Ni in high-purity glasses by DDTC extraction and determination by GF-AAS [63] Dissolution procedure for glasses: glass samples are placed in a platinum dish and heated in a furnace at 800 –9008C for 0.5 –2 min. To shatter them, a few (cm3) of distilled water is dropped onto the pieces of glass. The shattered samples are ground to ﬁne powder using an agate pestle and mortar. One gram of the sample is placed in a platinum crucible, and 4 cm3 of hydrochloric acid and 5 cm3 of perchloric acid are added. The crucible is heated gently to dryness. The residue is dissolved in a mixture of 0.1 cm3 of perchloric acid and 4 cm3 of water. Extraction procedure: the sample solution is transferred to a 10 cm3 volumetric ﬂask, and 5 cm3 of a solution containing 5% (w/v) Na –DDTC and 10% (w/v) sodium acetate is added (the solution is buffered at pH 6). After the ﬂask is allowed to stand for 15 min, 1 cm3 of MIBK is added. The mixture is shaken thoroughly for 3 min and the phases are allowed to separate; the organic phase is analyzed by GFAAS. Other information: extraction efﬁciency: about 100% except Cr. Chromium results in poor extraction, because of losses as chromyl chloride and/or ﬂuoride during the dissolution stage. Each element can be measured down to 0.01 mg g21.

488

Chelation solvent extraction for separation of metal ions

15.4.2.3 Separation of U from apatite minerals by 3-phenyl-4-benzoyl-5-isooxa zolone (HPBI) extraction and determination by ICP-OES [67] Dissolution procedure for apatite: 1–10 g of a powdered sample in a covered 100 cm3 beaker is heated with 10 cm3 of nitric acid at 150 –2008C for 30– 60 min. After cooling, the sample solution is diluted with pure water, passed through a ﬁlter paper (No. 5c) to remove any residue and then diluted to 100 cm3 (10–100 mg sample cm23) with pure water. Extraction procedure: the apatite sample solution (apatite 0.1 –0.2 g) is placed in a separatory funnel and the pH is adjusted to 0.3. Five cubic centimeters of 0.06 mol dm23 HPBI in diisobutyl ketone is added to the funnel, and the mixture is shaken for 10–20 min. The organic phase is transferred to a centrifuge tube and separated. The centrifuged organic phase is analyzed by ICP-OES. Other information: detection limit: 0.02 mg dm23. Recovery of U is about 100% at pH . 0. When the content of the apatite mineral is higher than 0.8 g/10 cm3, recovery decreases with increase in the apatite content. 15.4.2.4 Removal of Fe matrix from dissolved steel sample by di(2-ethylhexyl) phosphate (D2EHPA) extraction [92] Extraction procedure: 20 cm3 of a dissolved Fe sample solution ([Fe] , 0.1 mol dm23, pH ¼ 2.0) are taken in a 100 cm3 separatory funnel and 20 cm3 of MIBK containing 1 mol dm23 D2EHPA is added. The mixture is shaken for 4 min, and then 2 cm3 of a sodium hydroxide solution (ca. 2 mol dm23) are added to the funnel in order to adjust the pH to 2. The mixture is shaken for 4 min. After the phases are allowed to separate, the aqueous phase is transferred to another 100 cm3 separatory funnel (the organic phase is discarded). Twenty cubic centimeters of the MIBK solution containing 1 mol dm23 D2EHPA are added to the aqueous phase, and the mixture is shaken for 4 min. After the phases are allowed to separate, the aqueous phase (containing almost no Fe) is used for analysis. Other information: a ﬂow system using this extraction method has recently been developed [93]. 15.4.2.5 Separation of Al, Cu, Fe, Si, Ti, V from high-purity graphite using poly(dithiocarbamate) or poly(acrylamidoxime) resins and determination by ICP-OES [72] Dissolution procedure for high-purity graphite: a graphite sample is dried in an oven at 1008C for 2 h. One to ﬁve grams of the graphite sample is placed in a platinum crucible, and 2 cm3 of a 5% magnesium nitrate solution are added as an ashing aid, especially to prevent volatilization loss of V. Prior to its use, the magnesium nitrate solution was puriﬁed by shaking for 24 h with 500 mg of the poly(dithiocarbamate) resin. The graphite sample is then dried on a hot plate and kept in a mufﬂe furnace at 8008C for 12 h until ashing is complete.

489

H. Itabashi and T. Nakahara

The ash is dissolved in 10 cm3 of hydrochloric and nitric acids (3:1) and diluted with distilled water to a ﬁnal volume of 100 cm3. Separation procedure for Cu, Fe, Si, Ti, V: pH of the sample solution is increased to 5 (for Cu, Fe, Ti, V) or 8 (for Si) with ammonium hydroxide, and the solution is passed through a column containing 70 mg of 70 – 80 mesh poly(dithiocarbamate) resin. The resin is initially digested with 30% hydrogen peroxide at room temperature for 5 min followed by slow heating in 2 cm3 of nitric acid at 1008C for 15–20 min. The ﬁnal volume (5 cm3) is adjusted using 0.1 mol dm23 nitric acid. This solution is analyzed by ICP-OES. Separation procedure for Al: the pH of the sample solution is adjusted to 6 using ammonium hydroxide. The poly(acrylamidoxime) resin (100 mg) is added, and the mixture is stirred for 10 h. The mixture is then ﬁltered and the resin is washed with pure water. The resin is added to 2 cm3 of a mixture of 20% hydrochloric and 20% nitric acid, and the mixture is shaken for 3 h. The mixture is ﬁltered and the resin is washed with pure water. The ﬁltrate is collected and diluted to 5 cm3, which is used as a sample for ICP-OES analysis. Other information: detection limit (ng g21 in graphite): 8.4 for Al, 5.0 for Cu, 8.0 for Fe, 20 for Si, 4.0 for Ti, 8.0 for V. Recovery: 96– 103%. 15.4.3 Procedure for the extraction of metal ions from biological samples 15.4.3.1 Separation of Ag, Cd, Co, Cu, Ni, Pb, Zn from biological tissue by dithizone extraction and determination by AAS [37] Preparation of tissue sample: a sample is freeze-dried and ground in a ball mill and a 0.5 –3 g subsample is weighed into a 100 cm3 conical silica ﬂask. Nitric acid (10–20 cm3) is added to the ﬂask and the sample is left covered on a sand bath at ca. 1008C for 3 h. The cover is removed and the solution is evaporated to dryness. If the solution is still colored when evaporated, the treatment is repeated with 5–10 cm3 of nitric acid. When the solution is nearly colorless, 5 cm3 of (1 þ 1) nitric-perchloric acid are added and the solution is evaporated to dryness. Fifty cubic centimeters of water and 10 cm3 of hydrochloric acid are added and the solution is heated on a hot plate. If a residue remains, hydrochloric acid is added until a clear solution results. Three cubic centimeters of a 50% (w/v) citric acid solution are added and the pH of the solution is adjusted to 8 by adding ammonia. Extraction procedure: the sample solution (pH ¼ 8) is placed in a 150 cm3 separatory funnel and 5 cm3 of a chloroform solution containing 0.2% (w/v) dithizone are added. The mixture is shaken vigorously for 2 min and then the organic phase is transferred to a 100 cm3 separatory funnel. The pH of the aqueous phase is adjusted to 9.5 with ammonia and the extraction is repeated with 5 cm3 of a 0.2% dithizone solution. Finally, the extraction is performed using a 0.02% dithizone solution. If the dithizone still changes color, the extraction is repeated with the 0.02% dithizone solution until it does not.

490

Chelation solvent extraction for separation of metal ions

The dithizone extracts are combined in a 100 cm3 separatory funnel and washed with dilute ammonia. After the phases are allowed to separate, the organic phase is transferred into a second 100 cm3 separatory funnel and 50 cm3 of 0.2 mol dm23 hydrochloric acid are added. The mixture is shaken vigorously for 2 min. After the phases are allowed to separate, the aqueous phase is transferred to a beaker and evaporated to dryness. The residue is dissolved in 5 cm3 of 2 mol dm23 hydrochloric acid. The solution is analyzed by AAS for determination of Cd, Pb, and Zn. Conversely, the organic phase is transferred to another beaker, and 3 cm3 of perchloric acid is added. The organic phase is evaporated to dryness, and then 2 cm3 of 60% perchloric acid are added. The solution is evaporated to dryness, and the residue is dissolved in 5 cm3 of 2 mol dm23 hydrochloric acid. The solution is analyzed by AAS for determination of Ag, Co, Cu, and Ni. 15.4.3.2 Separation of rare earth elements from human blood serum using poly(iminodiacetate) resin and determination by ICP-MS [23] Preparation of human blood serum sample: 8 cm3 of a blood serum sample are placed in a 100 cm3 Teﬂon beaker and heated, after adding 2 cm3 of nitric acid, almost to dryness on a hot plate at 1108C. Then 2 cm3 of nitric acid are again added to the residue and the solution is heated at 1508C for 2 h. After adding a further 2 cm3 of nitric acid and 1 cm3 of 60% perchloric acid, the solution is heated at 1508C for 4 h until white fumes appear. This procedure is repeated twice. Finally, 0.76 cm3 of nitric acid and ca. 1 cm3 of pure water are added to dissolve the residue with heating at 1108C for 1 h, and the solution is diluted to 100 cm3 with pure water. Separation procedure: 100 cm3 of the sample solution is diluted to 300 cm3 with pure water. The pH of the solution is adjusted to 6 by adding either acetic acid or ammonia solutions, and then 0.5 g of the poly(iminodiacetate) resin (Chelex-100) is added to the solution. After stirring the solution at 808C for 3 h, the resin is collected on a glass ﬁlter and rinsed carefully with 10 cm3 of a 1 mol dm23 ammonium acetate solution and 2 cm3 of pure water. The rare earth elements adsorbed on the resin are eluted with 10 cm3 of 2 mol dm23 nitric acid. The efﬂuent is evaporated to dryness and the residue is dissolved in 2 cm3 of 0.1 mol dm23 nitric acid in which Rh and Re have been added as internal standard elements. The solution is analyzed by ICP-MS. Other information: detection limit (ng dm23): 0.2 for Ce, Sm, Gd; 0.1 for Nd, Dy; 0.07 for La, Yb; 0.05 for Eu; 0.04 for Pr, Ho, Er; 0.03 for Tm, Lu; 0.02 for Tb. Recovery: 92–102%. 15.4.4 Procedure for the speciation of metal ions in natural waters 15.4.4.1 Speciation of Cu(II) in river water samples by TTA extraction [11] Experimental procedure: 1 cm3 of a 5 mol dm23 sodium perchlorate aqueous solution is added to 50 cm3 of a water sample solution in a 100 cm3 separatory

491

H. Itabashi and T. Nakahara

funnel. Then, 5 cm3 of a carbon tetrachloride solution containing 4.0 – 12.0 £ 1024 mol dm23 TTA are added and the mixture is shaken vigorously for 30 min. After the phases are allowed to separate, the organic phase is analyzed by GFAAS and the pH of the aqueous phase is measured. According to the plot of Eq. (15.37), aL and M(mol) values are calculated, where Kex ¼ 0:10 is used. 15.4.4.2 Estimation of ligand concentrations and stability constants of Cu(II) complexes by TTA extraction [12] Preparation of Cu(II) – TTA complex: 50 cm3 of an ethanol solution containing 0.04 mol TTA are added to a 100 cm3 aqueous solution containing 0.02 mol copper(II) acetate. The resulting mixture is agitated for 30 min with a magnetic stirrer. After having stood for 3 h in a refrigerator, the crude product is washed with water and cold ethanol. Finally, Cu(tta)2 is obtained by recrystallization from ethanol. Extraction procedure: water samples are ﬁltered through a 0.45 mm membrane ﬁlter. Twenty cubic centimeters of the sample solution containing 0.1 mol dm23 sodium perchlorate, whose pH is preliminarily adjusted to 7 by a 0.01 mol dm23 3-morpholinopropanesulfonic acid buffer solution, are taken in a 100 cm3 separatory funnel, and an equal volume of benzene solution containing 0–5.0 £ 1023 mol dm23 TTA and 5.0 £ 1025 mol dm23 Cu(tta)2 are added. The mixture is then shaken vigorously for 1 h. After the phases are allowed to separate, Cu(II) in the aqueous phase is determined by AAS or GFAAS, followed by a pH measurement. According to Eq. (15.45), Cu(II) concentrations in the aqueous phase (corresponding to [ML] in Eq. (15.45)) are plotted as a function of log x, and concentrations of the ligands, Li , and conditional stability constants of the copper(II) complexes, bLi ð¼ ½CuLi ½Cu2þ 21 ½Li 21 Þ, are calculated using a nonlinear regression analysis.

REFERENCES 1

2 3 4 5 6 7 8 9

492

N. Suzuki, K. Akiba, K. Saitoh, Y. Hasegawa, H. Watarai, M. Kato and S. Tsukahara, Database on Solvent Extraction of Metal Ions, http://sedatant. chem.sci.osaka-u.ac.jp. H. Itabashi, Y. Takazawa, N. Niibe and H. Kawamoto, Anal. Sci., 13 (1997) 921. H. Itabashi, M. Yoshida and H. Kawamoto, Anal. Sci., 17 (2001) 1301. R.G. Pearson, J. Chem. Ed., 45 (1968) 581. E.B. Sandell and H. Onishi, Photometric Determination of Traces of Metals, Part 1. John Wiley & Sons, New York, 1978. T.V. Healy, Nucl. Sci. Eng., 16 (1963) 413. T. Sekine and Y. Hasegawa, Solvent Extraction Chemistry. Marcel Dekker, New York, 1977. H. Irving and D.N. Edgington, J. Inorg. Nucl. Chem., 15 (1960) 158. A. Ringbom, Complexation in Analytical Chemistry. John Wiley & Sons, New York, 1963, p. 353.

Chelation solvent extraction for separation of metal ions 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50

A. Tessier and D.R. Turner, Metal Speciation and Bioavailability in Aquatic Systems. John Wiley & Sons, Chichester, 1995. H. Itabashi, H. Kawamoto and H. Akaiwa, Anal. Sci., 10 (1994) 341. H. Itabashi, Y. Shigeta, H. Kawamoto and H. Akaiwa, Anal. Sci., 16 (2000) 1179. H. Itabashi, D. Yamazaki, H. Kawamoto and H. Akaiwa, Anal. Sci., 17 (2001) 785. Z.B. Alfassi and C.M. Wai, Preconcentration Techniques for Trace Elements. CRC Press, Florida, 1992. B.M. Marhol, Ion Exchangers in Analytical Chemistry. Their Properties and Use in Inorganic Chemistry. Elsevier, Amsterdam, 1982. S. Pai, Anal. Chim. Acta, 211 (1988) 271. C.J. Cheng, T. Akagi and H. Haraguchi, Bull. Chem. Soc. Jpn, 58 (1985) 3229. C.J. Cheng, T. Akagi and H. Haraguchi, Anal. Chim. Acta, 198 (1987) 173. H. Sawatari, T. Toda, T. Saizuka, C. Kimata, A. Itoh and H. Haraguchi, Bull. Chem. Soc. Jpn, 68 (1995) 3065. A. Itoh, C. Kimata, H. Miwa, H. Sawatari and H. Haraguchi, Bull. Chem. Soc. Jpn, 69 (1996) 3469. H. Haraguchi, A. Itoh, C. Kimata and H. Miwa, Analyst, 123 (1998) 773. T. Yabutani, F. Mouri and H. Haraguchi, Anal. Sci., 16 (2000) 675. K. Inagaki and H. Haraguchi, Analyst, 125 (2000) 191. K. Inagaki, N. Mikuriya, S. Morita, H. Haraguchi, Y. Nakahara, M. Hattori, T. Kinoshita and H. Saito, Analyst, 125 (2000) 197. T. Yabutani, K. Chiba and H. Haraguchi, Bull. Chem. Soc. Jpn, 74 (2001) 31. T. Yabutani, F. Mouri, A. Itoh and H. Haraguchi, Anal. Sci., 17 (2001) 399. E. Blasius and B. Brozio, Z. Anal. Chem., 192 (1963) 364. E. Blasius and B. Brozio, J. Chromatogr., 18 (1965) 572. Y. Inoue, H. Kumagai, Y. Shimomura, T. Yokoyama and T.M. Suzuki, Anal. Chem., 68 (1996) 1517. M.B. Colella, S. Siggia and R.M. Barnes, Anal. Chem., 52 (1980) 967. M.B. Colella, S. Siggia and R.M. Barnes, Anal. Chem., 52 (1980) 2347. G.V. Myasoedova, I.I. Antokol’skaya and S.B. Savvin, Talanta, 32 (1985) 1105. G.V. Myasoedova, Fresenius’ J. Anal. Chem., 341 (1991) 586. E. Yamagami, S. Tateishi and A. Hashimoto, Analyst, 105 (1980) 491. Z. Su, X. Chang, K. Xu, X. Luo and G. Zhan, Anal. Chim. Acta, 268 (1992) 323. G.P. Klinkhammer, Anal. Chem., 52 (1980) 117. H. Armannsson, Anal. Chim. Acta, 110 (1979) 21. R.G. Smith, Jr. and H.L. Windom, Anal. Chim. Acta, 113 (1980) 39. J.M. Lo, J.C. Wei and S.J. Yeh, Anal. Chem., 49 (1977) 1146. T.R. Gilbert and A.M. Clay, Anal. Chim. Acta, 67 (1973) 289. T.K. Jan and D.R. Young, Anal. Chem., 50 (1978) 1250. S.A. Wood and D. Vlassopoulos, Anal. Chim. Acta, 229 (1990) 227. C.W. McLeod, A. Otsuka, K. Okamoto, H. Haraguchi and K. Fuwa, Analyst, 106 (1981) 419. L. Danielsson, B. Magnusson and S. Westerlund, Anal. Chim. Acta, 98 (1978) 47. M. Gardner and S. Comber, Analyst, 127 (2002) 153. T.N. Tweeten and J.W. Knoeck, Anal. Chem., 48 (1976) 64. K.M. Bone and W.D. Hibbert, Anal. Chim. Acta, 107 (1979) 219. J. Korkisch and A. Sorio, Anal. Chim. Acta, 79 (1975) 207. K.M. Aldous, D.G. Mitchell and K.W. Jackson, Anal. Chem., 47 (1975) 1034. R.E. Sturgeon, S.S. Berman, J.A.H. Desaulniers, A.P. Mykytiuk, J.W. McLaren and D.S. Russell, Anal. Chem., 52 (1980) 1585.

493

H. Itabashi and T. Nakahara 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93

494

H.M. Kingston, I.L. Barnes, T.J. Brady and T.C. Rains, Anal. Chem., 50 (1978) 2064. K. Minagawa and Y. Takizawa, Anal. Chim. Acta, 115 (1980) 103. S. Caroli, A. Alimonti, F. Petrucci and Z. Horvath, Anal. Chim. Acta, 248 (1991) 241. K.J. Orians and E.A. Boyle, Anal. Chim. Acta, 282 (1993) 63. K.W. Warnken, D. Tang, G.A. Gill and P.H. Santschi, Anal. Chim. Acta, 423 (2000) 265. M. Nicolai, C. Rosin, N. Tousset and Y. Nicolai, Talanta, 50 (1999) 433. H. Narasaki, Anal. Sci., 2 (1986) 371. K. Lee, M. Oshima, T. Takayanagi and S. Motomizu, Anal. Sci., 16 (2000) 731. M. Yanagisawa, M. Suzuki and T. Takeuchi, Talanta, 14 (1967) 933. K. Watanabe and K. Kawagaki, Bull. Chem. Soc. Jpn, 48 (1975) 1812. J. Merecek and E. Singer, Fresenius Z. Anal. Chem., 203 (1964) 336. G.L. Hubbard and T.E. Green, Anal. Chem., 38 (1966) 428. C.W. Fuller and J. Whitehead, Anal. Chim. Acta, 68 (1974) 407. R.F. Sanzolone, T.T. Chao and G.L. Crenshaw, Anal. Chim. Acta, 105 (1979) 247. R.F. Apple and J.C. White, Talanta, 13 (1966) 43. Y.K. Agrawal and P. Shrivastav, Talanta, 44 (1997) 1307. O. Fujino, S. Umetani and M. Matsui, Anal. Chim. Acta, 296 (1994) 63. T. Nakahara, M. Munemori and S. Musha, Bull. Chem. Soc. Jpn, 46 (1973) 1127. A.J. Aller, Anal. Chim. Acta, 134 (1982) 293. S.A. Popova, S.P. Bratinova and C.R. Ivanova, Analyst, 116 (1991) 525. M. Ikeda, Anal. Chim. Acta, 170 (1985) 217. H.S. Mahanti and R.M. Barnes, Anal. Chem., 55 (1983) 403. X. Chang, B. Gong, Z.S.D. Yang and X. Luo, Fresenius J. Anal. Chem., 360 (1998) 736. C. Lee, M. Suh, J. Kim, D. Kim, W. Kim and T. Eom, Anal. Chim. Acta, 382 (1999) 199. T. Uchida, M. Nagase, I. Kojima and C. Iida, Anal. Chim. Acta, 94 (1977) 275. S. Hirata, K. Honda and T. Kumamaru, Anal. Chim. Acta, 221 (1989) 65. W.J. Blanchﬂower, A. Cannavan and D.G. Kennedy, Analyst, 115 (1990) 1323. L.H.J. Lajunen and A. Kubin, Talanta, 33 (1986) 265. H. Tao, A. Miyazaki and K. Bansho, Anal. Sci., 1 (1985) 169. P. Hocquellet and M. Candillier, Analyst, 116 (1991) 505. L. Liang, P.C. D’Haese, L.V. Lamberts, F.L.V. Vyver and M.E.D. Broe, Anal. Chem., 63 (1991) 423. J.M.E. Almendro, C.B. Ojeda, A.G. Torres and J.M.C. Pavon, Talanta, 40 (1993) 1643. F. Amore, Anal. Chem., 46 (1974) 1597. I. Kojima and S. Nomura, Anal. Sci., 11 (1995) 17. S.S. Leitner and J. Savory, Anal. Chim. Acta, 74 (1975) 133. J.B. Willis, Anal. Chem., 34 (1962) 614. H.T. Delves, G. Shepherd and P. Vinter, Analyst, 96 (1971) 260. T. Kumamaru, K. Murakami, F. Nakata, H. Sunahara, M. Kiboku and H. Matsuo, Anal. Sci., 3 (1987) 161. M.F. Enriquez-Dominguez, M.C. Yebra-Biurrun and M.P. Bermejo-Barrera, Analyst, 123 (1998) 105. C. Huang, M. Yang and T. Shih, Anal. Chem., 69 (1997) 3930. R.M. Barnes and J.S. Genna, Anal. Chem., 51 (1979) 1065. H. Asano, H. Itabashi and H. Kawamoto, Tetsu-to-Hagane, 87 (2001) 623. H. Asano, H. Itabashi and H. Kawamoto, Anal. Sci., 17(Suppl.) (2001) a347.

Chapter 16

Cryogenic trapping for speciation analysis M.P. Pavageau, E. Krupp, A. de Diego, C. Pe´cheyran and O.F.X. Donard

16.1

INTRODUCTION

Volatile metals and metalloid compounds (VMCs) and volatile organic compounds (VOCs) are important trace gases that inﬂuence atmospheric chemistry in many ways. Their distribution in the atmosphere is very complex and ﬂuctuates signiﬁcantly, and their concentrations vary from a few ppt to several ppb. The determination of VMCs is gaining increasing attention due to its ability to correctly assess global metal contamination budgets from emission sources. If the detection mode is well understood, most of the success for correct quantiﬁcation relies on using the appropriate sampling methods; these make use of careful cryofocusing after appropriate drying of the gaseous sample matrix. This chapter deals with the cryofocusing of VMCs. The procedures and analytical strategies largely take their inspiration from those used for VOCs. The sources and ﬂux of VMCs are becoming better known due to recent improvements in the whole analytical chain. VMCs have been identiﬁed as the result and signature of anthropogenic activities following their identiﬁcation and the recording of their emission from landﬁll and sewage gas or combustion ﬂue gas. Anaerobic fermentation processes are the main anthropogenic source of volatile organometallic compounds, whereas volatile inorganic compounds are emitted from combustion processes. In all cases, VMCs are diluted in a complex matrix. The primary constituents and their composition in various atmospheres are given in Table 16.1. Anaerobic fermentation processes take place in closed systems containing biomass (e.g. municipal landﬁlls and sludge digesters from wastewater treatment plants) wherein all the oxygen content has been consumed. Anaerobic bacteria then digest carbohydrates and produce large amounts of CH4 and CO2, as well as VMCs in smaller quantities. Concerning combustion processes, analysis of coal, wastes and fuel indicates that several metals are present at trace levels (e.g. Hg, Pb, Sn, Se, As, Cu, Sb). During combustion, these elements are redistributed in the bottom Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

495

M.P. Pavageau et al. TABLE 16.1 Gaseous constituents of the atmosphere compared to a typical ﬂue gas analysis of a coalﬁred power plant and biogas Gaseous constituents of the atmosphere N2 O2 Ar CO2 Ne He CH4 Kr H2 CO Xe O3 NO2 N2O NO NH3 SO2 CH3Cl C2H4 CCl4 CCl3F H2O HCl HF Hg H2S Hydrocarbons Halocarbons

78.1% 20.9% 934 ppm 330 ppm 20 ppm 5 ppm 2 ppm 1 ppm 500 ppb 100 ppb 90 ppb 40 ppb 20 ppb 300 ppb 6 ppb 2 ppb 0.5 ppb 0.2 ppb 0.1 ppb 0.1 ppb 0.1– 5%

Flue gas analysis of a coal-ﬁred power plant

12%

Sewage gas

Landﬁll gas

,4% ,1%

,1% ,1%

40%

35– 45%

54%

50– 60%

,1%

,1%

,100 ppm ,200 ppm ,100 ppm

,100 ppm

100 ppm

20 ppm 40 ppm 500 ppm 1500 ppm

8% 250 ppm 20 ppm 3 ppb

ash and ﬂy ash, and some are emitted with the main gaseous components (i.e., H2O, SO2, CO2) as volatile compounds. Volatile compounds of As, Sb, Sn, Pb, Hg [1,2], Mo, W, Ni, Fe, [3,4], and P [5,6] were recently detected in landﬁll gas, sewage gas and biogas. VMCs were also identiﬁed in natural environments such as hot springs [7], estuaries [8,9], natural waters [10] and soil gas [11], and these compounds can be used to trace the biogeochemical processes taking place in these environments. Concerning

496

Cryogenic trapping for speciation analysis

combustion processes, analysis of coal, wastes and fuel indicate that several metals are present in traces (e.g. Hg, Pb, Sn, Se, As, Cu, Sb). During combustion, these elements are redistributed in the bottom ash and the ﬂy ash, and some are emitted with the main gaseous components (i.e. H2O, SO2, CO2) as volatile compounds. Volatile compounds of Se, Hg, Sn, and Cu were found in coal combustion ﬂue gas [12]. Most of the analytical challenges associated with these anthropogenic signature assessments will be linked to the precise and accurate determination of the VMCs that are diluted in an important matrix, i.e., mainly CO2 or CH4. Sample preparation strategies will thus rely on cryogenic handling of the gases prior to their introduction into GC-ICP-OES or -MS systems. Various identiﬁed VMCs, together with their sources (either natural or anthropogenic) and the analytical procedures used by the various authors, are summarized in Table 16.2. Volatile metal(loid) species can be classiﬁed into four broad groups of gaseous species: hydrides, alkyl-, carbonyl- and halogenated species. Some VMCs and their chemical properties are also presented in Table 1 (Chapter 38). The identiﬁed metals associated with the formulations are: As, P, Sb, Bi, Ge, Ga, Sn, Cd, Tl, Pb, Hg, Ni, Fe, Mo or W. The most widely used technique for the measurement of VMCs is gas chromatography combined with different sensitive and selective detectors. Nevertheless, due to the low concentrations of the trace volatile compounds compared to the detection limits of the detectors used, it is most often necessary to employ preconcentration techniques before the analytical step. The preconcentration step can be performed either in the ﬁeld or in the laboratory. The volatile species can be concentrated on-site using various sampling techniques. These procedures, as well as the following analytical methodologies, are best known for VOCs and have been recently reviewed by Dewulf and Langenhove [13]. These techniques include preconcentration on various solid supports (e.g. sampling tubes ﬁlled with carboxen, tenax or SPME ﬁbres) at ambient or low temperatures, dissolution in speciﬁc absorption solutions, and cryogenic sampling procedures. Air samples can also be collected in canisters or plastic bags (e.g. Tedlarw bags) and later preconcentrated in the laboratory. The strategy associated with the sampling and handling procedure essentially depends on the nature of the emission and the concentration of the species involved. The whole analytical chain should prevent species degradation and allow the analysis of the integrity of the sample after the desorption step if preconcentration techniques are used. Indeed, the correct assessment of the fate of any metal in the environment requires exact species identiﬁcation to understand processes involved in its global cycling. This essential information can only be obtained if the sample preservation guarantees the integrity of the molecule. Such results require the combination of four different steps: 1. preconcentration of the gaseous analytes to gain enough sensitivity; 2. sample preservation prior to analysis;

497

498

TABLE 16.2 Natural and anthropogenic volatile metal(loid) species Volatile species Pb(CH3 )4, Pb(CH3)3(C2H5), Pb(CH3 )2(C2H5 )2, Pb(CH3)(C2H5)3, Pb(C2H5)4

Urban area, country, sea, sewage gas

Urban area, country, sea, industry landﬁll, sewage combustion ﬂue gas

Sampling technique Cryogenic trapping (2728C)/chromosorb— S.T.: 16 h, V.: 70 l Cryogenic trapping (21308C)/glass wool— S.T.: 1 h, V.: 360 l Porapack—S.T.:24 h, V.: 140 l Cryogenic trapping (2808C)/Supelcoport— S.T.: 20 min, V.: 20 l Cryogenic trapping (21758C)/glass wool— S.T.: 18 min, V.: 15 l Water purge and cryogenic trapping (21968C)/ glass wool Tenax—S.T.: 3.5 days, V.: 10 m3 Gold trap—S.T.: 16 h, V: 200 l Gold trap—S.T.: 1 h, V: 50 l Absorption solutions K2Cr2O7/HNO3

Analysis

Detection limit

Reference(s)

TD-GC-GF-AAS

500 pg/m

3

[64]

TD-LT-GC– AAS

200 pg/m3

[65]

TD-LT-GC– AAS

250 –370 pg/m3

[66]

TD-LT-GC– ICPMS

43 pg/m3

[1,2]

TD-LT-GC– ICPMS

6 ng/m3

[23]

TD-LT-GC– ICPMS

0.4 pg/l of water

[10]

TD-LE (benzene) -GC–AAS TD-CV-AAS, TD -CV-AFS TD-LT-GC– ICPMS CV-AFS

200 pg/m3 NC 600 pg/m3

[67,68] [69] [70]

NC

[16]

M.P. Pavageau et al.

Hg8, HgCH3Cl, Hg(CH3)2, HgCl2

Environment

TABLE 16.2 (continuation) Volatile species

AsH3, As(CH3 )H2, As(CH3)2H, As(CH3)3, As(C2H5)(CH3)2

Sampling technique

Carbotrap—S.T.: 13 j, V: 2 –9 m3 Chromosorb ambient temperature (HgCH3Cl) and at 2808C (Hg(CH3)2)—S.T.: 4– 69 h, V.: 0.3 –5.4 m3 Water purge and cryogenic trapping (21968C)/glass wool Country, sea, forest, lake, Cryogenic trapping natural waters, estuarine (21308C)/glass wool —S.T.: 4 h, V.: 700 l waters, geothermal and soil gases Water purge and cryo-

Landﬁll, sewage, microelectronic industry soil and geothermal gases

genic trapping (21968C)/glass wool w Tedlar bag followed by cryogenic trapping (2808C)/chromosorb—V.: 3l Cryogenic trapping (2808C)/Supelcoport— S.T.: 20 min, V.: 20 l

Analysis

Detection limit

Reference(s)

TD-LT-GC– CVAFS TD-GC– MIP-AES

20 pg/m3

[69]

100 pg/m3

[71]

TD-LT-GC– ICPMS

0.2 pg/l of water

[10]

TD-LT-GC– AAS

200 pg/m3

[72,73]

TD-LT-GC– ICPMS

0.8 pg/l of water

[10]

LT-GC– ICP-MS

NC

[11]

TD-LT-GC– ICPMS

27 pg/m3

[1,2]

499

continued

Cryogenic trapping for speciation analysis

Se(CH3)2 , Se2(CH3)2, (CH3) SeS(CH3)

Environment

500

TABLE 16.2 (continuation) Volatile species

Environment

Sampling technique

SnH4, Sn(CH3 )4 , Sn(C4H 9)H3, Sn(C4H9)H2

SbH3, Sb(CH3)3

Landﬁll, sewage, anoxic sediments, estuarine waters

Bi(CH3)3

Sewage, metal industry, geothermal gases, landﬁll, sewage Landﬁll, sewage

Te(CH3)2

Landﬁll, sewage

PH3

Landﬁll, sewage, fumigation processes

Detection limit 3

Reference(s)

TD-LT-GC– ICPMS

70 pg/m

[74]

TD-LT-GC– ICPMS

100 pg/m3

[11]

TD-LT-GC– ICPMS

14 pg/m3

[1,2]

TD-LT-GC– ICPMS

0.05 pg/l

[10]

GC–AAS

NC

[75]

TD-LT-GC– ICPMS

8 pg/m3

[1,2]

TD-LT-GC– ICPMS

7 pg/m3

[1,2,76]

TD-LT-GC– ICPMS

5 pg/m3

[1,2]

TD-LT-GC– ICPMS

250 pg/m3

[17]

M.P. Pavageau et al.

Cryogenic trapping (21758C)/chromosorb— S.T.: 20 min, V.: 10 l þ NaOH cartridge Cryogenic trapping (2808C)/Supelcoport— S.T.: 20 min, V.: 20 l Cryogenic trapping (2808C)/Supelcoport— S.T.: 20 min, V.: 20 l Water purge and cryogenic trapping (21968C)/glass wool Direct gas injection (0.5 ml) Cryogenic trapping (2808C)/Supelcoport— S.T.: 20 min, V.: 20 l Cryogenic trapping (2808C)/Supelcoport— S.T.: 20 min, V.: 20 l Cryogenic trapping (2808C)/Supelcoport— S.T.: 20 min, V.: 20 l Cryogenic trapping (21808C)/glass wool— S.T.:40 min, V.: 20 l þ NaOH cartridge

Analysis

TABLE 16.2 (continuation) Volatile species

Environment

Sampling technique

Direct injection

Ni(CO)4, Fe(CO)5, Mo(CO)6, W(CO)6 CH3I

Sewage

Geothermal gas, estuarine waters

S.T.: sampling time; V.: volume, NC: not calculated.

Detection limit

Reference(s)

GC– FPD, GC(nitrogen–phos phorus)– FID GC-nitrogen–phos phorus detector

NC

[5]

70 pg/m3

[6]

LT-GC–ICP-MS

100 –105 pg/m3

[4]

LT-GC–ICP-MS

NC

[7]

TD-LT-GC– ICPMS

0.1 pg/l of water

[8]

TD-LT-GC– ICPMS

4 pg/m3

[8]

Cryogenic trapping for speciation analysis

Tedlarw bag þ NaOH catridge, cryotrapping, V.: 50 ml w Tedlar bag followed by cryogenic trapping (2808C)/chromosorb V.: 1 l w Tedlar bag followed by cryogenic trapping (2808C)/chromosorb V.: 1 l Water purge and cryogenic trapping (21968C)/glass wool Cryogenic trapping (21968C)/glass wool—S.T.:30 min, V.: 30 l

Analysis

501

M.P. Pavageau et al.

3. efﬁcient qualitative and quantitative desorption procedures; 4. selective detection and quantiﬁcation of the analytes. These requirements are best achieved using cryogenic trapping techniques followed by appropriate hyphenated detection systems. Cryofocusing techniques are usually selected for the clean and complete thermal efﬁciency of desorption of the trace gas of interest. The aim of this chapter is to review and present the physico-chemical principles of cryotrapping along with their associated analytical constraints. The last part of the chapter presents an overview of recent cryogenic method developments in the ﬁeld of sampling and the analysis of gaseous, liquid and solid samples for species determination. 16.2

DEFINITION OF VOLATILE SPECIES

A general and global deﬁnition characterising all volatile species ranging from VOCs to VMCs does not exist. Similar to what has been adopted for VOCs by the US-EPA, both VMCs and VOCs can be generally classiﬁed as chemical compounds having a vapour pressure greater than 0.1 mmHg (0.01 kPa) at 258C and at atmospheric pressure. The theory assumes that, under these conditions, molecules in gases are largely dispersed and can therefore be considered independent of one another. The properties of gases can therefore be described using the ideal gas law ðPV ¼ nRTÞ: This hypothesis is needed as it is used for the calculation of partial pressures. In terms of units, the relationship between ppm and mg/m3 at 258C is given by the following equation: concentration ðppmÞ ¼ concentration ðmg=m3 Þ £ molar mass=24:45: We will brieﬂy review some of the terms and deﬁnitions used later in the text to correctly assess the processes associated with cryofocusing. Vapour pressure: The vapour pressure of a compound is a direct measure of the ability of molecules to transfer from the liquid or solid phase and remain in the gaseous state at a given temperature. The higher the vapour pressure for a given temperature, the more volatile the compound will be. Boiling point: The boiling point of a compound is deﬁned as the temperature at which the saturated vapour pressure of a liquid is equal to the surrounding atmospheric pressure. Because the vapour pressure increases with temperature, it follows that for an ambient pressure greater than 760 mmHg, the boiling point of water is above 1008C and for pressures less than 760 mmHg, the boiling point is lower than 1008C. The stability of the boiling point makes it a convenient calibration temperature for temperature scales. As an example, Fig. 16.1 displays and compares the vapour pressures of PH3, Hg8, CO2 and H2O at different temperatures [14]. PH3 is deﬁnitely the most volatile species, whereas Hg8 is the least volatile of the four

502

Cryogenic trapping for speciation analysis

Fig. 16.1. Effect of temperature on the partial vapour pressure of Hg8, H2O, CO2 and PH3.

compounds. At ambient temperature and atmospheric pressure, PH3 is in the gaseous state whereas Hg8 is in the liquid state. Using the above deﬁnitions and a temperature of 258C under atmospheric pressure, a large variety of volatile metal(loid) species (hydride or alkyl) can occur in the gaseous or liquid state in natural or anthropogenic environments. Some of them are listed in Table 16.1. The volatile nature of these metal(loid) species, which possess relatively high molecular weights, is essentially due to their low intermolecular interactions. Volatile compounds: We will refer throughout this chapter to volatile compounds as those species corresponding to the deﬁnition and conditions described above. This is in contrast to what is commonly referred as “volatile metals” in the major literature. Indeed, in most other cases, volatile species are deﬁned according to their ability to pass through a ﬁltration device. For example, normative sampling procedures concerning combustion ﬂue gas for total metal determination make full use of this deﬁnition [15,16]. The method is based on chemical absorption of the species in solutions after ﬁltration. According to this speciﬁcation, gaseous species and all small particles which pass through the ﬁlter will be classiﬁed as “volatile species”. In this case, classiﬁcation according to the term “volatile” directly and only depends on the ﬁltration efﬁciency. This deﬁnition does not take into account the thermodynamic properties of the molecules. Under these conditions, some of the species classiﬁed as “volatile species” are very small but in the solid state. It is in fact very important to be able to classify the species of interest according to their physical state to be able to really assess the environmental and health impact. Therefore, in this chapter, the deﬁnition used for “volatile metal species” will only make reference to those species displaying gaseous physico-chemical properties at ambient temperature.

503

M.P. Pavageau et al.

16.3

PHYSICO-CHEMICAL PRINCIPLES AND PROCESSES ASSOCIATED WITH CRYOFOCUSING

Sample preparation using cryofocusing can be used in a wide array of situations ranging from “on-site” trapping of VMCs to direct trapping prior to detection during the analytical procedures. In all cases, it is necessary to understand the physico-chemical processes taking place. An ideal situation is presented in Fig. 16.2. The cryogenic trapping technique is based on the condensation of the molecules on a cooled surface. The condensation depends on the surface temperature and on the partial vapour pressure of the molecules in the gas phase. If the surface is free of adsorption sites, the trapping efﬁciency will only depend on the above two parameters. In this case, cryotrapping should consist of a simple change of physical phase of the compounds of interest. A theoretical ﬂow of molecules is presented to an inert surface maintained at 2 1408C. The gaseous compounds having partial vapour pressures much less than their saturated vapour pressure at 2 1408C will change phase and will deposit from the gaseous state as solid or liquid compounds on the cooled surface. When the partial pressures of the compounds of interest are higher than their saturated vapour pressures, they will then change phase again and convert from the solid to the gaseous state. The role of the temperature in this respect is essential, because it serves to condense and preconcentrate the analytes of interest upon cooling. During a second subsequent stage, it can be used to selectively release the analytes.

Fig. 16.2. Molecules in the gas phase upon contact with a cryo-cooled surface.

504

Cryogenic trapping for speciation analysis

However, ideal situations are never the case, and potential interferences associated with the solid substrate must be taken into account. To support the understanding of cryogenic trapping mechanisms, we will refer to two different studies dealing with the inﬂuence of the sampling temperature for the determination of PH3 and detailing the trapping efﬁciency of Hg8 under cryogenic conditions [17,18]. Both experiments were conducted using different sampling temperatures based on a “home-made” cryogenic air sampler illustrated in Fig. 16.3. Brieﬂy, in this device, the sample is collected in a Pyrex tube ﬁlled with silanized glass wool, and the trapping temperature in the tube is regulated by combined ﬂow of liquid nitrogen and a surrounding heating wire. Appropriate programming of both temperature regulation systems (cooling and heating) allows a precise trapping temperature ranging from 220 to 21908C (^28C). The gas standard mixture of Hg8/air was generated using a permeation chamber (Calibrage, Saint Chalmas, France). The PH3/air mixture was prepared by diluting a PH3 gas standard (produced at 10.7 ppm in helium, Air Liquide) in a mixing chamber. The samples were then analysed by LT-GC –ICP-MS, and the principal observations and conclusions are presented below. Figure 16.4A shows the inﬂuence of sampling temperature on the trapping efﬁciency of phosphine from 2 190 to 2 358C. Three different regions can be distinguished on this graph: (1) from 2 190 to 21608C, the trapping efﬁciency for phosphine clearly is independent of sampling temperature. The results concerning the study of the inﬂuence of phosphine concentration, sampling ﬂow and sampling time versus the quantity of PH3 collected at 21808C indicate that the quantity of collected phosphine is proportional to the variations of concentration, sampling ﬂow and sampling time. This means that gaseous phosphine is quantitatively collected below 2 1608C. The saturated vapour pressure diagram [19] also indicates that below 21348C phosphine exists in the solid state. (2) from 2 160 to 2 1108C, the trapping efﬁciency for phosphine decreases from 100 to about 5% (phosphine melting point is 2 1348C). (3) from 2 110 to 2358C, the trapping efﬁciency for phosphine is about 5%, despite the fact that the phosphine vapour pressure diagram [19] indicates that the partial pressure of phosphine (1.5 £ 1025 mmHg) in the sampled ﬂow (C ¼ 0.028 mg/l at 258C) is lower than the saturated vapour pressures referenced in the range of sampling temperatures from 2110 to 2 358C. The vapour pressure of phosphine at 21108C is about 200 mmHg. This means that between 2110 and 2358C, phosphine exists in the gaseous state and not in the liquid state, and no trapping should be obtained. Therefore, these results highlight the occurrence of phosphine adsorption mechanisms on the surface of the glass wool and sampling tube in this temperature range. Free molecules in the gas phase and adsorbed molecules on the surface are in dynamic equilibrium, and the surface coverage, u, depends on the pressure of the adsorbed gas. The variation of u with pressure at a given temperature is described by an adsorption isotherm. Different adsorption isotherms are

505

506 M.P. Pavageau et al. Fig. 16.3. Laboratory made cryogenic air sampler—the conﬁguration presented (drying device) was used for sampling ﬂue gas during coal combustion.

Cryogenic trapping for speciation analysis

Fig. 16.4. (A) Inﬂuence of sampling temperature from 2190 to 2358C (^28C) on the trapping efﬁciency for phosphine (phosphine concentration: 0.028 mg/l at 258C, sampling ﬂow: 0.1 Nl/min and sampling time: 2 min). RSD calculated for n ¼ 3: (B) Inﬂuence of sampling temperature from 2190 to 2208C (^28C) on the trapping efﬁciency for elemental mercury (concentration: 2 ng/l at 258C, sampling ﬂow: 0.1 Nl/min and sampling time: 2 min). RSD calculated for n ¼ 3:

507

M.P. Pavageau et al.

given in the literature [20]. In order to further characterize the adsorption mechanisms, the samples were collected at 21008C and the experiments were performed using different phosphine concentrations, sampling times and quantities of glass wool support. These experiments showed that the quantity of trapped phosphine is proportional to the phosphine concentration, but independent of sampling time, and decreases with lower quantities of glass wool. Therefore, we assumed that the trapping mechanism of phosphine at 21008C should follow a Langmuir isotherm [21]. The Langmuir isotherm is based on three hypotheses, namely that all the adsorption sites are equivalent and independent of each other and there is monolayer adsorption. The dynamic equilibrium is: AðgÞ þ MðsurfaceÞ $ AMðsurfaceÞ: The adsorption is proportional to the partial vapour pressure of A and to the number of unoccupied sites, Nð1 2 uÞ; with N being the total number of sites. The rate of desorption is proportional to the number of adsorbed molecules, Nu: When equilibrium is reached, u is constant and can be expressed by the following equation: u¼

KP 1 þ KP

ð16:1Þ

with K ¼ Ka =Kd and u being the surface coverage (the ratio of adsorbed molecules to adsorbing sites), P is the partial pressure (gaseous ﬂow concentration) (Pa), and Ka and Kd are the rates of adsorption and desorption, respectively. The BET isotherm, derived from Langmuir isotherm, is less probable. It considers a multilayer adsorption phenomenon wherein the condensation of molecules on the covered surface is unlimited. In our case, according to the PH3 concentration in the gas, the surface occupied by the molecules of PH3 is about 1026 m2, and the speciﬁc surface area of the glass wool can be estimated to be about 0.7 m2/g, so a monolayer adsorption phenomenon is the most probable. Above 21108C, the adsorption phenomenon is proportional to the ﬂow of gaseous phosphine at a ﬁxed concentration but depends on the quantity of glass wool support. This fact implies that the quantity of glass wool support should be rigorously the same for each sampling tube to be able to quantify the trapped phosphine above 2 1108C. This property can be used for trace metal quantiﬁcation in CO2-enriched atmospheres. Similar types of results were obtained for the cryotrapping of gaseous Hg8. The inﬂuence of sampling temperature on the trapping efﬁciency of elemental mercury from 2190 to 2 258C is presented in Fig. 16.4B. In order to generate a continuous ﬂow of Hg8, a permeation chamber was used. A polytetraﬂuoroethylene (PTFE) tube (internal diameter: 6 mm, thickness: 1 mm, length: 114 mm) was ﬁlled with pure Hg8 and placed into an oven set at 80 ^ 0.18C. Hg8 vapours were diluted in a puriﬁed nitrogen ﬂow. The resulting concentration of elemental mercury in the gas mixture was 2 mg/m3 at 258C.

508

Cryogenic trapping for speciation analysis

As observed for phosphine (Fig. 16.4A), three different regions can be detailed on the graph: (1) from 2 190 to 2 1408C, to obtain quantitative trapping of Hg8, the temperature must be lower than 2 1408C. At 21808C, the trapping efﬁciency is proportional to the sampling time. (2) from 2 140 to 21008C, the trapping efﬁciency decreases from 100 to about 20%. (3) from 2 100 to 2208C, the trapping efﬁciency decreases slowly. It may be noted that at 2808C, less than 10% of total mercury is collected and the trapping is then independent of sampling time. No literature data [22] referring to the vapour pressure of elemental mercury could be found for temperatures below 2 388C. The vapour pressure of Hg8 at 2 388C corresponds to 1.5 £ 1026 mmHg. In the gaseous mixture sampled, the partial pressure of elemental mercury is 1.8 £ 1027 mmHg. Therefore, at 21008C, Hg8 is in gaseous state; nevertheless, some Hg8 was still trapped and recovered at temperatures above 2 388C (about 5%). Then, as observed for phosphine, adsorption mechanisms should be involved in this range of sampling temperatures. In conclusion, two points have to be mentioned regarding the cryogenic trapping procedure. First, for a ﬁxed sampling temperature, the trapping efﬁciency for different analytes present in the sample depends on their partial vapour pressure. For complete collection, the sampling temperature must be maintained lower than the temperature corresponding to an equilibrium vapour pressure of the analytes of interest. Second, the role of the trapping surface is important because it will not be totally inert with regards to adsorption of the analyte. Silanized glass surfaces were found to be the least “adsorptive” or present the smallest “memory effect”. Other solid sorbents, such as Chromosorb and Tenax, can also be used for the sampling of volatile species at low temperature in order to enhance the trapping efﬁciency. Desorption efﬁciency is, however, most often less than that observed from less inert solid supports.

16.4

ANALYTICAL CONSTRAINTS

If, in general, cryofocusing appears to be an attractive and effective technique for preconcentrating volatile species, the gaseous matrix in which they are present during cryocollection can create a large array of problems that need to be addressed in order to obtain qualitative and quantitative information. Volatile metal species occur in natural and anthropogenic gaseous emissions within an array of different matrix gases. Major constituents of matrix gases generally present in combustion ﬂue gas, biogas or atmospheric air are listed in Table 16.1.

509

M.P. Pavageau et al.

The most common gaseous species in which the analytes are diluted are CO2 and/or water. These are the two main entities that are ubiquitous in all natural and anthropogenic gaseous samples, and they can be considered as severe problems for the quantitative determination of the analytes. Indeed, water, CO2 or other major matrix compounds trapped simultaneously during the sample collection are often inevitable and will create problems during the sampling step or later, during the analytical run. These common problems include peak distortion, shifts in retention time, a rising baseline during elution of the analyte or direct physical problems such as clogging of the traps. Much of the success of the sample preparation will depend on its ability to either remove them or to fully minimize their inﬂuence during the detection step of the analytical procedure. The use of a selective hydrophobic sorbent support could potentially avoid some of the above problems, but both the adsorption and desorption processes related to the speciﬁc sorbent support involve chemical interactions with analytes that are likely to change their chemical identities, and hence alter the quality of the determination. In order to investigate trace volatile species in these atmospheres, different sampling strategies have to be used to avoid major matrix compounds, according to the species of interest. The cases of CO2 and water are detailed in this chapter. For most successful determinations, the “sample preparation” associated with these types of matrix will refer to the efﬁcient “removal” of their effect before or during the analytical run. Some of the main analytical strategies dealing with appropriate “sample preparation” for the correct determination of gaseous compounds are reviewed below. 16.4.1 Removal of CO2 Two strategies can be used. The ﬁrst one makes use of differential temperature trapping and the second one makes use of direct and selective removal of the CO2 matrix onto solid NaOH during the detection stage. 16.4.1.1 Selective temperature strategy to avoid CO2 trapping Gaseous samples can generally be collected at a set temperature in U-shaped tubes ﬁlled with glass wool cooled at different temperatures depending on various bath mixtures. The type of mixture and resulting trapping temperatures are referenced in Table 4-9-2 of Chapter 4-9 and also by Namiesnik and Wardencki [27]. Automated cryosampling devices may also be used with adjustable temperature settings similar to the one developed “in house” and presented in Fig. 16.2. Trapping of VMCs in ambient air: In the case of collection of VMCs in ambient air, the partial pressure of CO2 is about 0.25 mmHg, and CO2 will only be efﬁciently collected together with trace compounds at sampling temperatures lower than 21408C. Therefore, above 2 1408C, CO2 will diffuse

510

Cryogenic trapping for speciation analysis

unaffected through the trapping device. Nevertheless, the vapour pressure of VMCs in ambient air is generally too low to ensure quantitative trapping below 21408C (see PH3 and Hg8). The use of selective temperatures requires that the considered matrix compound is signiﬁcantly more or less volatile than the species of interest. According to Pe´cheyran et al. [23], tetraalkyl lead species and Hg8 were sampled at 21758C, and the collected volume was 15 l; this corresponds to about 5 ml of pure CO2. The samples were analysed by LTGC –ICP-MS. Under these sampling and analytical conditions, the CO2 collected with trace compounds was not described as a problem. Trapping of VCMs under harsh conditions: In some cases, it may be necessary to trap all the gases, which are present in order, to collect and characterize very volatile analytes. In this case, all gases are cryocondensed simultaneously in the sampling device at very low temperatures, such as at 21758C. These collection temperatures, however, generate some severe problems. Depending on the sampling temperature, major gases, such as CO2 (12%) or SO2 (1500 ppm) in combustion ﬂue gas, may clog the sampling tube during cryogenic sampling, preventing quantitative and reproducible analyte collection [12]. Similar drawbacks have been observed for VMCs when sampling at 21968C in atmospheres enriched in CH4 (36%) and CO2 (24%) [24]. During experiments using cryogenic sampling at 21758C for collection of volatile metal species in coal combustion ﬂue gas [12], it has been demonstrated that CO2 is trapped together with the trace compounds. It disrupts the analytical procedure by extinguishing the plasma when using LT-GC –ICP-MS analysis. This problem can then be overcome by refocusing the samples and the analytes of interest at 2808C. Further, under these refocusing temperatures, only a very small fraction of CO2 should be collected as the saturated vapour pressure of CO2 at 2808C is about 400 mmHg, and the partial pressure of CO2 in the case of a ﬂue gas sample is lower than 100 mmHg. It should be noted that, at 2808C, some volatile compounds (having a partial vapour pressure lower than the corresponding saturated vapour pressure) will not be trapped quantitatively.

16.4.1.2 Chemical reagents for removal of CO2 Making use of different boiling points is not sufﬁcient when the analyte(s) and the major component(s) of the gaseous matrix have similar vapour pressures (e.g., CO2 and PH3 have similar vapour pressure) and cannot be separated by a temperature difference. In such case, a chemical reagent has to be used during the sampling procedure prior to cryogenic trapping at 2 1908C. This procedure can be repeated during the analytical step to directly eliminate the CO2 matrix online. The perturbation effect of CO2 on PH3 quantiﬁcation using cryogenic trapping followed by GC – ICP-MS (dry plasma conditions) was recently reported [17]. Determination of phosphine requires the complete elimination of CO2. Its concomitant elution will signiﬁcantly alter the plasma stability. The

511

M.P. Pavageau et al.

introduction of a cartridge of solid NaOH into the analytical device selectively removes CO2 and permits quantitative species determination without plasma perturbation. These important effects are illustrated in Fig. 16.5A and B, where the efﬁciency of removal of CO2 during determination of phosphine using a solid cartridge of NaOH is presented. The effect of CO2 on plasma was studied by injecting 1 and 5 ml of pure CO2 into the analytical device (which corresponds to the quantity of CO2 trapped at 21808C for an air sampling volume from 3 to 17 l at 350 ppm). Phosphine interaction with NaOH was evaluated by injecting a deﬁned volume of PH3 [100 ml of 10.7 ppm PH3 in helium, stored in a Tedlarw bag (Supelco)] and performing analysis with and without the NaOH cartridge (Teﬂon tube, length: 30 cm, i.d. 8 mm, 20 g of NaOH beads). The NaOH cartridge was connected between the injection port and the chromatographic column and was left in place during all analytical runs. Results presented in Fig. 16.5A clearly illustrate the inﬂuence of CO2 on the plasma stability. When CO2 elutes, an increase of intensity at the m=z 31 and m=z 13 signals is detected, while the intensity of the signal of the internal gaseous standard (124Xe) decreases. Feldmann et al. [25] also observed this effect using wet plasma conditions and after nebulising an indium solution as an internal standard. When CO2 is present at the detection limit (ICP-MS), the m=z 31 signal increases even without phosphine injection. 13C18Oþ polyatomic ions could be formed in the plasma and produce an isobaric interference on the phosphorus 31 m=z signal when CO2 elutes. Under these conditions, the plasma is so perturbed that quantiﬁcation is not possible during the elution of CO2. The problems then lie with the fact that CO2 and PH3 are co-eluted (Fig. 16.5B). Further, the CO2 elution changes the real retention time of phosphine (Fig. 16.5A and B). This type of effect can be a problem when an accurate retention time estimate is needed for species identiﬁcation [26]. When an NaOH cartridge was inserted into the analytical line, CO2 was very efﬁciently absorbed; the internal standard presents a steady-state signal and the RSD of the retention time values is lower than 3%. However, disadvantages of using a solid NaOH cartridge exist because unstable compounds or slightly acidic compounds can react with the solid substrate. Feldmann et al. [25] have indeed observed that some organometallic and hydride compounds of arsenic and antimony can be affected by the presence of the NaOH cartridge, whereas no signiﬁcant effect was observed for tin compounds. In the case of phosphine, no interactions with NaOH could be detected. Indeed, when NaOH cartridge was used, a recovery of about 97% was calculated from the area of the chromatographic peaks. Area RSD values , 4% both with and without the NaOH cartridge were achieved. 16.4.2 Water removal The drying process from natural gases, besides removing the largest amount of water present in the sample, should not affect the initial distribution of the

512

Cryogenic trapping for speciation analysis

Fig. 16.5. (A) Typical chromatogram of PH3 (100 ml, 10.7 ppm) with CO2 injection (1 ml) without NaOH cartridge. (B) Chromatogram of PH3 (100 ml, 10.7 ppm) using an NaOH cartridge.

513

M.P. Pavageau et al.

gaseous analytes of interest. Many procedures have been used with varying degrees of success. The most efﬁcient procedures used for drying a gas stream can be divided into four groups [27]: (1) a wide variety of desiccants using cartridges ﬁlled with hygroscopic solids such as P2O5, K2CO3, Na2CO3, Mg(ClO4), CaCl2, Na2SO4 or MgSO4. Previous studies, comparing the drying efﬁciencies of CaCl2, Na2CO3 and MgSO4 for a VOC mixture, concluded that only the Na2CO3 reagent did not result in signiﬁcant losses [28] under the conditions tested and for the selected VOCs. Among the variety of drying sorbents described in the literature, only some of them have been applied in combination with cryotrapping of volatile organometallic species: potassium carbonate (K2CO3), calcium chloride (CaCl2), sodium hydroxide (NaOH) and magnesium perchlorate (MgClO4). For the use of calcium chloride, no loss was reported for SnH4 [29], but 15 –20% loss occurred for methylated arsenic hydrides, whereas the use of sodium hydroxide or magnesium perchlorate as the drying agent did not result in loss of these arsenic compounds [30]. Potassium carbonate was reported to be very efﬁcient at low gas ﬂows below 100 ml/min, showing no loss for methyliodide and organic selenium species [31]. Similar results were obtained for the use of magnesium perchlorate [32]. For several volatile tin compounds, calcium chloride, as well as magnesium perchlorate, did not cause species loss, but magnesium perchlorate was found to be a very efﬁcient drying sorbent using gas ﬂows of 1 l/min. Thus, Feldmann [24] systematically used magnesium perchlorate for the probing of environmental gases. (2) classical adsorbents, such as silica gel, molecular sieves (zeolites), calcium or sodium silicates. (3) the second last series of devices are based on drying the analytes by condensation of water while passing the gas stream through a cooled trap. This ensures a decreased water vapour content if the residence time is sufﬁcient. Under these conditions, the partial pressure of water vapour is related to the temperature of the trap as given in Table 16.3. The drying capacity of these three different classes of dryers, adapted from the review of Namiesnik and Wardencki [27], are reported in Table 16.3. The ﬁrst and second series are very popular because of their high drying efﬁciency, but they can induce total or partial losses of some trace compounds, as indicated by the following experiment. The previous different approaches have been tested and compared for determinations of VOCs. In a recent study on a mixture of VOCs (concentration range ng/l), two different hygroscopic agents (soda lime and Na2CO3), one adsorbent (molecular sieve) and drying by cryocondensation were selected [33]. The recoveries obtained by comparing the results to those obtained without using any dryers are reported in Table 16.4. The RSD values are lower than 10% when the compounds of the mixture are not affected by the dryer. The results demonstrate that drying with soda lime, Na2CO3 and a molecular sieve affects the sample composition. This is due to adsorption of analytes together

514

Cryogenic trapping for speciation analysis TABLE 16.3 Drying efﬁciencies of different types of water trap systems at different temperatures Temperature (8C) Drying agent P2O5 Silica gel CaCl2

þ 20 þ 20 þ 20

Partial pressure of water (mmHg)

0.00002 0.01 0.2

Drying by condensation þ 20 0 220 270 2100

17.5 4.6 0.77 0.002 0.00001

Adapted from Namiesnik and Wardencki [27].

with water or chemical reactions of the VOCs on the surface of the desiccant. A cryogenic water trap held at 2 208C (acetone and nitrogen bath), however, did not result in signiﬁcant losses of the considered VOCs. When passing through the water trap, the gas mixture components will condense (or not) according to their relative vapour pressures. In this case, the partial vapour pressures of all the VOCs are below their saturated vapour pressures at 2208C. Therefore, it can be concluded that, generally, a water trap at 2 208C can be used for the determination of low concentration mixtures. (4) drying by permeation. This technique is quite efﬁcient and has seen many developments. It generally makes use of a tube made of Naﬁonw, a copolymer of tetraﬂuoroethylene and perﬂuoro-poly(triﬂuoromethylo(oxapropylo))-sulphonic acid. The driving force of water movement through a Naﬁonw membrane for its removal from the eluting gas is based on the difference in the partial pressures of water on both sides of the Naﬁonw wall. The role of temperature and the drying efﬁciency of the Naﬁonw drying system have been reported in the literature [34]. The permeation or drying process can be increased by using a counter current of dry gas. This device has been developed in the laboratory and uses the boil-off from the cryostat as this gas. These types of drying systems do not prevent partial or total lost of some polar compounds such as ketones, amines, ammonia, alcohol, acids or dimethyl sulfoxide due to absorption and/or permeation processes [35]. Recent studies on the efﬁciency of Naﬁonw and the recovery of volatile compounds were conducted with a synthetic mixture of PH3 in air, as well as a VOCs mixture diluted in air [17,36]. Air moisture was removed before cryogenic sampling at 2 1808C using a cooled Naﬁonw membrane. The Naﬁonw

515

516 TABLE 16.4 Recovery results for VOCs mixture using four different drying system before cryogenic trapping (21808C)

Methanol

Acetone

Dichloro-

Methyl-

Ethyl

Dichloro-

Methyliso-

methane

ethylketone

acetate

ethane

butylketone

Toluene

Ethyl-

Paraxylene

benzene

Without dryer

100

100

100

100

100

100

100

100

100

100

Soda lime (5 g)

66 (36%)

76 (1%)

100 (1%)

62 (1%)

42 (17%)

, DL

10 (24%)

99 (1%)

96 (3%)

87 (4%)

Na2CO3 (5 g)

44 (28%)

100 (52%)

98 (4%)

, DL

, DL

98 (1%)

, DL

99 (1%)

99 (1%)

99 (1%)

Cryocondensation

96 (29%)

Blank problem

95 (7%)

99 (2%)

96 (5%)

94 (7%)

91 (5%)

98 (1%)

97 (1%)

97 (1%)

, DL

, DL

99 (1%)

, DL

, DL

86 (6%)

3 (8%)

81 (3%)

60 (8%)

60 (8%)

at 2 208C Molecular sieve 5A (20 g)

(RSD%).

M.P. Pavageau et al.

Recovery (%)

Cryogenic trapping for speciation analysis

membrane dryer tube (length 24 cm, i.d. 2 mm and o.d. 3 mm; Perma Pure Products, New Jersey) was maintained at 2108C in order to enhance drying efﬁciency. Experiments were conducted with and without a Naﬁonw membrane in order to detect possible losses of compounds at concentrations in the range of ng/l. Results concerning a PH3 and VOCs mixture are reported in Table 16.5. The use of the Naﬁonw membrane for drying led to the partial loss of the polar VOCs from the mixture (i.e., alcohols and ketones). These losses were attributed to hydrogen bonding with the sulfonic acid groups surrounding the ﬂuorocarbon matrix (http://www.permapure.com). In the case of PH3, the losses were estimated to be less than 15% due to the slight polarity of the PH3 compound. Finally, the Naﬁonw membrane can also be used for the drying of gases containing VMCs. These statements can be illustrated by the following tests and results. To test the recovery of volatile metal species and Hg8, a mixed solution of organometallic compounds (Me4Pb, Et4Pb, Me4Sn, Et4Sn, Me2Se and Me2Se2) and Hg8 vapour from a closed vial were vaporized and injected through the air sampler (Fig. 16.3) into a clean synthetic air stream (2 Nl/ min). The drying Naﬁonw membrane was set at different temperatures. The results, in terms of concentrations following application of different drying conditions, did not show any loss of the analytes of interest, as presented in Table 16.5. In general, all adsorbents have a speciﬁc and limited drying capacity. Therefore, after several operating cycles they should be regenerated. The important advantage of molecular sorbents is the possibility for their multiple regeneration (over 2000 cycles) without changing their adsorption capacity [27]. In contrast to other dryers, the Naﬁonw membrane drying capacity can be auto-regenerated [37]. In conclusion, water removal is an essential step in sample preparation for the determination of gaseous species. The array of drying systems should be evaluated according to the species of interest. Drying processes, such as selective condensation or drying using Naﬁonw membranes, appear to generate the best results for VMCs determination.

16.5

SAMPLE PRESERVATION AND STABILITY

Once collected by various methods (i.e., by cryogenic sampling or Tedlarw bags at ambient temperature), the samples must be stored prior to detection. Thus, stability of the gaseous analytes is of paramount importance for the validity of the results generated and only few studies have been made on this topic. Cryoconservation greatly decreases reaction time. For example, for a single-step reaction with an activation energy of 53 kJ/mol that takes 1 s at room temperature (258C), the reaction time is prolonged to 1018 years at 21908C [38].

517

518

TABLE 16.5 Recovery results for VOCs mixture, Hg8 and organometallic compounds using a Naﬁonw membrane set at different temperatures before cryogenic trapping (21808C)

(RSD%).

Naﬁon (þ108C)

Naﬁon (þ08C)

Naﬁon (2108C)

Naﬁon (2208C)

Naﬁon (2308C)

Naﬁon (2508C)

95 (1%) 92 (1%) 120 (2%) 95 (2%) 92 (4%) 85 (7%) 97 (5%) 91 (1%)

94 (3%) 92 (2%) 120 (5%) 90 (2%) 98 (5%) 76 (2%) 93 (5%) 90 (2%)

96 (2%) 94 (1%) 110 (4%) 95 (6%) 94 (4%) 77 (3%) 95 (4%) 92 (4%)

89 (4%) 89 (5%) 99 (2%) 87 (7%) 97 (8%) 81 (3%) 91 (1%) 89 (2%)

94 (2%) 95 (2%) 107 (1%) 95 (5%) 99 (1%) 80 (4%) 96 (6%) 91 (3%)

93 (2%) 90 (3%) 107 (2%) 91 (2%) 95 (5%) 84 (3%) 95 (2%) 87 (1%)

93 (14%)

86 (16%)

74 (22%)

90 (3%) 90 (3%) 120 (2%) 88 (20%) 88 (5%) 80 (6%) 90 (5%) 87 (1%) 95 (10%) 87 (18%) 35 (34%) 81 (31%) 99 (1%) 58 (29%) 76 (12%) 98 (1%) 72 (16%) 99 (1%) 99 (1%) 99 (1%)

88 (9%)

96 (2%)

107 (14%)

M.P. Pavageau et al.

Me2Hg Et2Hg Me2Se Me2Se2 Me4Pb Et4Pb Me4Sn Et4Sn PH3 Hg8 Methanol Acetone Dichloromethane Methylethylketone Ethyl acetate Dichloroethane Methylisobutylketone Toluene Ethylbenzene Paraxylene

Naﬁon (þ208C)

Cryogenic trapping for speciation analysis

After cryogenic sampling, the cryogenic traps are usually stored at 21908C in a special cryogenic container ﬁlled with liquid nitrogen. This is to prevent degradation and volatilization of the volatile analytes. The stability of elemental mercury and organometallic species at temperatures below 21908C has been recently reported by Le Gac and Amouroux [39]. In that study, samples were kept at 21908C in a special cryogenic ﬁeld container (Voyageur, L’air Liquide). A gas mixture of Hg8, Et2Hg, Me4Sn and Me2Se in N2 was generated using a permeation chamber (Calibrage, Saint Chalmas, France). The gas mixture was collected at 2 1708C with the air sampler illustrated in Fig. 16.3. The samples were analysed by GC –ICP-MS after 8 days storage at 21908C. Over this period, no signiﬁcant losses were detected. More recently, the cryogenic conservation of PH3 (a very volatile species) was reported by Pavageau et al. [17]. The sampling and analytical procedure and storage conditions were the same as those described above. Samples were kept at 2 1908C and analysed after 8 and 15 days of storage. Regarding the overall method uncertainty, the estimated PH3 losses were not signiﬁcant. Volatile compounds can also be collected in non-rigid chambers, such as Tedlarw bags, and concentrated in the laboratory using a cryogenic preconcentration system [3,4]. Conservation of volatile compounds in Tedlarw bags has also been evaluated for VOCs and VMCs. Conservation of VOCs in Tedlarw bags is widely documented in the literature [40–45], whereas only one study concerning VMCs was found [46]. In this last referenced experiment, various volatile arsenic, tin and antimony species (AsH3, MeAsH2, Me2AsH, Me3As, SnH4, MeSnH3, Me2SnH2, Me3SnH, Me4Sn, BuSnH3, SbH3, MeSbH2, Me2SbH and Me3Sb) have been generated and mixed with humidiﬁed air. The concentrations of VMCs varied from 0.3 to 18 ng/l at 258C. The samples were stored in the dark at 20 and 508C, and the stability of the VMCs was monitored over a period of 5 weeks. Recoveries depended on the species, and it seems that chemical reactions, such as oxidation or demethylation, are more likely to affect the stability of these compounds in the gas samples. They concluded that Tedlarw bags are suitable for sampling VMCs with the condition that the analysis is completed within a few days of sampling to ensure the integrity of the sample. Finally, the stability of PH3 in a Tedlarw bag (PH3 diluted in puriﬁed compressed air) at a concentration of 28 ng/l at room temperature in the dark was recently investigated. The samples were analysed by GC –ICP-MS over a period of 15 days. The results showed that PH3 is stable during this period, with a recovery of 95% [17]. All the studies conducted have demonstrated that either under gaseous conditions or stored at liquid nitrogen temperature, the VMCs are stable over a short period. In both cases, the fact that interactions are limited (in the gaseous state) or when cryofocused (in the solid state) allows separation of the sampling and sample preparation stages prior to their detection. However, in all cases, sample preservation is always kept to the minimum.

519

M.P. Pavageau et al.

16.6

INSTRUMENTATION FOR CRYOGENIC TRAPPING AND SELECTED APPLICATIONS

Sample preparation for VMCs requires a fully automated handling of the phase transfer of the analytes. Cryofocusing devices have been developed and automated for maximum performance and reproducibility. These devices, and their automation, are an essential part of the sample preparation procedure when analyzing trace metals in the gaseous state together with more abundant gaseous matrix. Further, most of the volatile compounds are extremely toxic even at very low concentrations (,mg/m3); therefore, an important instrumentation demand exists for the trapping and analysis of these species in industrial hygiene and environmental monitoring. In this last section, a review of some of the analytical devices used for cryogenic trapping and handling of the gaseous analytes of interest is presented. 16.6.1 Cryosampler for determination of industrial and environmental VMCs A temperature regulated cryogenic sampling system was developed in the laboratory for a large array of industrial and environmental applications. A schematic of this gas sampler has already been presented in Fig. 16.3. After the ﬁltration stage, the collected gas is dried by the reversed cryogenic Naﬁonw system mentioned earlier in the text. The cryostat is cooled using a liquid nitrogen ﬂow associated with a heating wire. This process allows the trapping of analytes with precise temperatures ranging between 2 20 and 2 1908C. The gaseous species are sampled by cryogenic trapping on silanized glass wool (Supelco, pesticide grade) packed in a glass tube (i.d. 5 mm, length 19 cm). Two samples can be collected simultaneously. For each sampling tube, the ﬂow can be regulated from 0.1 to 1 Nl/min by a mass ﬂow controller (Aalborg). After collection, the sampling tubes are placed in a PVC tube, sealed with silicon caps and stored at 2 1908C in a dry atmosphere cryocontainer (Voyageur 12, L’Air Liquide, Paris, France). The sampling lines were made of Teﬂon and washed with detergent, soaked one day in 10% nitric acid and rinsed with MilliQ Millipore water (18.2 MV.cm). Prior to use, the sampling tubes were cleaned by heating for 10 min at 3008C in a ﬂow of helium. This air sampler was adapted for air measurements in an urban atmosphere, combustion ﬂue gas and work place environment. For the selected applications, the samples were analysed by low temperature gas chromatography coupled with ICP-MS detection. This technique has an absolute detection limit of about 5 pg, depending on the element monitored. (1) Urban environment. Ambient air measurements of tetraethyllead compounds and elemental mercury were conducted in an urban environment [23]. A ﬁltration cartridge (Millipore) equipped with a quartz microﬁbre ﬁlter (Whatmann) was used to remove the smallest particles from the gas, and the

520

Cryogenic trapping for speciation analysis

gas was dried using an empty U-shaped glass trap held at 2 208C before cryogenic trapping at 21758C. The air was sampled at 0.8 l/min for a total collected volume of 15 l. The detected concentrations ranged between 3 and 15 ng/m3. (2) Combustion ﬂue gas. Combustion ﬂue gas samples were collected directly in the stack after ﬂue gas treatment (electrostatic precipitator). Cryogenic trapping was performed at 21758C using a complex drying system in order to avoid clogging during the sampling step [12] after a quartz ﬁltration cartridge. The outlet of the ﬁlter was connected to a Teﬂon (PFA) sampling tube (i.d. 8 mm, length 1 m), which serves as the inlet to the air sampler drying system. The gas was dried using different drying systems connected in series: ﬁrst, a water trap made of Teﬂon tubing (length 10 cm, i.d. 8 mm) set at ambient temperature; next, two empty U-shaped glass traps held at 2 208C, and ﬁnally, a semi-permeable Naﬁonw membrane dryer tube (length 24 cm, i.d. 2 mm and o.d. 3 mm; Perma pure products, New Jersey, USA) set at 2 108C. The Naﬁonw drying efﬁciency is temperature dependent. In our laboratory, we have evaluated the ﬁeld conditions with ambient air, for the same levels of humidity (8.2% vol.) and using the sampling ﬂows (2 Nl/min) implemented during the sampling campaign. Under these conditions, the dew point of the exhausted air was found to be 2 108C. During the sampling time, variable amounts of gas were collected due to partial clogging of the column, and careful attention should be paid to this problem. These harsh conditions can saturate the drying unit in the presence of extremely humid gas (8.2% vol. water content). Further, the trapping of CO2 will be very important because it is the most abundant gas under these conditions. The collected volume for each sample varied between 3 and 5 l. The GC –ICP-MS analysis could not be performed directly due to the high amount of CO2 trapped at 21758C. A recondensation of the samples at 2808C before sample analysis was needed, as described earlier. Under these analytical conditions, seven volatile metal species were detected in the coal combustion ﬂue gas: one for tin, one for copper, two for mercury and three for selenium [12]. Fig. 16.6 presents typical chromatograms obtained for these species. More recently, cryogenic sampling was performed during combustion of metal-enriched fuel [47]. Samples were directly collected at 2 808C in order to avoid trapping of CO2. Volatile species of mercury and selenium were evident in the ﬂue gas. (3) Industrial hygiene. A cryosampling system, together with LT-GC–ICPMS method, including CO2 trapping, has been developed for the determination of phosphine [17] and arsine (unpublished data) for industrial hygiene purposes. Samples were collected in the fumigation room of a tobacco factory after fumigation and outside the fumigation room during fumigation at 21808C using a Naﬁonw membrane dryer set at 2 108C. The collected volume for each sample varied between 5 and 40 l. The phosphine concentrations were below the concentration limit values for occupational exposure to phosphine [VME (France) and TLV-TWA (USA)]. They were estimated to be about 1 ng/m3 in

521

M.P. Pavageau et al.

Fig. 16.6. Typical chromatograms obtained for volatile species of tin (Sn1), selenium (Se1, Se2 and Se3), mercury (Hg1 and Hg2) and copper (Cu1) detected in coal combustion ﬂue gas.

ambient air at 208C and lower than 10 ng/m3 in the vicinity of the fumigation room during the fumigation process. The fourth application concerns the characterization of organometallic species in landﬁll and sewage gases with a simple cryotrapping device using a U-shaped trap cooled with cryogenic baths. The sampling device is illustrated in Fig. 16.7. In this setup, a second cryotrap may be installed in line after the ﬁrst one in order to check for trapping efﬁciency. (4) Landﬁll and sewage gases. Sampling was performed at 2808C by immersing the cryotraps [U-shaped glass tubes ﬁlled with diatomaceous support (Supelcoport)] into a mixture of acetone and liquid N2. This was necessary to avoid the condensation of CO2 and methane, which leads to perturbation during the analysis conducted by desorption in a low temperature gas chromatograph coupled to ICP-MS. For most compounds, identiﬁcation was accomplished by comparison of retention times with standard species. Where standards were not available, species were identiﬁed using the linear relationship between boiling point and retention time. Semi-quantiﬁcation of

522

Cryogenic trapping for speciation analysis

Fig. 16.7. Simple cryotrapping landﬁll or sewage gas: (A) gas glass tube, i.d. 6 mm, length (acetone/liquid N2), (E) pump

device for sampling of volatile organometallic species in inlet, (B) drying tube (MgClO4), (C) cryotrap (U-shaped 20 cm, ﬁlled with Supelcoport), (D) Dewar at 2808C with gas ﬂow regulation (1 l/min), (F) ﬂow meter and (G) outlet.

the species found was performed using a special cross-calibration procedure with liquid element standard solutions [48]. The results listed in Table 16.6 summarize the measurements on ﬁve municipal waste deposits and ﬁve municipal wastewater treatment plants in Germany. The given concentration ranges are mean values of several measurements, and concentrations are given in ng/m3 gas, with reference to the metal. Where no range is given, the low number is below the detection limit. Sewage and landﬁll gases were also collected in Tedlarw Bags, preconcentrated on chromosorb at 2 808C and analyzed by LT-GC –ICP-MS [3,4].

16.6.2 Cryogenic trapping for speciation analysis Cryofocusing has been also used in applications other than collection of volatile species in air sample measurements. Using cryogenic traps in online systems for trace metal analysis results in high preconcentration factors for the determination of metals in various environmental samples. The main issues that present important similarities to the gaseous handling strategy of the analytes are reviewed below. Many elements can form volatile species and this property is used to extract them from the digestion leachate. The most popular derivatization agents include sodium borohydride (NaBH4) and sodium tetraethylborate (NaBEt4). Some other reagents, such as sodium tetrapropylborate (NaBPr4), sodium tetraphenylborate (NaBPh4) and lithium triethylborohydride (LiBEt3H), have been incorporated over the last few years. The experimental conditions that result in optimal derivatization must be deﬁned in each case, taking into account

523

M.P. Pavageau et al. TABLE 16.6 Volatile organometallic compounds in gases from sewage fermentation and municipal landﬁll Species

Sewage gas (range in ng/m3)

Landﬁll gas (range in ng/m3)

AsH3 MeH2As Me2HAs Me3As Me3Sb Me4Sn BuH3Sn Me2Te Me2Hg Me4Pb Me3EtPbb Me2Et2Pbb MeEt3Pbb Et4Pb Me3Bi D4 D5

21– 133a 6 –121 508 46– 1.935 483 –4.526 22 1 47 16– 57 7 –29 13 – 37 99 1665–24.237 1000 £ 103 –2205 £ 103 234 £ 103 – 600 £ 103

15 3–75513 40– 87000 120–35000 n.q. 12– 1761 1–792 1–5 0,2 0,2 0,6 29– 510 2–927 n.q. n.q.

D 4: octamethylcyclotetrasiloxane; D 5: decamethylcyclopentasiloxane; n.q.: not quantiﬁed. a Cryotrapping not quantitative. b Species identiﬁcation by retention time/boiling point relationship.

the nature of the analytes, the derivatization reagent and the sample matrix. Once generated in the gaseous states, many of the precautions and procedures detailed earlier apply to this very versatile analytical procedure. Figure 16.8 shows a scheme of a versatile automatic online system for metal(loid) speciation analysis of liquid samples. The sample is placed in a round-bottom borosilicate glass reaction vessel. The geometry of the vessel is important in order to obtain quantitative purging of the volatile compounds formed during the reaction. Although the volume of a typical reaction vessel is about 250 ml, the system is also operative with vessels of higher volumes, which facilitates the analysis of very diluted samples. The derivatization reagent is added to the vessel by means of a peristaltic pump, and a controlled ﬂow of inert gas is used to sweep the volatile compounds formed during the reaction from the vessel. A fritted glass junction enhances the contact between the gas and the solution. The volatile forms of the analytes are purged from the vessel and preconcentrated

524

Cryogenic trapping for speciation analysis

Fig. 16.8. Schematic of a versatile automatic on-line system for metal(loid) speciation analysis of liquid samples, based on cryogenic trapping of the volatile analytes generated after an appropriate reaction and their separation by packed gas chromatography. He: helium or other inert gas as argon; F: ﬂow meter; R: reagent; P: peristaltic pump; M: magnetic stirrer; EP: electronic (or pneumatic) pump; D: detector.

into a U-shaped tubular column made of Pyrex glass (typically 30–45 cm length, 6–8 mm i.d.) packed with an appropriate adsorbent and immersed in a cold bath at the desired temperature. A precise control of the temperature of the bath is of outstanding importance, in order to achieve selective trapping of the species of interest and to separate them from the sample matrix. As explained earlier (see Section 16.3), the trapping efﬁciency mainly depends on the temperature of the trap. Selecting this precisely results in high preconcentration rates and a signiﬁcant reduction of interferences. Temperatures as low as 2 1968C can be achieved using baths of liquid nitrogen. Mixtures between other cryogenic liquids and the temperatures obtained in each case have been listed elsewhere in Table 16.2. The analytes are sequentially released from the column to the detector in a stream of an inert carrier gas. If a non-inert material has been selected to ﬁll the trap, improved separation capabilities may be achieved, greater than those obtained by simple differences in boiling points of the analytes. Non-polar silicon-based stationary phases, such as OV 1, OV 3, OV 101, SE 50, SE 54 and SP 2100, have been used in the determination of organometallic compounds of tin, lead, and mercury [49–51]. Lighter loadings, from 3 to 5%, are necessary for methylated arsenic [52] or selenium [53] and for species with higher boiling points, such as butyltin compounds [54]. Handling of analytes in the gas phase results in quantitative introduction of the sample into the detector, signiﬁcantly improving the sensitivity of the technique. Very selective atomic detectors, such as atomic absorption [50,55–57] or ﬂuorescence [58,59], have been hyphenated with such derivatization– preconcentration–separation online setups, providing signals nearly free from

525

M.P. Pavageau et al.

spectral interferences. Multielemental analysis is also possible if techniques such as mass spectrometry [10,60] and atomic emission spectroscopy [61], both with plasma sources (MIP-AED or ICP-MS), are incorporated. Versatility is the most outstanding feature of the system described above. It is possible to select not only the best derivatization reagent in each case, but also the most appropriate adsorbent, the temperature of the trap, the temperature program to warm up the column and, what is more important, the detector to be used. A correct selection of these parameters results in very sensitive and selective methods for metal and metalloid speciation analysis of a large array of matrices. The high preconcentration factors obtained provide very low detection limits, which are usually enough to measure more habitual organometallic forms in most environmental samples. A series of points deserve special care; otherwise, they could give rise to signiﬁcant problems during the analysis. For example, formation of foam during the derivatization reaction is usual in samples with high organic content. Foam should be prevented from reaching the column, if necessary, by adding any anti-foaming agent. Problems may also arise from deﬁcient packing of the column. This should be done in a way that the column is not homogeneously ﬁlled. The column ﬁlled with the adsorbent should be correctly silanized prior to use, in order to deactivate any active sites that may serve as adsorption sites for the analytes, negatively affecting the reproducibility of the analysis. Memory effects may also be observed as a result of a deﬁcient silanization process. The separation capacity of the packed column in this way is limited [62,63]. This is one of the more serious limitations of the technique. Satisfactory separations are usually achieved, however, in the case of analytes with boiling points below 2508C, if the correct adsorbent and warming program are selected. Special care must be paid to detection and elimination of leaks, and all the connectors between components should be made of an inert material, kept to a minimum and warmed, in order to avoid condensation of analytes, memory effects and peak tailing. In some applications, elimination of humidity is mandatory. There are several ways to prevent water reaching the column. The most popular ones are the use of a Naﬁonw drier and cold traps (2 208C) immediately before the cryogenic trap, as used with the direct analyses of gaseous samples. As very selective detectors are normally used, spectral interferences are minimised. The system is not free, however, from chemical interference, which may occur during different steps of the analysis. Online preconcentration systems similar to those described here have been used in the speciation analysis of different matrices (Table 16.7).

526

TABLE 16.7 Speciation analysis of metals and metalloids by volatilization, cryogenic preconcentration and gas chromatographic separation using different detectors (Me: methyl-, Et: ethyl-, Bu: butyl-) Species

Reagent

Detector

Matrix

Reference

Sn

Sn4þ, MexSn(4-x)þ

NaBH4

QF-AAS

[49]

Sn Sn Sn

NaBH4 NaBEt4 NaBEt4

QF-AAS QF-AAS QF-AAS

KBH4 NaBEt4 NaBEt4

GC-FPD QF-AAS AFS

NaBEt4 NaBH4

Hg Hg Hg Hg Hg Hg

n-BuxSn(4-x)þ n-BuxSn(4-x)þ MeSn3þ, Et2Sn2þ, Bu2Sn2þ MexSn(4-x)þ MexPb(4-x)þ Hg2þ, MeHgþ, Me2Hg Hg2þ, MeHgþ Hg2þ, MeHgþ, Me2Hg, Et2Hg Hg2þ, MeHgþ MeHgþ MeHgþ Hg2þ, MeHgþ Hg2þ, MeHgþ Hg2þ, MeHgþ

Natural water, sediment, biological tissue Natural water, sediment Oyster Synthetic solution

Hg Hg

Hg2þ, MeHgþ Hg2þ, MeHgþ

NaBH4 NaBEt4 NaBH4 NaBH4 NaBEt4 NaBH4/NaBEt4 NaBH4 NaBEt4

Sn Pb Hg Hg Hg

[54] [77] [55] [78] [50] [79]

QF-AAS QF-AAS

Natural water Synthetic solution Natural water, biological tissue Biological tissue Biological tissue

AFS AFS AFS QF-AAS QF-AAS QF-AAS

Natural water Biological tissue Sediment Biological tissue Sediment Synthetic solution

[82] [83] [84] [85] [85] [86]

AFS GC-MIP-AED

Biological tissue Natural water

[87] [88]

[80] [81]

527

continued

Cryogenic trapping for speciation analysis

Element

528

TABLE 16.7 (continuation) Species

Reagent

Detector

Matrix

Reference

Hg Hg Hg As

MeHgþ Hg2þ, MeHgþ Hg2þ, MeHgþ As(III), As(V), MMA, DMA As(III), As(V), MMA, DMA As(III), As(V), MMA, DMA MMA As(III), As(V), MMA, DMA MexSn(4-x)þ, Hgþ2, MeHgþ, MexPb(4-x)þ Total As, Sb, Se and Sn As(III), As(V), Se(IV), Se(VI) Hg2þ, MexHg(2-x)þ, Sn4þ, MexSn(4-x)þ, Et4Pb Alkyl species of As, Ge, Hg, Sn Inorganic and methylated Ge, Sb, As

NaBEt4 NaBEt4 NaBH4 NaBH4

GC– ICP-MS QF-AAS AFS QF-AAS

Sediment Biological tissue Natural water Natural water

[89] [57] [59] [90]

NaBH4

QF-AAS

Natural water

[56]

NaBH4

QF-AAS

Synthetic solution

[91]

NaBH4 NaBH4

GP-Spec QF-AAS

Synthetic solution Natural water

[92] [56]

NaBEt4

GC-MIP-AED

Natural water

[61]

NaBH4

GP-Spec

Natural spiked water

[93]

NaBH4

QF-AAS

Synthetic solution

[94]]

NaBH4/NaBEt4

GC-MIP-AED

Synthetic solution, soil

[95]

NaBH4

ICP-MS

Natural water

[26]

NaBH4

ICP-MS

Natural water

[96]

As As As As Multielement Multielement Multielement Multielement

Multielement Multielement

M.P. Pavageau et al.

Element

Cryogenic trapping for speciation analysis

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33

J. Feldmann, R. Gru¨mping and A.V. Hirner, Fresenius J. Anal. Chem., 350 (1994) 228. J. Feldmann and A.V. Hirner, Int. J. Environ. Anal. Chem., 60 (1995) 339. J. Feldmann and W.R. Cullen, Environ. Sci. Technol., 31 (1997) 2125. J. Feldmann, J. Environ. Monit., 1 (1999) 33. I. Devai, L. Felfo¨ldy, I. Wittner and S. Plosz, Nature, 333 (1988) 343. D. Glindemann, U. Stottmeister and A. Bergmann, Environ. Sci. Pollut. Res., 3 (1996) 17. A.V. Hirner, J. Feldmann, E. Krupp, R. Gru¨mping, R. Goguel and W.R. Cullen, Org. Geochem., 29 (1998) 1765. E. Tessier, D. Amouroux, G. Abril, E. Lemaire and O.F.X. Donard, Biogeochemistry, 59 (2002) 183. E. Tessier, D. Amouroux and O.F.X. Donard, Biogeochemistry, 59 (2002) 161. D. Amouroux, E. Tessier, C. Pe´cheyran and O.F.X. Donard, Anal. Chim. Acta, 377 (1998) 241. A.V. Hirner, E. Krupp, F. Schulz, M. Koziol and W. Hofmeister, J. Geochem. Expl., 64 (1998) 133. M.P. Pavageau, C. Pe´cheyran, E. Krupp and O.F.X. Donard, Environ. Sci. Technol., 36 (2002) 1561. J. Dewulf and H. Van Langenhove, J. Chromatogr. A, 843 (1999) 163. Handbook of Chemistry and Physics, 79th edition, Lide D.; CRC Press, Boca Raton, LF, 1998–1999. AFNOR XP X 43-051, Air quality—stationary source emissions—determination of the total emission of heavy metals and other speciﬁc elements, June 2001. AFNOR NF EN 13211, Air quality—stationary source emissions—manual method of determination of the concentration of the total mercury, July 2001. M.P. Pavageau, C. Pe´cheyran, M. Demange and O.F.X. Donard, J. Anal. At. Spectrom., 18 (2003) 323. M.P. Pavageau et al. Unpublished data. Handbook of Chemistry and Physics, 56th edition, Lide D.; CRC Press, Boca Raton, FL, 1975–1976. P.W. Atkins, Chimie Physique. DeBoeck Universite´, Bruxelles, 2000. M.R. Riazi and A.R. Khan, J. Colloid Interf. Sci., 210 (1999) 309. P. Pascal, Nouveau Traite´rale. Tome V, Masson and Cie, Paris, 1957. C. Pe´cheyran, B. Lale`re and O.F.X. Donard, Environ. Sci. Technol., 34 (2000) 27. J. Feldmann, Ph.D. Dissertation, University of Essen, Germany, 1995. J. Feldmann, L. Nae¨ls and K. Haas, J. Anal. At. Spectrom., 16 (2001) 1040. C.M. Tseng, D. Amouroux, I.D. Brindle and O.F.X. Donard, J. Environ. Monit., 2 (2000) 603. J. Namiesnik and W. Wardencki, Int. J. Environ. Anal. Chem., 73 (1999) 269. B. Fernandez, Ph.D. Dissertation, Universite´ de Pau et des Pays de l’Adour, France, 1997. U. Busche, Ph.D. Dissertation, University of Essen, Germany, 1993. Y. Odanaka, N. Tsuchiya, O. Matano and S. Goto, Anal. Chem., 55 (1983) 929. D. Tanzer and K.G. Heumann, Int. J. Environ. Anal. Chem., 48 (1992) 17. W. Reifenhauser and K.G. Heumann, Atmos. Environ., 26A (1992) 2905. M.P. Pavageau, C. Pe´cheyran, V. Dezausiers, E. Krupp, H. Pinaly and O.F.X. Donard, InfoChim. Magazine, 434 (2002) 92.

529

M.P. Pavageau et al. 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52

53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69

530

K.J. Leckrone and J.M. Hayes, Anal. Chem., 69 (1997) 911. Naﬁonw gas sample dryers, Technical note, Permapure Inc., New Jersey, 1995. M.P. Pavageau et al., to be published. Website—http://www.permapure.com/newweb/FAQ.htm K. Hoppstock and H. Emons, Chem. Unserer Zeit, 2 (1999) 95. M. Le Gac and D. Amouroux, unpublished data, 2000. EPA, Sampling of principal organic hazardous constituents from combustion sources using Tedlarw bags, method 0040, December 1996. J.M. Andino and J.W. Butler, Environ. Sci. Technol., 25 (1991) 1644. L.J. Mc Garvey and C.V. Shorten, Am. Ind. Hyg. Assoc. J., 61 (2000) 375. R.L. Seila, W.A. Lonneman and S.A. Meeks, J. Environ. Sci. Health, 2 (1976) 121. Y. Wang, T.S. Raihala, A.P. Jackman and R. St John, Environ. Sci. Technol., 30 (1996) 3115. P.F. Wilson, C.G. Freeman, M.J. McEwan, D.B. Milligan, R.A. Allardyce and G.M. Shaw, Rapid Commun. Mass Spectrom., 15 (2001) 413. K. Hass and J. Feldmann, Anal. Chem., 72 (2000) 4205. M.P. Pavageau, A. Morin, F. Se´by, C. Guimon, C. Pecheyran, E. Krupp, J. Poulleau and O.F.X. Donard, Environ. Sci. Technol., in press. J. Feldmann, J. Anal. At. Spectrom., 12 (1997) 1069. O.F.X. Donard, S. Rapsomanikis and J.H. Weber, Anal. Chem., 58 (1986) 772. S. Rapsomanikis, O.F.X. Donard and J.H. Weber, Anal. Chem., 58 (1986) 35. C.M. Tseng, A. de Diego, H. Pinally, D. Amouroux and O.F.X. Donard, J. Anal. At. Spectrom., 13 (1998) 755. S.C. Apte, A.G. Howard and A.T. Campbell, Environmental Analysis Using Chromatography Interfaced with Atomic Spectrometry. Ellis Horwood, Chichester, 1989, 258. A.G. Howard, Environmental Analysis Using Chromatography Interfaced with Atomic Spectrometry. Ellis Horwood, Chichester, 1989, 318. L. Randall, O.F.X. Donard and J.H. Weber, Anal. Chim. Acta, 184 (1986) 197. F.M. Martin and O.F.X. Donard, Fresenius J. Anal. Chem., 351 (1995) 230. J.Y. Cabon and N. Cabon, Fresenius J. Anal. Chem., 368 (2000) 484. I. Va¨lima¨ki and P. Pera¨ma¨ki, Mikrochim. Acta, 137 (2001) 191. C. Pe´cheyran, D. Amouroux and O.F.X. Donard, J. Anal. At. Spectrom., 13 (1998) 615. T. Stoı¨chev, R.C.R. Martin, D. Amouroux, N. Molenat and O.F.X. Donard, J. Environ. Monit., 4 (2002) 517. C. Pe´cheyran, C.R. Quetel, F.M.M. Lecuyer and O.F.X. Donard, Anal. Chem., 70 (1998) 2639. M. Ceulemans and F.J. Adams, J. Anal. At. Spectrom., 11 (1996) 201. O.F.X. Donard and R. Pinel, Environmental Analysis Using Chromatography Interfaced with Atomic Spectrometry. Ellis Horwood, Chichester, 1989, 189. O.F.X. Donard and F. Martin, Trends Anal. Chem., 1 (1992) 47. B. Radziuk, Y. Thomassen, J.C. Van Loon and Y.K. Chau, Anal. Chim. Acta, 105 (1979) 255. W.R.A. De Jonghe, D. Chakraborti and F.C. Adams, Environ. Sci. Technol., 15 (1981) 1217. C.N. Hewitt and R.M. Harrison, Anal. Chim. Acta, 167 (1985) 277. A. Bzezinska, J.V. Loon, D. Williams, K. Oguma, K. Fuwa and I.H. Haraguchi, Spectrochim. Acta, 38 (1983) 1339. A. Paudyn and J.C. Van Loon, Fresenius Z. Anal. Chem., 325 (1986) 369. N. Bloom and W. Fitzgerald, Anal. Chim. Acta, 208 (1988) 151.

Cryogenic trapping for speciation analysis 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96

D. Amouroux, J.C. Wasserman, E. Tessier and O.F.X. Donard, Environ. Sci. Technol., 33 (1999) 3044. D.S. Ballantine and W.H. Zoller, Anal. Chem., 56 (1984) 1288. S.G. Jiang, H. Robberecht and F. Adams, Appl. Organomet. Chem., 3 (1989) 99. S.G. Jiang, H. Robberecht and F. Adams, Atmos. Environ., 17 (1983) 111. M.P. Pavageau, unpublished data. O.F.X. Donard and J.H. Weber, Nature, 332 (1988) 339. J. Feldmann, E. Krupp, D. Glindemann, A.V. Hirner and W.R. Cullen, Appl. Organomet. Chem., 13 (1999) 739. J.S. Han and J.H. Weber, Anal. Chem., 60 (1988) 316. J.M. Liu, G.B. Jiang, Q.F. Zhou and Z.W. Yao, Anal. Sci., 17 (2001) 1279. N. Bloom, Can. J. Fish. Aquat., 248 (1989) 149. R. Fisher, S. Rapsomanikis and O. Andreae, Anal. Chem., 65 (1993) 763. R. Puk and J.H. Weber, Anal. Chim. Acta, 292 (1994) 175. R. Ritsema and O.F.X. Donard, Appl. Organomet. Chem., 8 (1994) 571. E. Saouter and B. Blattmann, Anal. Chem., 66 (1994) 2031. M.A. Morrison and J.H. Weber, App. Organomet. Chem., 11 (1997) 761. C.M. Tseng, A. de Diego, D. Amouroux, F. Martin and O.F.X. Donard, J. Anal. At. Spectrom., 12 (1997) 743. A. de Diego, C.M. Tseng, T. Stoı¨chev, D. Amouroux and O.F.X. Donard, J. Anal. At. Spectrom., 13 (1998) 623. A.A. Heller and J.H. Weber, Sci. Total Environ., 221 (1998) 181. S. Slaets and F.C. Adams, Anal. Chim. Acta, 414 (2000) 141. L. Lambertsson, E. Lundberg, M. Nilsson and W. Frech, J. Anal. At. Spectrom., 16 (2001) 1296. A.G. Howard and S.D.W. Comber, Mikrochim. Acta, 109 (1992) 27. A.G. Howard and C. Salou, J. Anal. At. Spectrom., 13 (1998) 683. J. Sanz, M.T. Martinez, M. Plaza and M.P. Clavijo, Anal. Chim. Acta, 381 (1999) 331. S. Cabredo, J. Galban and J. Sanz, Talanta, 46 (1998) 631. U. Pyell, A. Dworschak, E. Nitschke and B. Neidhart, Fresenius J. Anal. Chem., 363 (1999) 495. R. Reuther, L. Jaeger and B. Allard, Anal. Chim. Acta, 394 (1999) 259. M.J. Ellwood and W.A. Maher, J. Anal. At. Spectrom., 17 (2002) 197.

531

This page intentionally left blank

Chapter 17

Biotrapping as an alternative to metal preconcentration and speciation Yolanda Madrid and Carmen Ca´mara

17.1

INTRODUCTION

Currently, the need for ultra-trace metal determination in complex matrices such as those of the environment, food, biology, etc., is signiﬁcantly increasing. This task usually requires highly sensitive, sophisticated and expensive analytical techniques such as ICP-MS, ETV-ICP and HG-ICP-MS. The use of a prior preconcentration step based on complex formation followed by solvent extraction, solid-phase extraction and knotted reactors, or co-precipitation, electro-deposition, etc., offers an alternative which has been widely evaluated [1]. As an example, the determination of metals at ultra-trace levels in sea water illustrates the difﬁculty of achieving accurate results even using ICP-MS or HR-ICP-MS techniques. The high salt content of the sample makes it necessary to dilute the sample prior to analysis, resulting in a low concentration of analytes. The use of chelating agents, adsorbents and other chemical agents, either in batches or on-line, has been used to separate and preconcentrate a large range of metals. Recently, metal preconcentration by living organisms, such as algae, fungi, bacteria, yeast, etc., has offered an attractive alternative to other chemical adsorbent procedures because of its low cost, high accumulation capacity, large quantities of microbiological waste production and the large variety of microorganisms available [1]. One of the reasons why the bio-absorption ability of living organisms is higher than that of chemical adsorbents is due to the many functional groups available (amine, hydroxyl, carboxyl groups, phosphate, and sulphydryl groups) to bind the metals and the high apparent diffusion coefﬁcients. The reported Cd and Pb values for a brown algae were 20- to 50-fold higher, respectively, than those obtained using strong-acid resins like Dowex 50 X8 [2]. Biomass metal uptake can take place either by using the organisms in an active mode (bioaccumulation, bioconcentration) and/or in a passive mode (biosorption), which implies that both living and dead organisms can be used. Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

533

Y. Madrid and C. Ca´mara

Analytical applications of living organisms extensively used for technological purposes, have been very limited for metal preconcentration and speciation in spite of their attractive characteristics: † † † †

They are very inexpensive and easily available. They have a wide diversity of metal-ion binding sites. They are highly selective for metal ions and/or chemical forms. Their activity is less subject to interference by high concentrations of alkali and alkaline earth metals than conventional compounds such as ionexchange resins.

The biosorption process depends not only on the type of living organism and analyte, but also on the concomitants. For instance, some biosorption studies of Cr(VI), Cu(II) and Cd(II) by Ryzopus arrithus showed that Cr(VI) and Cu(II) signiﬁcantly inhibited the biosorption of Cd(II), while the Cr(VI) þ Cu(II) þ Cd(II) combination showed synergistic interactions for Cr(VI) biosorption. The fact that the uptake process may also depend on the chemical form of the analyte (oxidation state, organic or inorganic forms, etc.) can be useful for speciation purposes. The biosorption efﬁciency may be affected by many factors, including the concentration of the analytes, amount of biomass, pH, etc., and biosorption can be achieved using both activated and inactivated substrates, as illustrated in Fig. 17.1. Experiments on biosorption of Hg(II) and Cd(II) in aqueous media conducted with both live and inactivated fungal Phanerochaete chryosporium have shown that the ion uptake increases as the concentration of analytes in the aqueous media increases. The biosorption efﬁciency was maximal within the 5.0 –6.0 pH range and 15–458C temperature range [3]. Living and dead organisms can be used as biosorbents with similar efﬁciency in both free and immobilised systems, as illustrated in Fig. 17.2. The most common biological substrates used for analytical purposes (sampling, preconcentration and speciation) are algae, fungi, yeast, plants, cells, etc. For on-line analytical applications, the biomass should be immobilised on an appropriate support and placed in a minicolumn. The uptake efﬁciency or retention capacity can be affected by the immobilisation process. The main

Fig. 17.1. Parameters affecting biosorption efﬁciency.

534

Biotrapping as an alternative to metal preconcentration and speciation

Fig. 17.2. Main biological substrates and their analytical applications and working procedures.

biological substrates and their analytical applications and working procedures are illustrated in Fig. 17.2. This chapter focuses on the uptake mechanisms, working procedures as well as analytical and technological applications of biomass. 17.2

GENERAL CHARACTERISTICS OF BIOLOGICAL SUBSTRATES

17.2.1 Algae Algae can be considered as microscopic organisms that subsist on inorganic nutrients and produce organic matter from carbon dioxide by photosynthesis. The general nutrient requirements of algae are carbon (obtained from CO2 or HCO3 2), nitrogen (generally as NO3 2), phosphorous, sulphur (SO4 22) and trace elements, including sodium, potassium, calcium, magnesium, iron, cobalt and molybdenum. In addition to single cells, algae can grow as ﬁlaments, sheets and colonies. Algae are native to a vast array of fresh water and marine environments, and can be grown in large quantities with relative ease. The ability of algae to accumulate trace metals by biosorption has been known for some years. Thus, they have been used in water treatment systems. When waters containing metal ions are passed through a body of water where algae are growing, the efﬂuent waters have a lower metal ion concentration than the inﬂowing waters and alga growth is often suppressed. However, algae

535

Y. Madrid and C. Ca´mara

have only recently been exploited for analytical measurements. Since favourable environments are needed for the growth and reproduction of algae, the use of living algae for metal preconcentration presents obvious limitations. Thus, dead cells may be used for the removal of metal ions from solution under conditions normally toxic to living organisms. Several studies have shown that non-living algae cells can reversibly bind signiﬁcant quantities of metal ions and function like ion-exchange resins. The possible advantage of algae is that the cell wall has many constituents that can be implicated in metal binding, including amine and carboxyl groups from aminoacids and polysaccharides, sulphydryl groups and unmethylated pectins. This range of binding sites means that by altering the elution conditions, different metal ions can be retained preferentially. Different algae exhibit different afﬁnities towards different metals and different metals exhibit different pH-binding proﬁles for a given algae. Metal ions absorbed by algae have been divided into three classes: Class I includes the metal ions (Al3þ, Cu2þ, Cd2þ, Cr3þ, Fe3þ, Ni2þ, Pb2þ, UO2 2þ and Zn2þ) that are bound strongly at near neutral pH values, but are not bound and can easily be stripped at pH , 2: Class II ions (PtCl4 22, CrO4 22 and SeO4 22) are bound strongly at low pH and weakly above pH 5. Class III ions (Agþ, Hg2þ and AuCl4 2) are the most strongly bound of all metal ions, and the binding is independent of pH. Some class II and III metal ions are reduced to elemental metal on the biomass surface; for example, Au3þ is rapidly reduced to Auþ and eventually to elemental gold [4]. The algae most commonly used for metal preconcentration are Chlorella vulgaris [5], Selenestrum capricornutum [6] and Stilloccocus bacillaries [7], either in batch or immobilised in different supports. 17.2.2 Bacteria Bacteria are single-celled prokaryotic microorganisms that may be shaped as rods (bacilli), spheres (cocci) or spirals (vibrios, spirilla, spirochetes). Bacteria cells may occur individually or grow as groups ranging from two to millions of individual cells. Most bacteria fall into the size range of 0.5 –3.0 mm. The metabolic activity of bacteria is greatly inﬂuenced by their small size. Their surface-to-volume ratio is extremely large, so that the inside of a bacterial cell is highly accessible to a chemical substance in the surrounding medium. Therefore, bacteria bring about very rapid chemical reactions compared to those mediated by larger organisms. Bacteria also excrete exoenzymes that break down solid food material into soluble components which can penetrate bacterial cell walls. The cell envelopes of the bacteria are multilayered. The inner layer is the cytoplasmic membrane composed of protein and phospholipids. The identity of the external layers depends on the bacterium. The inner membrane is in contact with a layer of peptidoglycan (a network of polysaccharides, cross-linked by short peptides).

536

Biotrapping as an alternative to metal preconcentration and speciation

Bacteria play an important role in the modiﬁcation, activation and detoxiﬁcation of heavy metals. However, they may themselves be subject to metal toxicity. Bacteria are generally the ﬁrst organisms to be affected by discharges of heavy metals into the environment. Obviously, some heavy metals are more toxic to bacteria than others. In a decreasing order of afﬁnity, this series is Fe3þ, Hg2þ, .Cu2þ, Al3 þ , . Ni2þ, Pb2þ, . Co2þ, Zn2þ . Fe2þ, Cd2þ . Mn2þ . Mg2þ . Ca2þ . Liþ . Naþ . Kþ [8]. The effect of heavy metals is also highly dependent on their chemical form. The major factor determining the toxicity of heavy metals to bacteria is the extent to which they penetrate the cytoplasm. Many bacteria appear to have the capacity for adsorbing metals in the outer layers of the cell. Metals may also be prevented from entering the cell by the formation of complexes or chelates with several metal-binding agents. Apart from the role in immobilising heavy metals, bacteria may affect their environment [9]. Some bacteria are thought to guard against mineral deﬁciencies by providing themselves with chelating agents which may aid the absorption and conservation of essential trace elements. Some of these compounds which are released into the medium in conditions of iron deﬁciency are the so-called siderophores [10]. Under oxidising conditions and physiological pH, Fe3þ is favoured over Fe2þ and so the stabilised iron ferric is easily quelated. In the hard and soft classiﬁcation of ions, Fe3þ is hard and therefore forms stable complexes with harder ligands such as oxygen. The d5 electronic conﬁguration of Fe3þ favours octahedral coordination with poly-oxo chelates. It is therefore not surprising that the most stable ferric ion chelates are based on functional groups like catechol, hydroxamate and a-hydroxyketone. Most natural siderophores and synthetic iron chelators contain such structures, thereby achieving a strong chelate effect as hexadentate ligands. Some naturally occurring iron chelates are desferrioxamine, enterobactin, rhodotorulic acid and desferrithiocin. The ferrioxamines are a group of structurally related polyhydroxamate siderophores produced by genera of the Actinomycetales. Desferrioxamine B (desferrioxamine, deferoxamine, desferal and DFO) is secreted in signiﬁcant quantities by Streptomyces pilosus under iron-limiting conditions and forms a hexadentate iron chelate with log b ¼ 35:9: This compound has been introduced commercially for the clinical treatment of thalassemia. Other siderophores are rhodotorulic acid produced by Rhodotorula pilimanae and enterobactin produced by Escherichia coli and Salmonella typhimmurium. 17.2.3 Fungi Fungi are non-photosynthetic organisms. Most frequently they possess a ﬁlamentous structure. The morphology of fungi covers a wide range. Some fungi are as simple as microscopic unicellular yeast, whereas other fungi form large, intricate toadstools. Fungi are aerobic (oxygen requiring) organisms and

537

Y. Madrid and C. Ca´mara

generally have a better capacity than bacteria to thrive in more acidic media. They are also more tolerant than bacteria of higher concentrations of heavy metal ions. For metal removal and recovery, dead fungi biomass seems to offer several advantages: it is not subject to metal toxicity or adverse operating conditions and needs no nutrient supply, and the recovery of surface-bound metals may be relatively simple by non-destructive treatment [11]. Yeast is more widely used due to its easy growth, non-hazardous nature, considerable tolerance towards metals and high cell-binding capacity. Yeast grows in a pH range from below pH 2 to above pH 9, with an optimum usually from 4 to 7. The temperature range for the growth of most types of yeast is also broad (5 –358C). Further, yeast can be obtained cheaply as a by-product from many industrial sources. Yeast has cellular walls that play an important role in metal accumulation. The main structural component of cell walls is either chitin or chitosan, which are polymers of N-acetylated or non-acetylated glucosamide, respectively. Other ligand groups may be phosphate, carboxyl, amine and hydroxy [12]. Sacharomyces cerevisiae is a type of yeast most commonly used for metal accumulation. S. cerevisiae is an inexpensive, readily available source of biomass, since it is obtained as a waste product of industrial fermentation activities. It also works over a wide range of experimental conditions with a minimum risk of infection and contamination. S. cerevisiae previously immobilised in calcium alginate, polyacrylamide, glutaraldehyde, agar or cellulose acetate has been used for the accumulation of metal. The ﬁlamentous fungus Rhizopus arrhizus has proved to be very useful in the removal of heavy metals, especially radioactive elements such as uranium and thorium. The recovery of uranium from efﬂuents from uranium mines and nuclear waste processing plants has been the most common application of R. arrhizus [13]. The cell wall of fungi contains amino or non-aminopolysaccharides and a high proportion of chitin. 17.3

UPTAKE MECHANISMS

The biosorption mechanisms are very complex and not always well understood. Several processes are involved in metal retention and accumulation in microorganisms (Fig. 17.3): † †

Adsorption of metal ions to cell surface or cell membrane (surface binding) through electrostatic interactions or complex formation with one or more functional groups (Fig. 17.3) Formation of metal precipitates by reaction with certain microbially produced anions (Sy, PO4 32)

Organisms contain binding groups with various charge distributions and geometries and may therefore selectively bind certain metal cations. The most common ionised groups on the cell surface are carboxylate, hydroxyl and phosphate. Nitrogen in uncharged groups, such as peptides, may act as a ligand to complete the coordination number requirements of the metal ion.

538

Biotrapping as an alternative to metal preconcentration and speciation

Fig. 17.3. Mechanisms and functional groups involved in the accumulation process.

Metabolism-independent biosorption is generally rapid and affected by pH (protonation of functional groups) and constant over a modest temperature range (0 –348C) [14]. Gardea [15] evaluated the pH effect on copper ion uptake by alfalfa and the increasing uptake with pH increases suggested that an ion-exchange mechanism was taking place. This trend in pH dependence also suggests that the actual binding sites may be carboxyl groups. At pH higher than 3 –4, the carboxyl groups are deprotonated and negatively charged. Consequently, the attraction of positively charged copper ions would be enhanced. However, at pH lower than 3, the carboxyl groups are protonated and, consequently, the attraction of positively charged copper ions would be inhibited. Kuyucat [16] has reported that at pH levels higher than 1, the functional group can occasionally be responsible for the binding. This is the case for gold retention by a bacterial surface in which the carbonylic and carboxylic groups are responsible for binding at low pH whereas the amino groups participate only at high pH. The pH may affect the binding process due to protonation of the anionic groups. Majidi and Holcombe [17] and Madrid et al. [18] proposed that the adsorption of Cu by algae, or of methylmercury by yeast, takes place through covalent bonds due to the negligible pH effect on the uptake efﬁciency. When living organisms act as ion-exchange materials, the equilibrium process is very fast (a few minutes), and pH-dependent and accumulated metals are easily stripped off with acids. These factors suggest that metals are only retained outside the cell wall. However, if longer incubation times are needed in the retention process, the active functional groups are inside the membrane. †

Metal translocation into the cell with subsequent transformation.

Many of the cellular homeostatic mechanisms are linked to solute transport due to the metabolic energy required in the process. Chemical species might be

539

Y. Madrid and C. Ca´mara

retained in living bacterial cells through an active process, which is assumed to be more selective. Hydrophobic compounds in solution dissolve in the cytoplasmic membrane, diffuse through it and dissolve in the cell’s inside compartment. Translocation of the cations across the wall and membrane into the cell is usually a slow, metabolism-dependent process. Essential metal ions have a speciﬁc transport system associated with their uptake that may involve channels or pores in the membrane. These transport pathways can also be used for foreign metal ions if their size and chemistry are similar to those of a natural cation. Neidhart [19] proposed a very complex mechanism for Cr(VI) biosorption by erythrocytes through the anion-permeation channel. Erythrocytes have an afﬁnity for Cr(VI) but not for Cr(III). Phosphate transport systems are responsible for translocation of arsenate and vanadate anions. Some toxic species are translocated in the same way as essential metals. Cadmium has the same uptake system as manganese, and talium is incorporated through certain potassium transport systems in bacteria and eukaryotic microbes. Metallothioneins and similar proteins play an active role in metal uptake by binding heavy metals such as Cd, Zn, Cu, Au, Hg, etc. They are also active in metal detoxiﬁcation, homeostasis and possibly metal transfer to apometalloproteins. †

Volatilisation of metals by biotransformation.

Some metallic and metaloid species (Pb, As, Sn, Hg, Tl) undergo biomethylation in the environment. The methylation of Hg(II) takes place, as shown in Eq. (17.1), through methylcobalamin. Hg2þ þ CH3 Co ! Hgþ CH3 ! HgðCH3 Þ2

ð17:1Þ

However, organic mercury compounds may also be volatilised by conversion to elemental mercury by cells resistant to the toxic effect of Hg compounds (enterobacteria, mycobacteria, pseudomonas, etc.) which have the enzyme organomercurialyase, responsible for converting organomercury compounds to Hg2þ and then reducing them to elemental mercury by the enzyme mercuryreductase. Other microorganisms have the ability to synthethize compounds with a high propensity to bind metals, such as iron-binding siderophores or metallothioneines, which bind copper and cadmium. Yeast metallothioneines (MT) have been proposed for the recovery of precious metals [20]. †

Biotransformation.

Biotransformation is another mechanism involving living organisms. Several bacteria (E. coli, Flavobacterium sp., Corynebacterium sp., Pseudomas

540

Biotrapping as an alternative to metal preconcentration and speciation

sp.) have demonstrated the ability to preconcentrate and biotransform inorganic arsenic into organometallic species. Microorganisms can reduce metal and metalloids to a lower oxidation state. Many of the organisms that catalyse such reactions use metal or metalloids as terminal electron acceptors in anaerobic respiration. The microbial reduction of Cr(VI) to Cr(III) is one of the most widely studied reactions, since a wide variety of heterotrophic organisms are capable of inducing this reduction. Microbes can also indirectly promote Cr(VI) reduction by producing Fe(II) and sulphide which can abiotically reduce Cr(VI) [20]. Signiﬁcant advances in the understanding of microbe –metal interactions have been made in recent years but more research is needed because new and interesting perspectives are anticipated in detoxiﬁcation, bioremediation, etc. 17.4

WORKING PROCEDURES

Living or dead microorganisms can be used in both batch and on-line (packed in a column) systems and can function free and/or immobilised on different supports. Most of the analytical work published on biological substrates used non-immobilised material through a batch procedure. The main advantages of immobilisation are as follows: † † † †

Allows working with “on-line” systems. The substrates can be easily regenerated and reused in several cycles. The use of certain supports increases the maximum capacity. Preconcentration and elution of the analyte is very fast.

However, the main drawback is the increase in the mass transfer resistance due to the polymeric matrix. 17.4.1 Immobilisation Immobilisation of the biomass can bestow ideal size, mechanical strength, rigidity and porosity on biological substrates, along with further protection against chemical attack and structural degradation. The aggregation or immobilisation treatment selected should not affect the surface structure of the cells. Different supports, such as alginate, sol –gel, silicagel, polyvinyl chloride and polyacrylamide gel, have been used to immobilise microorganisms, alginate being one of the most appropriate for analytical purposes. 17.4.1.1 Immobilisation in alginate The use of alginate is one of the simplest methods and it has been widely employed in laboratory and pilot-scale studies.

541

Y. Madrid and C. Ca´mara

Alginate has been used for yeast, fungus, etc. It is a natural polymer and may be converted into hydrogels via crosslinking with divalent calcium ions. It is preferred [3] over other materials because it provides several advantages, including biodegradability, hydrophilicity, the presence of carboxylic groups (enhancing metal ion adsorption compared with other supports) and natural origin. The general procedure for yeast immobilisation in alginate involves the following steps: About 1 g of baker’s yeast is hydrated with 3 ml of HEPES ðpH ¼ 7Þ: The resulting suspension is mixed into the sodium alginate solution (7 ml of 2% w/w) until homogeneous. The alginate yeast suspension is pumped through a syringe needle and dropped into a solution of 0.05 M CaCl2·2H2O where the alginate gels as beads. The drop height from the capillary tip to the solution must be adjusted to obtain spherical beads of an appropriate diameter (Fig. 17.4). After a 4-hour hardening period, the beads are transferred to a 0.005 M CaCl2 solution and, when necessary, stored at 48C prior to use. The bioabsorbent beads can be used in a batch or in a column for on-line preconcentration. The yeast is trapped in the polymeric alginate matrix [21]. A similar procedure was proposed by Kac¸ar [3] to immobilise the fungus P. chrysorporium. The SEM micrographs of fungus-immobilised alginate beads showed them to be completely different from the control alginate beads and a uniform fungal growth was observed on the bead surface, indicating that the immobilisation of basidiospores was not localised. This uniform distribution is an important criterion for the proper biosorption of heavy metal ions over the whole surface area of the fungus-immobilised beads. The amount of entrapped biomass depends on the microorganism, fungus providing about 0.107 g per gram of beads. 17.4.1.2 Immobilisation in silica gel Silica gel has the advantage of non-compressibility and availability in a variety of pore sizes. However, prior immobilisation (by an adsorption mechanism) is

Fig. 17.4. Yeast immobilisation in alginate.

542

Biotrapping as an alternative to metal preconcentration and speciation

necessary to establish the contribution of the silica gel support to the retention of metal ions, by running the metal solution through the column ﬁlled with the silica. Different microorganisms can be immobilised in silica gel by the procedure proposed by Mahan and Holcombe [22] for algae cells. Approximately 200 mg of biomass is mixed with 2 g of silica gel. The mixture is moistened with a minimum amount (1 –2 ml) of deionised water and mixed to homogeneity. Next, the resulting paste is heated in an oven at 1058C for 20 min to remove the water. It is recommended that the last two steps be repeated twice to maximise contact between the microorganism and the silica surface, thereby improving the immobilisation efﬁciency. Once the silica – microorganism briquette removed from the oven is cooled, it should be gently broken and passed through molecular sieves to obtain the desired particle size (Fig. 17.5). This immobilisation procedure is based on an adsorption process and improves the mechanical resistance stability of the substrate. However, in some cases, the heating required in this immobilisation decreases the activity of substrates such as yeast. Silica gel has been used as a support for the immobilisation of algae and bacteria such as E. coli and Pseudomonas putida [22,23]. Other immobilisation procedures for biomass, such as tomato roots in spherical gel beads are proposed by Scott et al. [24]. 17.4.1.3 Immobilisation in sol– gel Sol–gel chemistry is based on the hydrolysis and condensation of metal alkoxides [25]. The reactions are usually described as: SiðORÞ4 þ nH2 O ! SiðORÞ42n ðOHÞn þ nROH xSiOH þ HO – Si ; !xSi – O – Six þ H2 O

ðHydrolysisÞ ðWater condensationÞ

Fig. 17.5. Yeast immobilisation in silica gel.

543

Y. Madrid and C. Ca´mara

and/or xSi – OR þ HO – Sx ! xSi – O – Six þ H2 O

ðAlcohol condensationÞ

Such a description of sol–gel chemistry is far too simple and does not explain why chemical additives have to be added in order to control the growth of inorganic oxopolymers. Livage and Sa´nchez [26] have explained that most sol – gel reactions involve hydroxylated species and can be described as nucleophilic substitutions. The usefulness of sol–gel immobilisation is mainly due to the ease with which sol –gel-derived materials (proteins, enzymes, siderophores, microorganisms) can be prepared, modiﬁed and processed [27]. Trapping is achieved by doping the substrate in the sol –gel prior to its gelation [25] or by using organo-silicon derivatives R–Si (OR0 )3. The sol –gel materials are optically transparent, chemically and mechanically stable and can be prepared in different forms and shapes, being possible to control the pore size and pore distribution by varying the processing conditions, including the types and concentrations of silanes. The chemical reactions that occur during formation of the sol, gel and xerogel strongly inﬂuence the composition and properties of the ﬁnal product. The water-to-silane ratio, the nature and concentration of the catalyst (acid, base, nucleophile) and the alkoxide precursors highly affect the relative rate of the hydrolysis and condensation process. In a typical procedure, TMOS is combined with water and during the sol – gel transformation the sol interconnects to form (on the time scale of seconds to minutes to days to months) a rigid, porous network—the gel. After drying, a sol– gel is formed. While the sol –gel process is incredibly ﬂexible, it is also inherently complex. The preparation of these materials is often empirical and seems to involve a lot of trial and error combined with some chemical intuition. Several papers dealing with immobilisation procedures in sol –gel [25,27, 28] highlight the advantages of the sol –gel encapsulation method over covalent attachment or adsorption: the entrapment is not invasive and preserves the integrity of the entrapped compound; the absence of covalent bonding is important for maintaining stability and reactivity; selectivity is improved by controlling the mean pore size of the silica glass. Furthermore, encapsulated bio-molecules or microorganisms do not leach from the matrix even under harsh washing conditions but, because of the porous nature of the sol –gel, the analytes to be determined can diffuse into the pores and react with the entrapped compound. A standard sol –gel procedure for pyoverdine (a natural ﬂuorescence pigment) encapsulation [29] is shown in Fig. 17.6. After following this entrapment procedure, no leaching was detected when passing through a minicolumn packed with the doped sol –gel different acidic, basic and neutral solutions. Biomass trapping by sol–gel is another way proposed to immobilise substrates such as the siderophore pyoverdine [29] or desferrioxiamine [31].

544

Biotrapping as an alternative to metal preconcentration and speciation

Fig. 17.6. Procedure followed for pyoverdine encapsulation in sol– gel.

17.4.1.4 Controlled pore glass (CPG) immobilisation The immobilisation of chemical reagents, enzymes and microorganisms on CPG is performed by covalent attachment. CPG is an effective insoluble substrate that is fairly inert, and therefore not subject to microbiological degradation or swelling due to varying ionic strengths. It exhibits good mechanical properties in ﬂowing streams and has been widely used as a solid support for the immobilisation of enzymes and other reagents. The efﬁciency of immobilisation on CPG increases in proportion to the pore size used. Immobilisation on CPG requires a ﬁrst cleaning step with HNO3 followed by distilled water. Next, the CPG is activated or silanised with an aminoalkyl agent and then the bifunctional glutaraldehyde is used to cross-link the substrate to the microorganism (Spirulina platensis, yeast, bacteria, etc.). The procedure used by Maquieria et al. [30] for bacteria immobilisation on CPG is as follows: a mass of 0.5 g of CPG is activated by boiling with 5 ml of 5% HNO3 for 30 min. The procedure for bacteria immobilisation after washing the CPG with water and drying at 958C for 2 h is shown in Fig. 17.7. The reaction scheme is:

545

Y. Madrid and C. Ca´mara

Fig. 17.7. Procedure followed for cyanobacteria immobilisation on CPG.

Studies using an electron micrograph of the immobilised cyanobacteria revealed that it may be possible to increase the quantity of immobilised bacteria by improving the immobilisation methodology and, consequently, the absorbable surface of the material, which would increase the absorption capacity. Bacterial cells covalently immobilised on CPG and placed in a minicolumn have provided excellent reproducibility for trace metal enrichment. Although the retention capacity of this material is relatively low, this is not a problem for trace metal preconcentration purposes. 17.5

APPLICATIONS

17.5.1 Analytical applications 17.5.1.1 Metal preconcentration Both free and immobilised biological substrates have been used for several technological purposes; however, only recently have they been exploited as an analytical tool for metal preconcentration and speciation purposes. The published works dealt with batch or on-line preconcentration procedures, by using the substrate with or without immobilisation. In fact, a tendency arose of developing on-line preconcentration procedures by using the immobilised substrate. As previously described, the immobilisation of living organisms offers more advantages; however, the immobilisation of the substrate could reduce the available metal adsorption sites by approximately 40%. Most studies involve the optimisation of experimental parameters such as pH, sample volume, amount of substrate and type of eluent (Fig. 17.1). As a rule, the most critical parameter seems to be pH. The elution of metal with mineral acids is proposed in most procedures (HCl, HNO3). However, in some cases, for complete recovery the addition of a chelating agent is required. One of the drawbacks of using these types of substrates is their short life, which is of special concern in the case of continuous preconcentration processes. Algae has been the substrate most widely used for metal preconcentration, either in batch or in continuous mode. One of the ﬁrst works was published by Darnall et al. [4]. They used C. vulgaris for the accumulation of gold by forming complexes. The same alga was used by Shengjun and Holcombe [5] to concentrate Ni2þ and Co2þ from sea and river water samples. The sorption was constant between pH 6 and 9. Algae were immobilised in different supports for analytical application. The green alga S. capricornutum [6] was covalently immobilised on GPG for on-line preconcentration of Cu, Pb, Zn, Co, Hg and Cd

546

Biotrapping as an alternative to metal preconcentration and speciation

prior to ﬂame atomic absorption spectrometry. The optimum pH values for Cu, Hg and Pb were 7.5, 6.5 and 5.5, respectively, and those for Zn, Co and Cd were 7.5, 8 and 8.5, respectively, and were completely eluted by 100 ml of 0.1 – 1 mol l21. The detection limits were 0.05, 0.2 and 2.0 ng ml21 for Cu, Zn and Cd, respectively. Stiloccocus bacillaris was also immobilised on a silica gel substrate to separate and preconcentrate trace amounts of Pb, Cu, Zn and Cd from different solutions. The same type of immobilisation was used by Caban˜ero et al. [32] for on-line preconcentration of metals such as Cd, Cr, Mn and Pb by using diatomaceous earth. Preconcentration was based on the retention of Cr and Pb at pH ¼ 3 and subsequent elution with 3 mol l21 HCl. The preconcentration factors achieved for Cr and Pb were 150 and 100, respectively, and were applied to analyse water samples. Plant-derived materials have been employed for preconcentration. Lujan et al. [33] used the roots and stems of cattail plants (Typha latifolia), the leaves of young and mature tumble weeds (Salsoa spp.), Spanish moss and alfalfa sprouts (Medicago sativa). The Cu uptake was pH-dependent with a maximum between 4 and 6. Al(III) exhibited a 100% retention efﬁciency at pH 5 with alfalfa. Gardea [15] tested seven different varieties of alfalfa plant tissues for copper accumulation. The copper retention remained unaltered between pH 2 and 6, the uptake being higher in the shoots than in the roots. The recovery of copper was accomplished by treatment with 0.1 M HCl. The same authors [34] used the same substrate for Cr(III), Cu(II), Pb(II), Ni(II) and Zn(II) accumulation. An uptake efﬁciency of 90% was reported for all the elements except Cr(III), where only a 40% accumulation efﬁciency was achieved. Tomato (Lycopersicum esculetum) and tobacco (Nicotiana tobacum) roots were employed by Scott [35] to adsorb Sr from an aqueous solution. The roots were immobilised in carragen gel beads and packed into a small column. The maximum adsorption capacity was within a narrow pH range of between 5.3 and 5.9. The Sr was eluted by lowering the pH (, 2) or by adding concentrated chloride salt solution. Ximenez-Embu´n et al. [36] used Lupinus seeds of different species (albus, hispanicus, luteous and angustifolius) for Cr, Mn, Pb, Cd, Hg preconcentration from waste water using a batch procedure. For all metals, the uptake pH remained constant within the 4–8 pH range. Even though the plants were exposed for a relatively short time to the metal solution, metal concentrations of approximately 2 g kg21 of dry matter were detected in the young lupin plants. Although algae have been the most widely studied organisms in metal preconcentration for analytical purposes, bacteria and fungi have also been successfully employed for preconcentration purposes. Maquieria et al. [30] used the cyanobacterium S. platensis immobilised in CPG for batch preconcentration of Cu, Zn, Cd, Pb and Fe at optimum pH values of 7.0, 8.0, 7.5, 6.0 and 7.0, respectively. The adsorbed metals were eluted in an acid medium using 0.5 – 1.0 mol l21 nitric acid. The enrichment factor obtained was 100 –1000 times. The column retained its capacity for 3 months. Madrid et al. [37] used this bacterium for Cr and Sb preconcentration from environmental waters over a

547

Y. Madrid and C. Ca´mara

wide range of pH. The experimental procedure was conducted in batch mode, providing preconcentration factors of 20 and 40 for Cr and Sb, respectively. Robles et al. [38] used bacterial cells such as E. coli and P. putida to preconcentrate Au and Be, Cd, Se and Hg prior to determination by GF-AAS. The retention was highly dependent on pH. Immobilisation of the bacteria in silica gel improved the selectivity but reduced the number of metal adsorption sites available. The yeast S. cerevisiae has been one of the substrates most commonly applied for analytical purposes. S. cerevisiae has been immobilised in several supports. Covalent immobilisation on CPG has been used for on-line preconcentration of Cu, Zn, Pb and Cd [30]. The accumulation pH was between 6.0 and 7.5. Metal desorption was performed by lowering the pH to values below 2.0. The enhancement factors for Cd, Pb, Zn, Fe and Cu were 250, 125, 2000, 750 and 286, respectively. The lifetime of the column was 4 months without deterioration when stored at 48C. The same substrate was immobilised in sepiolite [39] for on-line determination of Cu, Zn, Cd, Fe and Ni in water by F-AAS. The optimum pH values were 6 for Zn and Cd, and 8 for Cu. The substrate seemed to be stable for at least 20 runs. Table 17.1 summarises the characteristics of some biological substrates for preconcentration purposes. 17.5.1.2 Speciation There is growing interest nowadays in developing simple analytical methods to discriminate among species of different toxicity. Speciation is usually carried out by using chromatographic techniques coupled to sensitive detectors, ICPMS being one of the most attractive. Although biological substrates have been applied mainly in analytical chemistry to preconcentrate metal ions, their selective bonding with several forms of metals of different toxicity was recently found to be an attractive feature to be used for speciation purposes. The use of these substrates for metal speciation requires, in most cases, the application of living cells, because metabolic processes and species transformation are involved. In other cases, the ability of most toxic forms of analytes, such as As(III), Se(IV), Sb(III), CH3Hgþ, to react with thiol groups (metal–thiol bonds) has also been used for speciation purposes. Despite the relatively long lapse of time—10 years—since the ﬁrst publication on this topic, only a few papers describing certain procedures for speciation have been published. The use of living organisms requires multidisciplinary collaboration (chemists, biologists, etc.) due to the need for speciﬁc knowledge, competence and experience in working with biological materials. Table 17.2 illustrates the main characteristics of some biological substrates used for speciation purposes. The most relevant applications published in the literature for speciation refer to the following analytes:

548

TABLE 17.1 Main applications of biological substrates for metal preconcentration Analytes

Type of sample

Remarks

References

Au and complexes Ni2þ and Co2þ

Batch procedure; optimum pH ¼ 3.0 Batch procedures; pH (6–9)

[4] [5]

Algae, Selenestrum capricornutum immobilized in CPG

Cu2þ, Cd2þ and Zn2þ

Natural waters River and sea water samples Natural waters

[6]

Diatomaceous earth immobilized on CPG Roots and stems of Typha latifolia, leaves of Salsoa spp. Spanish moss Alfalfa plant tissues

Cr(III) and Pb2þ

On-line procedure; pHs: 7.5, 6.5, 5.5, 8 and 8.5 for Cu and Zn, Hg, Pb, Co and Cd, respectively; elution with 100 ml 0.1 –1.0 M HCl On-line preconcentration; pH: 3.0; elution with 3 M HCl Batch procedure; Cu optimum pH: 4– 6; Cd optimum pH: 5.0

[34]

Cu2þ and Al3þ

River and drinking waters Natural waters

[32] [33]

Cu2þ, Pb2þ, Ni2þ, Zn2þ Sr2þ

Natural waters Aqueous solution

Batch procedure; optimum pH: 2–6; elution with 0.1 M HCl On-line preconcentration; critical retention pH: 5.3 –5.9; elution by lowering pH , 2

Wastewater samples

Batch procedure; uptake pH 4–8

[36]

Spirulina platensis immobilized on CPG

Mn2þ, Pb2þ, Cd2þ and Hg2þ Cu2þ, Zn2þ, Cd2þ, Pb2þ and Fe3þ

Water samples

[30]

Escherichia coli and Pseudoma putida immobilized on silica gel

Auþ, Be2þ, Cd2þ, Se(IV) and Hg2þ

Water samples

Saccharomyces cerevisiae immobilized on CPG S. cerevisiae immobilized on sepiolita

Cu2þ, Zn2þ, Pb2þ and Cd2þ Cu2þ, Zn2þ, Cd2þ, Fe3þ and Ni2þ

Water samples

On-line procedure; optimum pH from 6.0 to 8.0; elution with 0.5 – 1.0 M HNO3 Batch procedure; optimum retention pH: 6 for Be and Hg, and 5.0 for Cd and Se(IV); elution with 3.5 M HNO3 On-line preconcentration; optimum pH 6.0 – 7.5; elution lowering pH , 2 On-line preconcentration; optimum pH 6 for Zn and Cd and 8 for Cu

Tomato (Licopersicum sculetum) roots, tobacco (Nicotiana tobacum) roots immobilization in carragen gel beads Lupinus seeds

Water samples

[35]

[38]

[30] [39]

Biotrapping as an alternative to metal preconcentration and speciation

Biological substrate Algae, Chlorella vulgaris Algae, C. vulgaris

549

550

TABLE 17.2 Main characteristics of some biological substrates for speciation purposes Analyte

Sample

Retained species

Biological substrate

Remarks

Reference

As

Natural waters

As(III)

S. cerevisiae

[40]

Sb

Natural waters

Sb(III)

S. cerevisiae

Cr

Lake water, synthetic seawater Spiked river water samples Spiked river and seawaters Aqueous solution

Cr(III)

Dealginated seaweed

Cr(III) Cr(VI)

S. cerevisiae immobilized sephiolite Spilvine platensis

Cr(VI)

Spirogyra species

Aqueous solution

Cr(VI)

Tap water, mineral water Waters Waters

Fe(III)

Human erythrocytes, ghost erythrocytes Pyoverdine immobilized on CPG and sol –gel P. putida, E. coli S. cerevisiae immobilized in silica gel

Slight dependence on pH; batch procedure; retention and preconcentration of the most toxic species 608C, 30 min retention efﬁciency increases as pH increases On-line; pH (6–7); 0.8 ml/min; Cr(III) preconcentration On-line; Cr(VI) determined as difference between total Cr and Cr(III); highly selective method Batch; both species are retained over a wide pH range Retention capacity increases as pH decreases; slow kinetics (120 min); bioremediation Highly selective for Cr(VI)

Hg

Hg(I), Hg(II) CH3Hgþ (100%) CH3Hgþ (65%)

Se

Waters

Se(IV)

Surface waters Water

Se(VI) Selenocystamine (100%) (living and lyophilized cells) Se –Met, Se – Urea

Spiked tap water

S. cerevisiae immobilized in alginate S. cerevisiae Spirulina major P. putida

P. putida

High selectivity for Fe(III); allows Fe(III) and total inorganic Fe determination Hg(II) pH 1 (E. coli) and pH 3 (P. putida) CH3Hgþ retained in S. cerevisiae. It is stable for 1 month; elution with 0.02 M HCl; Hg retained in silica gel; elution with [HCl] . 0.1 M Easy preparation, minicolumn can be used only for one run Biotrapped Se(IV) is converted into organic selenium Very complex mechanism

Preconcentration depends on growth stage of the substrate

[53] [43] [44] [54] [19] [47] [48] [50]

[49] [21] [48] [51]

[52]

Y. Madrid and C. Ca´mara

Fe

[42]

Biotrapping as an alternative to metal preconcentration and speciation

Arsenic: Arsenic occurs in inorganic and organic forms which exhibit signiﬁcant differences in their metabolism and toxicity. The more common arsenic compounds have been found to have the following descending order of toxicity: As(III) . 70-fold As(V) . MMAs (monomethylarsonic acid) . DMAs (dimethylarsinic acid). Other arsenic compounds are believed to be non-toxic and are tolerated by living organisms. Recently, Smichowski et al. [40] have shown the tolerance of the yeast S. cerevisiae toward As(III) and used this property to separate, in batch mode, the two inorganic arsenic species in solution. A preconcentration factor of 7 was achieved for As(III) when 35 ml of water was processed. The method developed was applied to As(III) and As(V) determination in different kinds of spiked waters with a recovery within the 92–106% range. Antimony: The chemistry and toxicity of antimony are similar to those of arsenic [41]. Antimony may enter the environment through geological activity, including the weathering of rocks, run-off from soils, mining and industrial discharges. It is considered a pollutant of priority interest by the Environmental Protection Agency (EPA). The yeast S. cerevisiae has been shown to have a higher potential for Sb(III) retention than for Sb(V) within the pH range 3 –10 [42]. The afﬁnity to the former increases in proportion to the increase in pH, incubation time and amount of biomass. The pH dependence of Sb(III) biosampling suggests that Sb(III) interacts with yeast ligands through an ion-exchange mechanism rather than through covalent bonds, as occurs with selenium. Quantitative separation of Sb(III) and Sb(V) in an aqueous solution containing 5 mg of each species was achieved by adding 200 mg of baker’s yeast and allowing a 30 min incubation time at 608C. Next, the supernatant was separated from the yeast centrifugate and the amount of Sb(III) and Sb(V) in each phase (yeast and supernatant) was determined by HG-AAS. The differences observed between Sb(III) and Sb(V) could be explained by the different chemical properties (reactivity) which provide different degrees of toxicity. The main advantage of using this substrate for Sb speciation is that the sequestered Sb(III) is the more toxic and less abundant of the species. Furthermore, this speciation procedure allows quantitative separation and independent Sb(III) and Sb(V) determination. Chromium: Chromium is mobilised in the environment by both natural and anthropogenic processes. Of its two main oxidation states, Cr(III) is essential for humans and involved in glucose metabolism, while Cr(VI) is highly toxic, causing kidney tubule necrosis and lung cancer (if inhaled). The dealginated seaweed Ecklonia maxima immobilised in alginate has been used for Cr speciation because of its high afﬁnity for Cr(III), and negligible afﬁnity for Cr(VI). The retention of Cr(III) in dealginated seaweed is highly dependent on pH, being quantitative at pH 6. The functional group(s) responsible for binding Cr(III) at different pH is(are) yet to be identiﬁed, but it has been suggested that carboxyl groups may play a major role in metal

551

Y. Madrid and C. Ca´mara

binding (ion-exchange type mechanism). At low pH values, the carboxyl groups are protonated, thus reducing the available sites for cations. Because of the characteristics of dealginate as a cation exchanger, it can be used for on-line separation –determination of CrO4 22 and Cr(III) in environmental samples. When water containing a mixture of Cr(III) and Cr(VI) is passed through a minicolumn, retention of total Cr can be achieved after Cr(VI) reduction to Cr(III). Cr(III) is stripped with 500 ml of 1 M HCl and the concentration of Cr(VI) can be determined as the difference between total Cr and Cr(III). The preconcentration factor achieved for Cr(III) using an on-line system was about two, a higher number being obtained with an off-line conﬁguration. Another rapid, sensitive and accurate method for the separation and preconcentration of Cr(III) and Cr(VI), based on the biosorption of Cr(VI) at pH 2 and elution with 1 M HCl on S. cerevisiae immobilised on sepiolite, was proposed by Bag˘ [43]. Cr(VI) was determined as the difference between total Cr (after Cr(VI) reduction with 0.5 ml of concentrated H2SO4 and 0.5 ml of ethanol) and Cr(III). The difference in pH uptake between the above-mentioned alga and yeast is due to the action of different groups. The proposed method was applied to the separation of Cr(III) and Cr(VI) in spiked sample solutions in ﬁltered river water. One important feature of the system is its high selectivity (the ratio of interfering ions to chromium was 20 for Zn2þ, Cu2þ, Cd2þ, Ni2þ and Fe3þ, and 100 for Naþ, Kþ, Ca2þ and Mg2þ without signiﬁcantly affecting chromium retention). The system can be used for up to 20 runs. The relative error was lower than 5% for both Cr(III) and Cr(VI). The cyanobacterium S. platensis and the plant-derived material Phaseolus are other suitable substrates for chromium speciation. Their suitability has been demonstrated by Madrid et al. [44] for preconcentrating Cr(III) and Cr(VI) from synthetic sea and river water. The vegetable substrate Phaseolus is highly promising for speciation purposes. The biosorption of Cr(VI) from solutions by the green alga Spirogyre was evaluated by Gupta et al. [45] using batch experiments. This substrate has been proposed for Cr(VI) removal from solutions. The maximum chromium removed was found to be 14.7 £ 103 mg metal kg21 of dry weight biomass at pH 2 in 120 ml with 5.0 mg l21 initial concentration. This absorption capacity is much higher than that of most other types of sorbents reported earlier. The covalently immobilised alga Chlamydomonus has been shown to have a high afﬁnity for Cr(VI) [6] and has been used in an on-line system. As the presence of anions seriously interferes with Cr(VI) determination, the authors proposed the use of the standard addition method. The presence of a 10-fold excess of Cr(III) does not interfere with Cr(VI) determination. Neidhart et al. [19] used red blood cells immobilised for speciﬁc sampling of chromate even at high levels of Cr(III). Chromate is transported through the cell membrane via the bicarbonate channel since the erythrocyte membrane is not a transport channel for Cr(III).

552

Biotrapping as an alternative to metal preconcentration and speciation

The immobilised erythrocytes were used in batch mode and a quantitative Cr(VI) retention at pH 6.5 was achieved after a 60 min incubation period at 378C. Under these conditions, less than 0.5% of the Cr(III) ions that were adsorbed on the cell membranes were found in the erythrocyte pellet. Chromate ions are intracellularly reduced to Cr(III) by reductants such as glutathione at a concentration of 1.2– 2 mmol l21, ascorbate, cysteine and other substances which are present at much lower concentrations in the cell. After reduction, the chromate incorporated cannot leave the cell as Cr(III), since human erythrocytes do not provide a transport channel for Cr(III). The Cr(VI) gradient maintained by the intracellular reduction of Cr(VI) leads to nearly total accumulation of the chromate by the human erythrocytes. After Cr(VI) biosampling, the mixture was centrifuged and the supernatant removed. The erythrocyte pellets were haemolysed with Triton X-100 and the chromium content was determined by GF-AAS. It is important to emphasise that the immobilisation of the cells by gel entrapment did not affect the biological activity against the chromate. Rohling and Neidhart [46] have evaluated whether the concept behind Cr(VI) accumulation using erythrocytes can be extended to selenite, selenate, arsenite, and arsenate preconcentration using modiﬁed erythrocytes (ghosts) [47], hypothetically lysed, then ﬁlled with the desired media and released by warming. Erythrocytes ﬁlled with ascorbate or cysteine as intracellular reductants are able to accumulate almost all the chromate. Ghosts are comparable to unmodiﬁed erythrocytes as regards their shelf stability, the integrity of the membrane and the functioning of the anion transporter. Therefore, they offer many possibilities for the accumulation of different anions, such as uptake by the cell membrane and intracellular reaction processes that, in principle, could be reduction, oxidation, precipitation, complex formation, sorption or ion exchange with the only restriction being that the reactant and the product must neither leave nor damage the cell. This could be a promising application for speciﬁc accumulation of selenite; however, the adequate intracellular medium needs to be found. Iron: Iron speciation in environmental samples is of interest mainly to distinguish Fe(II) and Fe(III) species. The bacterium Pseudomonas ﬂuorescens is well known for the excretion of pyoverdine (siderophore). Siderophores are characterised by the ability to speciﬁcally complex Fe(III) ðK ¼ 1032 Þ; having a negligible afﬁnity for Fe(II). The application of pyoverdine immobilised in several supports for iron speciation has been extensively evaluated by Barrero et al. [29]. This pigment has been used immobilised on CPG and sol –gel. When immobilised on CPG, its high afﬁnity for Fe(III) allowed the development of a highly sensitive and selective biosensor for continuous Fe(III) determination (by molecular ﬂuorescence). The biosampling ﬂow rate of 5 ml min21 is highly selective and it has been used for Fe determination in mineral water. This pyoverdine immobilised on CPG was later proposed for a ﬂuorescent sensor to determine Fe(III) and total inorganic iron [48] in an on-line system. It can be used for 1000

553

Y. Madrid and C. Ca´mara

determinations without deterioration (10–15 iron determinations per hour) and quickly regenerated by pumping 1 M HCl through the ﬂow cell. The same authors have used pyoverdine immobilised on sol –gel glass for the spectroﬂuorimetric determination of Fe(III). It was applied to the abovementioned samples, the pyoverdine doped sol– gel glass remaining stable for up to 12 months (longer than CPG). Mercury: Mercury is a potential environmental toxicant because of four properties: the strong afﬁnity of Hg(II) and organomercurial compounds for thiol groups, a tendency to form covalent bonds with organic molecules, the high stability of the Hg– C bond, and a strong tendency to maximise bonding to two ligands in a linear stereochemical arrangement. Methylmercury is one of the most toxic species (100 times more toxic than Hg(II)) and it is present in the environment as a result of bio-methylation of inorganic mercury. Certain bacteria (P. putida and E. coli) have been widely used for mercury preconcentration and speciation. Aller et al. [49] have described the ability of the above two bacteria to preconcentrate Hg(I) and Hg(II). The retention capacity was pH and bacterial cell type dependent and was not the result of metabolic capacity. The retention of Hg(II) was maximum at low pH (pH 1 and 3 for E. coli and P. putida, respectively), whereas the retention of Hg(I) was favoured at pH 4 and 8 for E. coli and P. putida, respectively. Both inorganic mercury species bound to the bacterial cells were released upon exposure to a high HNO3 concentration (3.6 mol l21). The authors applied the analytical method developed to Hg(I) and Hg(II) speciation in natural water. Some authors have developed easy procedures to selectively determine CH3Hgþ and Hg(II). Madrid [18] used baker’s yeast cells S. cerevisiae to selectively separate methylmercury and Hg(II) using a batch procedure. The fraction of methylmercury bound to the yeast was 100% in all cases (30 min incubation time at pH 7.0 and 378C), whereas Hg(II) had less afﬁnity for yeast cells and remained in solution. The uptake of CH3Hgþ was accompanied by chemical transformation and it was rapidly reduced to more volatile species, such as Hg(I) and Hg0. Microorganisms (bacteria, yeast and other fungi) mediate three transformations of mercury: they reduce Hg(II) to Hg(I), they break down methylmercury or other organomercury compounds and they methylate Hg(II). The above authors have reported that the presence of high chloride levels in the sample accelerated the accumulation of Hg(II) in the cell through the formation of HgCl2 (more toxic than Hg2þ). Later, Pe´rez et al. [50] extended the application of baker’s yeast cells to both on-line speciation and preconcentration of mercury species by their immobilisation on silica gel. Both CH3Hgþ and Hg(II) species were retained when passed through a yeast doped silica gel minicolumn: inorganic mercury on the silica gel and CH3Hgþ on the yeast. Differentiation of CH3Hgþ and Hg2þ was possible through selective elution: the use of 0.02 M HCl selectively elutes the methylmercury with a 100% recovery in the presence of Hg2þ. Hg2þ can be quantitatively eluted with

554

Biotrapping as an alternative to metal preconcentration and speciation

an HCl concentration higher than 0.1 mol l21 and/or chelating agents such as CN2 (5 ml of 0.8 M CN2). In all cases, the recovery was around 90 –95%, the minicolumn being able to accumulate up to 4000 mg Hg2þ/g silica gel and 800 mg CH3Hgþ/g silica yeast before saturation. Preconcentration factors of 15and 100-fold were achieved for CH3Hgþ and Hg2þ, respectively. This method competes with sophisticated hyphenated techniques as a means of differentiating mercury species of different toxicities. The minicolumn could be used for several runs and has the advantage of allowing both preconcentration and separation of inorganic and organic mercury. Based on this work, the same authors [51] have demonstrated that yeast– silica gel columns are able to immobilise and stabilise mercury species. Furthermore, they showed their capabilities for ﬁeld sampling experiments involving sea and river water. Recovery for methylmercury after one month’s storage was similar to that from fresh columns (95 ^ 6%). However, the recovery for inorganic mercury after one month’s storage (70 ^ 3%) was lower than those obtained from fresh columns. The yeast –silica gel columns showed potential not only for ﬁeld sampling but also for CH3Hgþ preservation. The use of yeast –silica gel columns decreases the risk of sample contamination and the problem associated with species transformation and storage. When yeast was immobilised in alginate [42], the methylmercury retention decreased to 60–65%. Although the minicolumns can only be used for one run, their easy preparation (many minicolumns can be prepared in a short time) makes them recommendable in any laboratory for routine environmental analysis. Recently, Kac¸ar et al. [3] proposed the use of basidiosphores of P. chrysporium immobilised in Ca-alginate for Hg(II) preconcentration and removal from aqueous solution. The biosorption of Hg(II) ions by the biosorbents increased as the initial concentration of Hg(II) increased in the medium. The alginate fungus could be regenerated using 10 mM HCl. The biosorbents were rinsed in three biosorption and desorption cycles with negligible decrease (up to 97% recovery in biosorption capacities). Selenium: Selenium is an essential nutrient for humans at a low concentration, while at higher concentrations it becomes toxic for many animals and marine organisms. Selenium is today one of the most popular and studied bioelements because its toxicity and signiﬁcance in the environment depends on both concentration and chemical form. In many environmental matrices this element is present as inorganic selenium, Se(IV) and Se(VI) (toxic species) rather than organic species. Baker’s yeast has also been proposed as a substrate for differentiating Se(IV) and Se(VI) in aqueous samples. Se(IV) is selectively absorbed while Se(VI) remains in solution (pH 3–10) [21]. It was found that absorbed Se(IV) is converted into other selenium species (probably including selenomethionine and selenocysteine). Another batch procedure for Se speciation in natural waters and river sediment samples based on the use of algae (Spirulina major) was proposed for the separation of Se(IV) and Se(VI). This algae has the ability, after being washed with 1 M HCl for 10 min, to sequester Se(VI) at pH 5 while Se(IV)

555

Y. Madrid and C. Ca´mara

remains in the supernatant. The subsequent preconcentration of Se(IV) was carried out at pH 1. The recovery was dependent on the Se absolute amount (96.4% if the Se concentration does not exceed 1.7 mg). Aller and Robles [52] used P. putida for selective selenocystamine (Se-Cystm) uptake, using both living and lyophilised cells in a batch procedure. The percentage of Se-Cystm retained by the living cells was about 100% at low concentration levels (up to 100 ng ml21). The 20- to 80-fold concentration of other organic selenium compounds (such as Se-Methionine or Se-Urea) did not interfere. The retention mechanism is very complex, an energy expending process during cell growth probably being mainly responsible for the uptake process. Based on this mechanism, the same authors [53] proposed the use of P. putida for Se-Met and Se-U speciation. The methodology is based on the fact that the highest retention rate of Se-Met is in the early growth stages, while the highest uptake rates for Se-U occur later. The uptake process was monitored by measuring the selenium content in the biomass as a function of the growth time. The differences of the relative sorption of Se-Met and Se-U by living bacterial cells were applied to the determination of Se-Met and Se-U at trace levels in spiked tap water samples. Finally, a promising application of erythrocyte ghosts ﬁlled with appropriate intracellular reagents for the speciﬁc accumulation of SeO3 22 was proposed by Neidhart (uptake efﬁciencies less than 60%), but the methodology has not yet been fully applied. 17.5.2 Technological applications The vast majority of the applications of the biosorption phenomena exhibited by biological substrates are found in metal reclamation and remediation of industrial and contaminated streams, recovery of precious metals and radionuclides and biomonitoring processes. In fact, the analytical applications mentioned above are, to a certain extent, small-scale applications of these technological uses. Among the most important applications are bioremediation, phytoremediation and biomonitoring. Bioremediation strategies have been proposed as an attractive alternative to conventional methods due to their low cost and high efﬁciency. Microorganisms can remove toxic metals and metalloides from contaminated water and water streams by converting them to forms that are precipitated or volatilised from solution. This is accomplished either by biosorption of metals or enzymatically catalysed changes in the metal redox state [54]. Phytoremediation has been deﬁned as the use of green plants and their associated microorganisms, soil amendments and agronomic techniques to remove, contain or render harmless environmental contaminants. Phytoremediation is expected to be complementary to classical bioremediation techniques, based on the use of microorganisms. It should be particularly useful for the extraction of toxic metals from contaminated sites and the treatment of recalcitrant organic compounds [55].

556

Biotrapping as an alternative to metal preconcentration and speciation

Over the past few decades, the term bioindicator/biomonitor has become more popular. Bioindicators and biomonitors may be considered as organisms (part of an organism or community of organisms) that react to changes in environmental conditions with a change in (a) element concentrations or levels of certain compounds, (b) morphological structures, (c) intracellular and intercellular processes, and (d) dynamics of population [56]. 17.6

CONCLUSIONS

The property of metal accumulation by living organism is useful for trace metal preconcentration purposes. Their selective bonding capacity to different forms of metal is a promising approach for metal speciation. Besides these advantages, the use of these substrates for metal speciation is very scarce. Although the ﬁrst work of this subject appeared more than 10 years ago, only several papers have been published [1]. Considering the multidisciplinary character of this matter, a close cooperation of chemist and biologists is necessary, due to requirements of speciﬁc knowledge, competence and experience in the work with biological material. Automation has been recently included in this ﬁeld as the use of ﬂow procedures provide a better precision and reproducibility, easy operation and low time-consuming. Furthermore, the use of immobilised substrates allows regeneration of the substrates and an increase of their lifetime. REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17

B. Godelewska-Zylkiewicz, Crit. Rev. Anal. Chem., 31(3) (2001) 175–189. H. Seki and A. Suzuki, J. Colloid Interface Sci., 246(2) (2002) 259– 262. Y. Kac¸ar, C. Arpa, S. Tan, A. Denizli, O. Genc and M.Y. Arica, Process Biochem., 37(6) (2002) 601– 610. S.W. Darnall, B. Greene, M. Hosea, R.A. Mcpherson, M. Henzl and M.D. Alexander, in: R. Thompson (Ed.), Trace Metal Removal from Aqueous Solution. Burlington House, London, 1986, pp. 1 –24. J.A. Shengjun and J.A. Holcombe, Anal. Chem., 62 (1990) 1994. H.A.M. Elmahadi and G.M. Greenway, J. Anal. At. Spectrom., 6 (1991) 643. C.A. Mahan and J.A. Holcombe, Spectrochim. Acta, 47B (1992) 1483. A. Albert, Selective Toxicity, 5th edn. Chapman and Hall, London, 1973. R.M. Sterrit and J.N. Lester, Sci. Total Environ., 14 (1980) 5– 17. D.M. Templeton, in: R.A. Goyer and M.G. Cherian (Eds.), Toxicology of Metal. Biochemical Aspects. Springer-Verlag, Berlin, 1995, pp. 305 –331. Y. Sag, Sep. Purif. Methods, 30(1) (2001) 1. D. Brady and J.R. Duncan, Appl. Microbiol. Biotechnol., 41 (1994) 149. M. Tsezos, Z. Georgouis and E. Remoudaki, Biotechnol. Bioeng., 55 (1997) 16. G.M. Gadd, in: R.A. Herbert and A.G. Coods (Eds.), Microbes in Extreme Environment. Academic Press, London, 1986. J.L. Gardea, Solvent Extr. Ion Exch., 14(1) (1996) 119–140. N. Kuyucat and B. Volesky, Biosorption of Heavy Metals. CRC Press, Boca Raton, FL, 1990. V. Majidi and J.A. Holcombe, Spectrochim. Acta, 43B (1988) 1429.

557

Y. Madrid and C. Ca´mara 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56

558

Y. Madrid, C. Cabrera, T. Pe´rez Corona and C. Ca´mara, Anal. Chem., 67 (1995) 750. S. Neidhart, Ch. Herwald, B. Lippmann and B. Straka-Emden, Fresnius J. Anal. Chem., 337 (1990) 853–885. G.M. Gadd, Heavy metals pollutants: environmental and biotechnology aspects. Encyclopedia of Microbiology, 1992, pp. 351– 360. T. Pe´rez, Y. Madrid and C. Ca´mara, Quim. Anal., 20 (2001) 29– 36. C.A. Mahan and J.A. Holcombe, Anal. Chem., 64 (1992) 1933. L.C. Robles and A.J. Aller, Quim. Anal., 15 (1996) 21. C.D. Scott, C.A. Woodward and J.E. Thompson, Enzyme Microb. Technol., 11 (1989) 258 –262. D. Avnir, S. Braun, O. Lev and M. Ottolengui, Chem. Matter, 6 (1994) 1605. J. Livage and C. Sa´nchez, J. Non-Cryst. Solids, 145 (1992) 11– 19. M. Collison, Crit. Rev. Anal. Chem., 29(4) (1999) 289 –311. L.L. Hench and J.K. West, Chem. Rev., 90 (1990) 33. J.M. Barrero, C. Ca´mara, C. Perez-Conde, C. San Jose´ and L. Fernandez, Analyst, 120 (1995) 431– 435. A. Maquieria, H.A.M. Elmahadi and R. Puchades, Anal. Chem., 66 (1994) 1462. L. Ljunggren, I. Altrell, L. Risinger and G. Johansson, Anal. Chim. Acta, 256 (1992) 75– 80. A.I. Caban˜ero, Y. Madrid and C. Ca´mara, Anal. Bioanal. Chem., 244 (2002). J.R. Lujan, D.W. Darnall, P.C. Stark, G.D. Rayson and J.L. Gardea-Torresday, Solvent Extr. Ion Exch., 12 (1994) 803. J.L. Gardea, K.J. Tiemann, G. Ga´mez and K. Dokken, J. Hazard. Mater., 69(1) (1999) 41. C.D. Scott, Biotechnol. Bioeng., 39 (1992) 1064. P. Ximenez-Embu´n, Y. Madrid, C. Ca´mara, C. Cuadrado, C. Burbano and M. Muzquiz, Int. J. Phytoremediat., 3(4) (2001) 369. Y. Madrid, M.E. Barrero and C. Ca´mara, Analyst, 123 (1998) 1593. L.C. Robles, C. Garcı´a-Olalla and A.J. Aller, J. Anal. At. Spectrom., 8 (1993) 1015. H. Bag˘, M. Lake and D.R. Turker, Fresenius J. Anal. Chem., 363 (1999) 224. P. Smichowski, J. Marrero, A. Ledesma, G. Polla and D. Batistoni, J. Anal. At. Spectrom., 15(11) (2000) 1493. S. Maeda, in: S. Patai (Ed.), The Chemistry of Organic Arsenic, Antimony and Bismuth Compounds. John Wiley & Sons, Chischester, 1994. T. Pe´rez, Y. Madrid and C. Ca´mara, Anal. Chim. Acta, 345 (1997) 249. H. Bag˘, A.R. Tu¨rker, M. Lale and A. Tungel, Talanta, 51 (2000) 895. Y. Madrid, M.E. Barrero and C. Ca´mara, Analyst, 123 (1998) 1593. V.K. Gupta, A.K. Shrivastava and N. Jain, Water Res., 35(17) (2001) 4079. O. Rohling and B. Neidhart, Anal. Chem., 71 (1999) 1077. G. Schowoch and H. Passow, Mol. Cell. Biochem., 2 (1973) 197. P. Pulido, J.M. Barrero and M.C. Pe´rez-Conde, Anal. Chim. Acta, 429 (2001) 337. A.J. Aller, J.M. Lumbreras, L.C. Robles and G.M. Ferna´ndez, Anal. Proc., 32 (1995) 511. T. Pe´rez, Y. Madrid, C. Ca´mara and E. Beceiro, Spectrochim. Acta B, 53 (1998) 321 –329. T. Pe´rez, Y. Madrid and C. Ca´mara, Fresnius J. Anal. Chem., 368 (2000) 471– 474. A.J. Aller and L.C. Robles, J. Anal. At. Spectrom., 13 (1998) 469. A.J. Aller and L.C. Robles, Analyst, 123 (1998) 919 –927. Volesky, Hydrometallurgy, 59 (2001) 203. D.E. Salt, R.D. Smith and I. Raskin, Annu. Rev. Plant Physiol. Plant. Mol. Biol., 49 (1998) 643. B. Markert (Ed.), Plants as Biomonitors. Indicators for Heavy Metals in the Terrestrial Environment. VCH, Weinheim, 1993.

Chapter 18

Membrane extraction ˚ ke Jo¨nsson and Lennart Mathiasson Jan A

18.1

INTRODUCTION

In previous chapters, various extraction techniques, such as solvent extraction (LLE), solid phase extraction (SPE) or solid phase microextraction (SPME) have been treated. Membrane extraction can offer a number of advantages over other extraction techniques. There is nearly no need for organic solvents, while maintaining high enrichment factors, unsurpassed cleanup efﬁciency and high selectivity. In addition, the process is easily automated and adapted to various analytical instruments. A nonporous membrane is a liquid or a solid (e.g., polymeric) phase placed between two other liquid phases. One of these phases, the donor (or feed) phase, is the sample to be processed. On the other side of the membrane is the acceptor (or strip) phase, in which the extracted analytes are collected and subsequently transferred to an analytical instrument. This forms a three-phase system, in which the versatile chemistry of LLE can be employed (and extended) in a format that is amenable to automation. The following description will be focused on these techniques and their applications to trace metal analysis. 18.2

MEMBRANE EXTRACTION TECHNIQUES

A number of nonporous membrane techniques have been described for sample preparation in analytical chemistry. The main technique that has been applied to trace metal analysis is supported liquid membrane (SLM) extraction, but microporous membrane liquid –liquid extraction (MMLLE) has also been used for this purpose. Other versions of membrane extraction exist, but are mainly used for the extraction of organic compounds and these have been reviewed elsewhere [1,2]. Typical units for membrane extraction are shown in Fig. 18.1. Usually, they are constructed of two blocks of inert material with a machined groove in each. When the blocks are clamped together with a membrane between, a ﬂowthrough channel is formed on either side of the membrane, the donor channel, Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

559

˚ ke Jo¨nsson and Lennart Mathiasson Jan A

Fig. 18.1. Membrane units for liquid membrane extraction. (a) membrane module with 1 ml channel volume (A ¼ blocks of inert material, B ¼ membrane). (b) membrane module with 10 ml channel volume.

and the acceptor channel. For sample preparation use, channel volumes are in the range 10 –1000 ml. Different types of extraction units based on hollow ﬁber membranes have also been used. In one conﬁguration, the acceptor phase is inside the ﬁber lumen and the donor channel is the volume between the outside of the ﬁber and the inside of a surrounding tube or cylindrical hole. Another conﬁguration involves the use of two hollow ﬁbers, one containing the donor and the other, the acceptor phase. The membrane liquid then occupies the volume outside the ﬁbers. Other versions of liquid membrane conﬁgurations are also known, such as bulk liquid membranes or emulsion liquid membranes. The latter types are less suitable for analytical purposes and not further treated here; for information see Ref. [3]. 18.2.1 Supported liquid membrane extraction (SLM) SLM extraction has been used for quite some time for industrial separations, mainly for the extraction of metal ions from, e.g., wastewater [3,4]. The use of SLM for sample preparation in analytical chemistry was proposed by

560

Membrane extraction

Audunsson [5], who originally developed the technique for the extraction of amines in connection with the gas chromatography detection. The ﬁeld has further been expanded to many organic and inorganic species and is the subject of several reviews [1,2,6–10].

18.2.1.1 Principles of SLM extraction In SLM, the membrane consists of an organic solvent, which is held by capillary forces in the pores of a hydrophobic porous membrane. Typical solvents are long-chain hydrocarbons like n-undecane or kerosene and more polar compounds like di-hexyl ether, di-octyl phosphate and others. Various additives to the membrane phase are often used for the promotion of extraction, as described below. In Fig. 18.2, a generic principle for SLM extraction of metal ions is presented. To the sample is added an extractant (E), e.g., a chelating compound that forms uncharged species (ME) with the metal ions to be extracted. The acceptor channel on the other side of the membrane is ﬁlled with a solution of another chelating compound (C), capable of forming more stable, charged complexes (MC2 or MCþ) with the metal ions. When the sample is pumped through the donor channel, the ME species are extracted into the organic membrane phase. Having diffused through the membrane, the metal ions will be immediately captured at the membrane –acceptor interface as a charged species and therefore, prevented from re-entering the membrane. This is referred to as trapping and it results in a transport of metal ions from the donor to the acceptor phase, from which they are transferred to an analytical instrument after the extraction, either manually or on-line by a ﬂow system. In a typical realization of this scheme, E could be 8-hydroxyquinoline and C could be diethylenetriaminepentaacetic acid (DTPA) [11], but as will be seen below, there are many other possibilities.

Fig. 18.2. Schematic description of the SLM principle for the extraction of a metal ion M with an extractant E and a trapping reagent C. For details see the text.

561

˚ ke Jo¨nsson and Lennart Mathiasson Jan A

If the trapping is virtually complete, practically all the analytes in the acceptor will be in the form of MC2 species. Therefore, the concentration gradient (which controls the mass transfer rate) of the diffusing species (ME) will be practically unaffected by the total concentration of metal in the acceptor phase. This leads to a potentially high degree of concentration enrichment (several hundred or thousand times, depending on volumes and time), when more and more sample is pumped through the donor channel and collected in the acceptor channel. It is obvious that metal ions that do not form uncharged complexes with E in the donor phase will be excluded from the membrane. This also holds for all other charged species (organic or inorganic). Neutral compounds (N) may be extracted, but not enriched, as the concentration in the acceptor phase will never exceed that in the donor. Macromolecules, as proteins, will typically be charged and therefore, rejected. Finally, the extraction rate of uncharged macromolecules will be very low due to their low diffusion coefﬁcients. Thus, under the conditions mentioned, the SLM extraction will be highly selective for the metal ions that fulﬁll the condition of forming an uncharged complex under the donor conditions and a charged complex under the acceptor conditions. These principles provide many possibilities with which to tune the chemistry in the three phases to tailor the enrichment selectivity to different compounds and to various classes of organic analytes, as further detailed elsewhere [1,2,10]. Jiang et al. have described systems that are a combination of FIA and membrane extraction [12]. Those comprise an FIA-type LLE in a ﬂow system where the organic solvent is dispersed into the ﬂowing sample stream. The phases are then separated by a porous hydrophobic membrane and the analytes can be trapped in an aqueous acceptor. These systems are more complicated than SLM systems, but provide high extraction efﬁciency while maintaining a high degree of clean-up. 18.2.1.2 Theory of SLM extraction A thorough treatment of the theory and principles of the SLM process has been presented [13] and the following items are emphasized here: Extraction is usually evaluated as the extraction efﬁciency, E: This is the fraction of analyte amount input to the system that is collected in the acceptor, i.e., E ¼ nA =nI

ð18:1Þ

where nI and nA are the number of moles input during the extraction time and those collected in the acceptor, respectively. The extraction efﬁciency (fraction of analyte molecules that are transferred to the acceptor) is a function of many parameters. These include the magnitude of the partition coefﬁcient of the uncharged analyte species between the aqueous phase and the organic (membrane) phase, the trapping conditions in

562

Membrane extraction

the acceptor, ﬂow rate of the donor, as well as the characteristics and dimensions of the membrane. The inﬂuence of the partition coefﬁcient, K; is somewhat complex. At low values of K; E is small, as the analyte is insufﬁciently extracted into the organic membrane and the mass transfer is limited by the diffusion transfer through the membrane. At intermediate values, the mass transfer is limited by the transport properties in the ﬂowing donor phase, and in this region, the most efﬁcient extraction is obtained. At very high values of K; i.e., for very hydrophobic species, the stripping of analyte into the acceptor phase becomes the limiting factor, and the observed extraction efﬁciency will decrease, because signiﬁcant amounts of analyte will be left in the membrane. It was found [14] that the most efﬁcient extraction is obtained when the octanol –water partition coefﬁcient (as a measure of polarity) of the diffusing species is around 103. The speed of extraction, i.e., the rate of mass transfer from donor to acceptor, is proportional to the concentration difference, DC; over the membrane. With some simpliﬁcations (especially related to activity effects at different ionic strengths) it follows that [13,15] DC ¼ aD CD 2 aA CA

ð18:2Þ

where CD and CA are the concentrations in the donor (sample) and acceptor phase, respectively, and aD and aA are the fractions of the analytes that are in extractable (uncharged) form in the actual phase. Typically, extraction conditions are set up such that aD is close to 1 and aA is a very small value. CA is zero at the beginning of the extraction and increases during the operation, usually to values well over CD : As long as aA is sufﬁciently small, the second term in Eq. (18.2) is negligible and E will be constant during the course of the extraction; so the extracted amount will be directly proportional to the volume that is extracted and also to the concentration of analyte in the sample. This is the preferred situation and it is referred to as complete trapping. If the trapping is not complete, the extraction efﬁciency will decrease with time, leading to less precise quantitation. This was detailed in a recent paper [15]. However, incomplete trapping conditions can, as will be described below, provide a means of measuring free (nonbound) concentrations, as opposed to total concentrations. As could be expected, extraction efﬁciency is highest for very low donor ﬂow rates, and decreases as the ﬂow rate increases (see Fig. 18.3). Therefore, the most efﬁcient extractions are obtained at low donor ﬂow rates, as a low ﬂow rate increases the residence time of an analyte molecule in the donor channel. On the other hand, increasing the donor ﬂow rate also increases the amount of analyte that is introduced into the extraction system and the net result often is an increase in the amount of accumulated analyte in the acceptor during a given time. In practical work, time is an important issue, and it is therefore, often more relevant to maximize the concentration enrichment factor rather

563

˚ ke Jo¨nsson and Lennart Mathiasson Jan A

Fig. 18.3. Extraction efﬁciency, E and enrichment factor Ee (arbitrary units) as functions of a reduced donor ﬂow parameter f; (see text).

than to maximize the extraction efﬁciency. This will lead to larger instrumental signals (e.g. peak areas) in the analysis of the extract and thus to a more timeefﬁcient analysis. The concentration enrichment factor, Ee ; is related to the extraction efﬁciency as Ee ¼ CA =CS ¼ EVS =VA

ð18:3Þ

CS and CA are the analyte concentrations in the extracted sample and the acceptor channel, respectively, and VS and VA are the corresponding volumes. When the donor ﬂow rate is increased, E decreases, as shown in Fig. 18.3, but this is compensated for by an increasing amount of analyte being input to the system. The result is that, under donor-controlled conditions, the enrichment factor typically increases with donor ﬂow rate for a given time, as seen in Fig. 18.3. However, a high ﬂow rate will obviously consume a large volume of sample, which usually is not a problem for the extraction of environmental samples (e.g. river water or similar samples). If the available sample volume is limited (e.g. biological samples), extraction could be better performed at low ﬂow rate to maximize the extraction efﬁciency. Thus, it is not necessary to strive for the maximum value of E; and this parameter should not be confused with recovery. For good quantitative performance, the important issue is to ﬁnd conditions that lead to reproducible values of E and the value of this parameter will be included in the calibration. 18.2.1.3 Equilibrium extraction in SLM If the extraction is conducted for an increased amount of time, the concentration in the acceptor CA increases and eventually the second term in

564

Membrane extraction

Eq. (18.2) will become signiﬁcant and DC decreases. This leads to a lower rate of mass transfer and a decrease in E: When DC has reached zero, all three phases are in equilibrium and the maximum concentration enrichment factor possible is reached. This is given by EeðmaxÞ ¼ ðCA =CD Þmax ¼ aD =aA

ð18:4Þ

With aD < 1 and aA # 0:0005; signifying complete trapping, Ee ðmaxÞ $ 2000; it is possible to enrich the analyte at least several hundred times until the E is appreciably inﬂuenced. The maximum value is not attained in practice. However, if the trapping conditions are set to aA ¼ 0:1; the maximum enrichment factor is only 10, which is easily reached, and the system is at equilibrium. The concentration of the acceptor is then 10 times higher than that in the donor and will not increase if more of the sample is passed through the donor channel. Equation (18.4) was experimentally veriﬁed [15] for the extraction of organic bases and also for organic acids and for copper extraction using diethylhexyl phosphoric acid (DEHPA) (unpublished). These acceptor-controlled conditions can be used to measure equilibrium fractions of the analytes taking part in secondary equilibria. One example is the binding of different environmental pollutants to humic compounds, particles, etc., in natural waters. By setting up acceptor-controlled extraction conditions and carrying out the extraction until equilibrium is attained, the analysis of the acceptor directly gives a known factor times the free equilibrium concentration in the sample. As described above, the factor is aD =aA ; which can be set to a suitable value by selecting appropriate conditions in the donor and acceptor phases. 18.2.2 Microporous membrane liquid liquid extraction (MMLLE) For cases where extraction is made from water into an organic solvent, but a further extraction into a second aqueous acceptor is not required, the technique of MMLLE replaces SLM. Here, the acceptor phase is an organic solvent and the same solvent forms the liquid membrane by ﬁlling the pores in the porous hydrophobic membrane [16]. Chemically, this is analogous to conventional liquid –liquid extraction, but performed in a ﬂow system, easily automated and interfaced to analytical instruments. LLE in a ﬂow system (in the form of FIA) has been described many times, as reviewed by Valca´rcel [17] and others, but the organic and aqueous phases are then mixed in the same ﬂow channel and later separated. The practical problems with the phase separation seem to have prevented this technique being widely used. In MMLLE, the phases are never mixed and all mass transfer between the phases take place at the membrane surface.

565

˚ ke Jo¨nsson and Lennart Mathiasson Jan A

In MMLLE, there is no trapping as in SLM, so the extraction efﬁciency is limited by the partition coefﬁcient. If it is very high, it is possible to work with a stagnant acceptor and still get a considerable enrichment into a very small extract volume. With smaller partition coefﬁcients, it might be necessary to arrange a slow ﬂow of the acceptor phase to remove the extracted analyte and maintain diffusion through the membrane.

18.3

CHEMICAL PRINCIPLES FOR METAL EXTRACTION

Addition of ion-pairing reagents or chelating reagents to the donor phase, permits the SLM extraction systems to be used for various metal ions. Various carrier molecules or ions can be incorporated in the membrane phase to enhance selectivity and mass transfer, as well as trapping reagents in the acceptor phase preventing analytes to be extracted back into the membrane. This was reviewed by Sastre et al. [4] and by Bufﬂe et al. [18]. As an analytical example of an addition of a reagent to the donor phase, we can mention extraction of metals from solutions containing a complex former as 8-hydroxyquinoline. This forms extractable complexes with many metals [11]. See Fig. 18.4a. The complex is transported through the membrane and the extracted analyte is trapped in the acceptor by another ligand, in the cited work, DTPA which forms a stronger and charged complex. The addition of a carrier to the membrane phase can be made in several ways. A common carrier that has been used both for extraction of metals and for organic acids is Aliquat-336 (methyltrioctylammonium chloride). It is a tertiary ammonium ion, i.e., permanently positively charged in ion pair with chloride and it can be added to a suitable membrane solvent. It has been used for the extraction of Cu, Cd, Co, Zn [11] and also for chromate [19]. With the addition of thiocyanate ions to the donor a negatively charged metal –thiocyanate complex is formed in the donor and is extracted as ion pair with the Aliquat-336 cation. The metal ion is eventually trapped in the acceptor using DTPA as described above and presented in Fig. 18.4b. A commonly used extractant for metal ions is DEHPA [20,21]. See Fig. 18.4c. In this case a pH gradient must exist over the membrane, so the acceptor is kept more acidic than the donor (typically pH < 1 and pH < 3, respectively). Speciation of different chromium species (chromate and chromium ion) has been performed by the combination of two membrane extraction systems, one working with DEHPA for extraction of Cr3þ and the other with Aliquat-336 for extraction of the chromate anions [19]. 18.4

PROPERTIES OF MEMBRANE EXTRACTION

In this section, the main advantages of membrane extraction in sample preparation are discussed. Compared to other techniques, membrane extraction

566

Membrane extraction

Fig. 18.4. Membrane extraction schemes of metal ions (Me). HQ ¼ 8-hydroxyquinoline, DTPA5- ¼ diethylenetriaminepentaacetic acid anion, R4Nþ ¼ methyltrioctylammonium cation, HD ¼ diethylhexyl phosphoric acid. For a, b, and c, see the text.

techniques provide advantages in various ways. Membrane extraction probably offers the highest degree of selectivity and cleanup of all known techniques for complicated matrices, and it is possible to achieve very high enrichment factors with preservation of the selectivity. Automation and on-line connection to instruments for ﬁnal analysis can be readily made. Compared to most other techniques for sample preparation, the use of organic solvent is much reduced, in most cases essentially to zero. This is true for a large number of compound classes, in various matrices and concentration ranges. Here, the main emphasis is on the extraction of metal ions. 18.4.1 Clean-up and selectivity All types of nonporous membrane extraction procedures will, in principle, lead to a high degree of cleanup, especially between small and large molecules. The analytes to be analyzed must dissolve into the membrane, pass through it and

567

˚ ke Jo¨nsson and Lennart Mathiasson Jan A

re-dissolve in the acceptor phase. In many cases, the conditions of extraction can be set so that this chain of events is possible only for a strictly limited range of compounds. The possibilities of achieving this tuning are best for the SLM technique, where selective reactions in all three phases can be utilized for this purpose. There are also a number of such possibilities for the other membrane extraction techniques. There are a number of other possible additives that have been used in SLM extraction for enhancing the extraction efﬁciency of different classes of compounds, including chelating or complexing reagents, crown ethers, ion pair formers, artiﬁcial receptors, etc. Some of these are mentioned in the applications section below. Both in biomedical analysis and in environmental analysis an important objective for the sample preparation is removal of high-molecular weight material. In the case of biomedical analysis, such material is usually proteins, and in environmental applications it is primarily humic substances. The membrane extraction techniques are all very efﬁcient in this respect; such high-molecular weight compounds are often charged and therefore, not extracted into organic liquids. Even if they are noncharged, the transport is so slow that their extraction is negligible. 18.4.2 Enrichment The different membrane extraction techniques behave differently when it comes to concentration enrichment factors. In aq/org types of extraction (MMLLE) the maximum concentration enrichment factor is limited to the value of the partition coefﬁcient between the donor and the acceptor phases. In such techniques, appreciable extraction factors are possible only when the partition coefﬁcient is large; the same situation as for ordinary LLE. This does not preclude that considerable enrichment factors can be obtained when the conditions are favorable. Extraction enrichment factors of about 250-fold were obtained in the MMLLE extraction of cationic surfactants from natural water [22]. In SLM, on the other hand, the enrichment factors are not limited by the partition coefﬁcient, but from the trapping conditions in the acceptor phase [15]. With some simpliﬁcations, the maximum enrichment factor, Ee ðmaxÞ; for SLM extraction is, in general, described by Eq. (18.4). Depending on the chemical principles and actual experimental conditions used, the actual values of aA and aD can be calculated and a value for Ee ðmaxÞ; can be estimated. For example, for the often used extraction system employing DEHPA, it is found for the extraction of a divalent metal ion that (unpublished): EeðmaxÞ ¼ 102ðpHD 2pHA Þ

ð18:5Þ

Thus, by assuring that the pH difference between the donor and acceptor is at, say, 3 units, the maximum enrichment factor is about one million, so for all

568

Membrane extraction

practical cases, it is possible to obtain high enrichment factors without approaching the limit set by Eq. (18.5). There are examples in the literature (extraction of aniline) where enrichment factors of more than 2000-fold are obtained after overnight extractions [15]. To really obtain high enrichment factors, a high sample/extract volume rate is necessary. This is evident from Eq. (18.3), showing that even if E approaches 1, the enrichment factor is never larger than the volume ratio. The strength of SLM in this context is that it can provide high extraction efﬁciencies at the same time as the extract volume is kept small. A prerequisite for high extraction efﬁciencies, in general, is that the extraction needs to be very selective, so only the analytes are enriched, not various interfering compounds. As noted above, high selectivity is a characteristic of the SLM technique. As was discussed above in Section 18.2.1.3, equilibrium extraction is sometimes attempted. Such conditions can easily be reached by setting the pH difference to 0.5 units, so the maximum enrichment factor (Eq. (18.5)), will be only 10 and the extraction process is acceptor-controlled. 18.4.3 Automation and unattended operation As the membrane extraction process can be most conveniently performed in ﬂow systems, it is easy to devise arrangements employing pumps, autosamplers, solenoid and rotary valves, etc., with computer control that can provide more or less automated operation. Below, a number of examples of such systems are discussed. For unattended operation of automated systems, the high selectivity and cleanup possible with membrane extraction is an advantage, as it prolongs the usable life-time of columns, etc. On the other hand, membrane stability can be a limiting factor. Sometimes, fouling by dirty samples can be seen, but it is possible to devise washing schemes in automated membrane extraction to diminish the problem. In any case, a smooth membrane surface is less amenable to fouling than an SPE column, for example, and it is easier to wash. For SLM extraction, the issue of membrane stability is often raised. As an organic solvent is held in the pores of a hydrophobic porous membrane placed between two aqueous ﬂowing streams, this demands that the solvent used is nonsoluble in water and that the capillary forces that hold the liquid in the pores are strong enough to withstand inevitable pressure differences over the membrane. Practically, these potential problems are not crucial. No serious problems with pressure differences have ever been seen with SLM set-ups, and simple calculations show that pressures of several bars are necessary to “blow” out the organic phase from the pores in typical cases. The solubility of nonpolar solvents in water is typically very small, and a solvent like n-undecane, which has been used extensively, forms membranes that are stable for months. Some problems may be encountered when more polar membranes are needed.

569

˚ ke Jo¨nsson and Lennart Mathiasson Jan A

A medium-polar solvent that has been used extensively is di-n-hexyl ether, whose stability in SLM membranes is nearly as good as n-undecane. The inclusion of various additives might compromise the stability and the matter calls for careful attention. Additives, which are as hydrophobic as possible (maybe modiﬁed with alkyl chains) are advantageous. For example, membranes containing the extractant DEHPA in kerosene are stable for at least a week of continuous operation [20]. The material of the liquid membrane support seems to somewhat inﬂuence the stability, the most commonly used PTFE membranes being slightly better than polypropylene membranes. The regeneration of the SLM is made in a few minutes by simply soaking the membrane support in the desired liquid, wiping and reinstalling the SLM in the membrane holder. In some cases, in situ regeneration has been shown to work well. 18.4.4 Solvent consumption Compared to alternative sample preparation techniques, membrane extraction demands very little solvent. This is a signiﬁcant advantage, as the cost for high purity solvent is high, both for purchasing and for disposal. Even more important, the environmental implications of these solvents are considerable, both for the laboratory workers and for the outer environment. This is especially true for chlorinated solvents, where different types of restrictions and bans have been proposed and already partly implemented in several countries. Only for MMLLE, small amounts of conventional organic solvents are needed, typically less than 1 ml of heptane, for example, is used for each sample. Thus, the membrane extraction techniques compare favorably with the alternative techniques in terms of solvent consumption.

18.5

EXPERIMENTAL SET-UP

18.5.1 Flow systems for membrane extraction Simple and inexpensive membrane extraction ﬂow systems for reasonably large sample volumes (100 ml and up) can be built up around a peristaltic pump and membrane units with channel volumes around 1 ml. An example of such a system is seen in Fig. 18.5. With this set-up, the acceptor phase is manually removed by the use of a syringe after each extraction. Such systems have been used both for laboratory work [11] and for sampling in natural waters [23] and in greenhouses [24]. Such systems can be completely or partially automated with pneumatically or electrically actuated valves controlled by timers or by computer systems [20,25].

570

Membrane extraction

Fig. 18.5. Manual off-line apparatus for liquid membrane extraction based on a peristaltic pump after Knutsson et al. [23].

For smaller samples (, 1 ml) the liquid delivery precision of peristaltic pumps is not adequate and membrane extraction equipment based on syringe pumps connected to the so-called robotic liquid handlers can be applied [26]. This principle does not seem to have been used yet for membrane extraction of metals.

18.5.2 How to set up a membrane extraction experiment for metal ions Extraction of metal ions from different types of environmental water matrices is relatively straightforward. There are several ways to extract metal ions using SLM. Here, one of the most versatile principles involving di-(2ethylhexyl) phosphoric acid (DEHPA; CAS 298-07-7), as shown in Fig. 18.4c, will be described. This works for many metal ions, including Cd, Co, Cr, Cu, Mn, Ni, Pb and Zn [19– 21,27]. The membrane contains DEHPA, which is insoluble in water. The acidic proton on this compound can be replaced by a metal ion, forming an extractable complex. This travels to the acceptor side, where protons force out the metal ion from the complex. The driving “force” is a proton gradient working from the acceptor to the donor, and it is necessary that the pH in the acceptor is kept sufﬁciently low during the entire extraction. The limiting factor for this extraction is the supply of protons in the acceptor—when these are “used up”, the extraction stops. Table 18.1 lists the equipment needed to set up a simple system for manual SLM. It should be mounted as shown in Fig. 18.5. Membrane units can be prepared in-house or obtained from the Lund University workshop via the authors of this paper. They might also be obtained from suppliers of FIA instrumentation. In some cases, the pump channel marked “Acid” is not needed, which removes the necessity for the mixing coil as well. If desired, a separate pump channel can be used for pumping the acceptor phase.

571

˚ ke Jo¨nsson and Lennart Mathiasson Jan A TABLE 18.1 18.1 Equipment needed for a manual SLM system for the extraction of metals (a) Membrane unit (1 ml channel volumes; see end of paper) (b) Peristaltic pump to provide a ﬂow of 0.5 –5 ml/min (c) Tubes for the pump, acid resistant if needed (d) PTFE tubing (1.6 mm) with ﬁttings to connect membrane unit to the pump (see Fig. 18.4a). (e) A plastic 10 ml syringe with screw ﬁtting

In this case, it is necessary that this channel can be stopped during the extraction. The analysis after extraction is made by AAS or ICP, typically in an off-line, discrete way, but direct connection has been demonstrated [21,25]. Ion-pair chromatography has also been described for this purpose [27]. The membrane is prepared in the following way: porous PTFE ﬁlters having a suitable diameter (typically 90 mm) and pore size 0.2 mm are soaked with the selected membrane liquid in a Petri dish for at least 15 min. The membrane typically contains, e.g., 40% DEHPA (in this example) in kerosene or in DHE. Suitable membrane ﬁlters are FG (Millipore, Bedford, MA, USA) or TE35 (Schelicher & Schuell, Dassel, Germany). Wipe away excess liquid with a tissue. Mount the membrane in the unit and tighten the screws evenly and ﬁrmly. Wash both acceptor and donor side with water (5 –10 ml) to remove the remaining organic liquid. The pH of the donor solution (i.e. sample) should be weakly acidic ðpH3 – 4Þ: If the pH of the sample needs adjustment, this can be done either manually by the addition of a suitable acid or by pumping acid with a separate pump channel, as shown in Fig. 18.5. A mixing coil is necessary in this case. The acceptor pH needs to be at least two units lower and have a sufﬁcient buffer capacity. Prepare a proper solution (typically 1 M mineral acid) and ﬁll the acceptor channel of the membrane unit using a syringe or a separate pump channel, as mentioned above. Adjust the pump to a suitable ﬂow rate, as a ﬁrst suggestion, to 1– 2 ml/min. Higher ﬂow rates (up to 7 ml/min) will lead to faster accumulation of analyte in the acceptor and thus to a faster extraction, but might lead to problems with leakage. Collect the outﬂow from the donor channel in a measuring cylinder to measure the ﬂow rate and the total extracted sample volume. For the ﬁrst extractions, process about 50 ml of pure water spiked with different concentrations. After each extraction, wait a short time (see below) and collect the acceptor in a 2 ml measuring ﬂask or graduated test tube. Rinse both the channels by passing at least 10 ml of water and acceptor buffer, respectively, before the next sample is extracted. Analyze an aliquot of the acceptor and determine the amount of analyte that has been extracted.

572

Membrane extraction

Calculate the extraction efﬁciency according to Eq. (18.1), considering the actual dilution factors to refer concentrations and amounts to the conditions in the membrane channels. With the conditions stated, E should be 50–70%. It is possible to increase it by decreasing the sample ﬂow rate, but that will lead to smaller concentrations in the acceptor than with a higher ﬂow rate, other conditions (e.g. extraction time) being the same. To optimize the extraction, the main parameter to be changed is the composition of the membrane liquid. The inﬂuence of the pH of the phases is well known, and if they are selected as above they will be optimal, the same is the case with the ﬂow rates. To increase the extracted amount, and thus the sensitivity of the analysis, larger sample volumes can be extracted, perhaps with a higher ﬂow rate. It is necessary to verify that the extraction efﬁciency is reasonably constant over the volume range up to the actual sample volume; if it decreases with sample volume (keeping ﬂow rate constant) the trapping is not efﬁcient, and a more acidic pH in the acceptor is needed. It can also be the case that the buffer capacity of the acceptor is not sufﬁcient. It is wise to check the pH of the acceptor phase after each extraction with a pH paper. If the pH is completely wrong, it is possible that there is a hole in the membrane leading to mixing of the phases, with detrimental results. It should be possible to extract liters of water (which, of course, will take some hours) to reach a concentration enrichment of several hundred times (with 1000 ml sample, acceptor volume of 1 ml and E ¼ 50%; Ee ¼ 500 according to Eq. (18.3)). Memory effects need to be considered. After the extraction is ﬁnished (i.e., when the pumping of the sample is stopped) there is a portion of analyte still remaining in the membrane, and this must have time to reach the acceptor in order to be recovered for analysis. A waiting time of 10 –15 min will allow most of this material to diffuse to the acceptor. This can be checked by harvesting several portions of acceptor after each other following the extraction of one sample. Still, there will be some analyte left in the membrane and in the apparatus, and it is necessary to wash both the channels thoroughly before the next extraction to avoid carry-over effects. It should be demonstrated that the carry-over is negligible by extracting blanks after extracting samples containing high analyte concentrations. The measurement of memory effects has been discussed in several papers [11,28,29]. There should be no major matrix problems running samples like surface water, waste water, urine, etc., with this set-up. It should provide a high degree of cleanup from humic compounds and other interfering materials, even at high enrichment factors. The lifetime of the membrane should be several weeks before it needs replacement. To further develop the extraction set-up, it is possible to automate it by connecting pneumatic or electrical valves and pass the acceptor directly to the AAS. For ideas on how this can be done, see Refs. [20,25].

573

˚ ke Jo¨nsson and Lennart Mathiasson Jan A

ACKNOWLEDGEMENTS The work of the Lund membrane extraction group was, over the years, ﬁnancially supported by grants from the following authorities: NFR, VR, SNV, SJFR, SIDA, the Swedish Institute, the Crafoord Foundation and the European Community (DG XII). A number of graduate and undergraduate students as well as guest researchers have made important contributions, which are highly appreciated. REFERENCES 1 2

3 4 5 6 7

8 9

10 11 12 13 14 15 16 17 18

19 20 21

574

˚ . Jo¨nsson and L. Mathiasson, J. Separation Sci., 24 (2001) 495 –507. J.A ˚ . Jo¨nsson, Liquid membrane extraction, In: J. Pawliszyn (Ed.), Sampling and J.A sample preparation for ﬁeld and laboratory. Elsevier Science, Amsterdam, 2002, pp. 503– 530. R.A. Bartsch and J.D. Way, Chemical Separations with Liquid Membranes, ACS Symposium Series 642, ACS, Washington, DC, 1996. A.M. Sastre, A. Kumar, J.P. Shukla and R.K. Singh, Sep. Puriﬁc. Meth., 27(2) (1998) 213– 298. G.A. Audunsson, Anal. Chem., 58 (1986) 2714–2723. ˚ . Jo¨nsson and L. Mathiasson, Trends Anal. Chem., 11 (1992) 106– 114. J.A H. Lingeman, In: E. Reid, H.M. Hill and I.D. Wilson (Eds.), Drug development assay approaches including molecular imprinting and biomarkers. The Royal Society of Chemistry, Cambridge, 1998. N.C. van de Merbel, J. Chromatogr. A, 856 (1999) 55– 82. B. Moreno Cordero, J.L. Pere´z Pavo´n, C. Garcı´a Pinto, E. Ferna´ndez Laespada, R. Carabias Martı´nez and E. Rodrı´guez Gonzalo, J. Chromatogr. A, 902 (2000) 195– 204. ˚ . Jo¨nsson and L. Mathiasson, J. Chromatogr. A, 902 (2000) 205– 225. J.A ˚ . Jo¨nsson and L. Mathiasson, Analyst, M. Papantoni, N.-K. Djane, K. Ndung’u, J.A 120 (1995) 1471– 1477. J.-F. Liu, J.-B. Chao and G.-B. Jiang, Anal. Chim. Acta, 455 (2002) 93–101. ˚ . Jo¨nsson, P. Lo¨vkvist, G. Audunsson and G. Nilve´, Anal. Chim. Acta, 227 J.A (1993) 9 –24. ˚ . Jo¨nsson, Anal. Chim. Acta, 416 (2000) 77–86. L. Chimuka, L. Mathiasson and J.A ˚ . Jo¨nsson, Anal. Chem., L. Chimuka, N. Megersa, J. Norberg, L. Mathiasson and J.A 70 (1998) 3906– 3911. ˚ . Jo¨nsson and L. Mathiasson, Anal. Chem., 70(5) (1998) 946– 953. Y. Shen, J.A M. Valca´rcel and M.D. Luque de Castro, Non-Chromatographic Continuous Separation Techniques. The Royal Society of Chemistry, Cambridge, 1991. J. Bufﬂe, N. Parthasarathy, N.-K. Djane and L. Mathiasson, In: J. Bufﬂe and G. Horvai (Eds.), In Situ Monitoring of Aquatic Systems: Chemical Analysis and Speciation. Wiley, Chichester, 2000, pp. 407– 493. N.-K. Djane, K. Ndung’u, C. Johnson, H. Sartz, T. To¨rnstro¨m and L. Mathiasson, Talanta, 48(5) (1999) 1121–1132. N.-K. Djane, K. Ndung’u, F. Malcus, G. Johansson and L. Mathiasson, Fresenius J. Anal. Chem., 358 (1997) 822–827. N.-K. Djane, I.A. Bergdahl, K. Ndung’u, A. Schu¨ tz, G. Johansson and L. Mathiasson, Analyst, 122 (1997) 1073–1077.

Membrane extraction 22 23 24 25 26 27 28 29

˚ . Jo¨nsson, J. Chromatogr. A, J. Norberg, E. Thordarson, L. Mathiasson and J.A 869(1/2) (2000) 523–529. ˚ . Jo¨nsson, J. Agric. Food Chem., 40 M. Knutsson, G. Nilve´, L. Mathiasson and J.A (1992) 2413– 2417. ˚ . Jo¨nsson, N. Niedack, P. Bowens and B. Alsanius, Anal. V. Jung, L. Chimuka, J.A Chim. Acta, 474 (2002) 49–57. F. Malcus, N.-K. Djane, L. Mathiasson and G. Johansson, Anal. Chim. Acta, 327 (1996) 295–300. ˚ . Jo¨nsson, L. Mathiasson and A.-M. Olsson, Anal. B. Lindega˚rd, H. Bjo¨rk, J.A Chem., 66 (1994) 4490– 4497. K. Ndung’u, N.-K. Djane and L. Mathiasson, J. Chromatogr. A, 826 (1998) 103– 108. ˚ . Jo¨nsson, Anal. Chim. Acta, 292 (1994) 31–39. Y. Shen, L. Gro¨nberg and J.A ˚ . Jo¨nsson and L. Mathiasson, Int. J. Environ. Anal. T. Miliotis, M. Knutsson, J.A Chem., 64 (1996) 35– 45.

575

This page intentionally left blank

Chapter 19

Derivatization and vapor generation methods for trace element analysis and speciation Yong Cai

19.1

INTRODUCTION

It is generally recognized that the determination of the total amount of an element is important, but not sufﬁcient to assess its toxicity and bioavailability. Information about concentrations of the individual species of an element, including its organic derivatives, is particularly crucial. Frequently, the lack of speciation information is the major limitation to our understanding of the biogeochemical cycling of the elements [1 –6]. Speciation analysis is usually deﬁned as the determination of the concentrations of the individual physicochemical forms of the element in a sample that together constitute its total concentration. Speciation analysis has become one of the fastest developing areas of analytical chemistry over the last two decades. Techniques for the determination of the total amount of an element have been well developed and documented [7,8]. The identiﬁcation of the different chemical forms of an element, however, has been a challenging research area in environmental and biomedical studies. Two complementary techniques are necessary for trace element speciation; one provides an efﬁcient and reliable separation, and the other provides adequate detection and quantitation [9,10]. In its various analytical manifestations, chromatography is a powerful tool for separation of a vast variety of chemical species of trace elements. As for the detection systems, atomic spectroscopy offers the possibility of selectively detecting a wide range of metals and non-metals. The use of detectors responsive only to selected elements in a multi-component mixture drastically reduces the constraints placed on the separation step, as only those components in the mixture that contain the element of interest will be detected [6]. It is not surprising that the coupling of chromatographic techniques [gas chromatography (GC) and high performance liquid chromatography (HPLC)] with a highly sensitive and selective atomic spectrometry detector has been widely Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

577

Y. Cai

exploited and accepted for the speciation of metals and organometallic compounds. GC has enjoyed particular attention because of its high sensitivity and simplicity of coupling. The native species of most metals and metalloids, such as mercury, lead, tin, arsenic, and selenium, are generally present as ionic or polar forms in sample matrices. Direct analysis of these species using chromatography-coupled methods is, therefore, limited. For GC-based coupling techniques, these compounds need to be extracted from the sample matrix and converted to volatile and thermally stable derivatives. A number of derivatization and vapor generation methods have so far been developed. Derivatization reactions, method development, and applications for some of the important approaches are discussed in this chapter.

19.2

THEORY

19.2.1 Grignard reactions Grignard reagents (RMgX), a series of magnesium-containing compounds, are probably the most important for both organic and inorganic syntheses [11]. These organomagnesium halides were discovered by the French chemist Victor Grignard in 1900. Grignard received the Nobel Prize for his discovery in 1912, and organomagnesium halides are called Grignard reagents in his honor [12]. The reaction of metal and organometallic halides with Grignard reagents provides a very useful synthetic route to fully alkylated organometallic compounds [13]. The Grignard reaction is one of the most widely used derivatization techniques for speciation of a number of elements [14 –16]. Examples for organotin and organomercury analyses are illustrated in Eqs. (19.1) and (19.2): Rn Snð42nÞþ þ R0 MgX ! Rn SnR0ð42nÞ

ð19:1Þ

where n ¼ 0; 1, 2, 3; R ¼ methyl, butyl, etc.; R0 ¼ methyl, ethyl, propyl, butyl, pentyl, etc.; X ¼ Cl2, Br2, I2. Rn Hgð22nÞþ þ R0 MgX ! Rn HgR0ð22nÞ

ð19:2Þ

where n ¼ 0; 1; R ¼ methyl, ethyl; R0 ¼ methyl, ethyl, propyl, butyl, pentyl, phenyl, etc.; X ¼ Cl2, Br2, I2. The main advantages of this reaction are that different alkyl groups can be chosen to make fully alkylated species and this method generally gives good yields. However, Grignard reagents are very strong bases, they react with any compound that has a hydrogen attached to an electronegative atom such as oxygen, nitrogen, or sulfur. Therefore, Grignard reagents are very sensitive to water. The reactions of Grignard reagents with water are nothing more than acid –base reactions; they lead to the formation of the weaker conjugate acid

578

Derivatization and vapor generation methods for trace element analysis and speciation

and weaker conjugate base [12]. As a consequence, metal and organometallic compounds in aqueous samples have to be extracted prior to derivatization into an organic solvent with the assistance of complexing reagents such as dithiocarbamates and tropolone. The sample preparation process can be tedious and time consuming. 19.2.2 Hydride generation Reduction of non-volatile compounds to volatile hydrides is one of the most widely used derivatization reactions for analysis and speciation of metals and metalloids. Although various reducing reagents have been used as a means of producing hydride, sodium borohydride (NaBH4) is currently used almost exclusively. Several elements (As, Bi, Cd, Ge, Pb, Sb, Se, Sn, and Te) are known to form volatile hydrides [17– 19]. When these metals/metalloids are present in inorganic forms, NaBH4 converts them into their hydrides (e.g. AsH3, SeH2, SbH3) except for Hg, where volatile Hg0 is formed. Hydride generation is also well established for analysis of some organometallic compounds. The general reactions for alkylated As and Sn compounds can be described as: Men AsOðOHÞ32n þ NaBH4 ! Men AsHð32nÞ

ð19:3Þ

where n ¼ 0; 1, 2. Rn Snð42nÞþ þ NaBH4 ! Rn SnHð42nÞ

ð19:4Þ

where n ¼ 0; 1, 2, 3; R ¼ methyl, butyl, etc. The pH of the reaction medium is important for the derivatization. In general, the reduction reactions are performed at a pH that is a few units below pKa of the species of interest [20]. It should be pointed out that NaBH4 has also been reported to convert methylmercury (MeHg) to MeHgH [21,22]. MeHgþ þ NaBH4 ! MeHgH

ð19:5Þ

This derivatization reaction is carried out at pH 2, while under alkaline condition MeHg will produce Hg0. Because MeHgH is quite unstable, it has never been purely separated. Its presence has been demonstrated by several groups using various techniques [21,22]. The greatest advantage of this method is its simplicity and high sensitivity. Volatile hydrides can be purged from the reaction solution using an inert gas, such as helium, trapped in a column and then analyzed using an elementspeciﬁc detector (detailed information on the cryogenic trapping technique can be found in Chapter 16 by Donard in this volume). The usefulness of this procedure for speciation analysis, however, is restricted by either a thermodynamic inability for hydride formation by some species, or a considerable kinetic limitation to hydride formation. Although little interferences were usually observed in the detection stage, due to the separation of volatile hydride from the matrix, the latter frequently causes interferences in the

579

Y. Cai

reduction reactions in the reaction vessel [19,23]. Side-reactions may also lead to analytical problems [19]. 19.2.3 Aqueous derivatization with tetraalkyl(aryl)borates Some limitations of the Grignard reaction and hydride generation can, to a certain degree, be overcome by replacing NaBH4 with alkyl(aryl)borates [1,16,24,25]. These derivatization reactions can be performed in aqueous media, subsequently reducing analysis time and eliminating organic solvent extraction required with Grignard reactions. They are also versatile, in terms of selecting alkyl(aryl) groups. The following aqueous derivatizations have been reported for analysis and speciation of metals/metalloids and their organic derivatives. 19.2.3.1 Ethylation The use of sodium tetraethylborate (NaBH4) as a derivatization reagent for speciation of organometallic compounds was initiated for the determination of ionic methyllead compounds [26]. NaBEt4 acts as an aqueous phase ethylation reagent, transferring Et2 ions to ionic metals and organometallic compounds and forming thermally stable products. These derivatives can be then analyzed by using purge-and-trap techniques, if they are sufﬁciently volatile, or extracted into organic solvents and analyzed with GC. Aqueous phase ethylation with NaBEt4 has been used for the speciation of a variety of metals, including lead, mercury, and tin in different environmental and biological samples. The theory and applications of this method have been thoroughly reviewed [24]. The representative reactions for Pb and Hg can be described as: Men Pbð42nÞþ þ NaBEt4 ! Men PbEtð42nÞ

ð19:6Þ

where n ¼ 0; 1, 2, 3. Men Hgð22nÞþ þ NaBEt4 ! Men HgEtð22nÞ

ð19:7Þ

where n ¼ 0; 1. Although this derivatization reaction is currently the most widely used method for methylmercury speciation, it suffers from a few major deﬁciencies that limit its application [16]. First, the reagent is air-sensitive and consequently requires careful handling during preparation of the solution. Because of this, in part, the cost of using NaBEt4 can be substantial. Second, this method cannot distinguish between monoethylmercury and inorganic mercury (Hg2þ) species, as evident from reaction (19.7), because both monoethylmercury and Hg2þ are converted to Et2Hg. Monoethylmercury has not generally been found in marine and freshwater animals [16], but its occurrence has been reported in soil and sediment, both polluted [27] and

580

Derivatization and vapor generation methods for trace element analysis and speciation

natural [15]. Similarly, the speciation of the native ethyllead species cannot be performed using NaBEt4, unless a deuterated reagent is speciﬁcally used [78]. 19.2.3.2 Propylation The versatility of sodium tetrapropylborate (NaBPr4) as an aqueous derivatization reagent for speciation of organometallic compounds was recently demonstrated [28]. As expected, NaBPr4 behaves similarly to NaBEt4, and can readily react with a number of organometallic compounds. In contrast to NaBEt4, it is possible to analyze the important ethylated species of mercury and lead with the use of NaBPr4 as shown in the following reactions: Rn Pbð42nÞþ þ NaBPr4 ! Rn PbPrð42nÞ

ð19:8Þ

where n ¼ 0; 1, 2, 3; R ¼ methyl, ethyl, etc. Rn Hgð22nÞþ þ NaBPr4 ! Rn HgPrð22nÞ

ð19:9Þ

where n ¼ 0; 1; R ¼ methyl or ethyl. However, NaBPr4 is also fairly air-sensitive. More research efforts are needed to evaluate its routine applications for organometallic speciation, especially for mercury and lead. 19.2.3.3 Phenylation Tetraphenylborate (NaBPh4) has also been evaluated for analysis of organometallic species [25,29 – 31]. Although this reagent reacts readily with inorganic mercury and alkylmercury, it does not derivatize organolead and organotin compounds [30]. Rn Hgð22nÞþ þ NaBPh4 ! Rn HgPhð22nÞ

ð19:10Þ

where n ¼ 0; 1; R ¼ methyl or ethyl. An analytical procedure has recently been described for the determination of methylmercury and ethylmercury compounds in ﬁsh and sediment samples using GC and atomic ﬂuorescence spectrometry (AFS) following aqueous phenylation with NaBPh4 [25]. The derivatization products were identiﬁed by GC –mass spectrometry (GC–MS). Several points should be mentioned when comparing phenylation to ethylation. The pKa values of NaBEt4 and NaBPh4 are 50 and 44, respectively, indicating that NaBEt4 is a stronger base and is, therefore, more reactive to water [16,25]. Indeed, while NaBPh4 can be prepared using regular laboratory procedures, NaBEt4 has to be prepared with care. It is frequently handled under an argon atmosphere to avoid contact with air. Aqueous solutions of NaBPh4 are much more stable than those of NaBEt4. This makes use of NaBPh4 much more cost-effective. Additionally, the thermal stability of ethylated and phenylated derivatives shows a different pattern. Theoretically, the RHg –C6H5 bond is stronger than that of the RHg –C2H5 bond, meaning the derivatives of NaBPh4 are more thermally stable [16,25].

581

582 Y. Cai Fig. 19.1. Flow diagram showing representative procedures for the determination of organotin compounds in environmental and biological samples using Grignard reactions. More details are reported in Ref. [33]. Abbreviations: NaDDC, sodium diethyldithiocarbamate; Pe3SnCl, tripentyltin chloride; PrMgBr, propylmagnesium bromide; TMAH, tetramethylammonium hydroxide.

Derivatization and vapor generation methods for trace element analysis and speciation

Finally, NaBPh4 can be used for both methyl- and ethylmercury analysis, while NaBEt4 cannot.

19.3

METHOD DEVELOPMENT

Procedures for derivation and experimental conditions for trace element speciation and analysis have been reported for a variety of applications. It is not the intention of this chapter to extensively review these methods, rather, representative derivatization schemes for the Grignard reactions and the aqueous derivatizations (hydride generation and alkyl(aryl)ation) are presented. Note that the general steps involved in the derivatization for different applications are usually similar. Detailed information for some speciﬁc applications can be found in the original publications summarized in Section 19.4. 19.3.1 Grignard reactions Analytical procedures utilized for the determination of trace elements and their organic derivatives using Grignard reactions have been discussed and reviewed [1,15,16,32,33]. A schematic diagram illustrating the representative procedures used for organotin speciation in environmental and biological samples is shown in Fig. 19.1 [33]. Similar procedures can be applied to the analysis of other organometallic compounds, such as mercury and lead. In order to extract the ionic organotin compounds from water, sediment/soil, and biological samples into an organic solvent, complexing reagents are needed. The most commonly used reagents are sodium diethyldithiocarbamate (NaDDC) and tropolone. Use of an internal standard is critical for quantitative analysis and should be added at the beginning of the extraction step. After the reaction, the excess Grignard reagent needs to be destroyed for subsequent GC analysis. This is normally done by using dilute acid, such as 1 M H2SO4. Because of the production of heat after adding acid to the mixture, it is strongly recommended that this process be carried out slowly by adding acid drop by drop and/or by cooling the reaction system with ice. If sufﬁcient acid is added, aqueous and organic phases will be clearly separated. For extraction of environmental and biological samples, especially sediment/soil and biological tissues, the organic phase after derivatization usually contains colored substances that interfere with the subsequent GC analysis. These substances can be removed by a clean-up procedure using a Pasteur pipette as a microcolumn (e.g. 15 cm £ 5 mm i.d.) containing ca. 1 g of silica gel (silica gel 60, heated 16 h at 1708C, 5% water deactivated). Organometallic compounds present in the clean organic phase can be analyzed using different detection methods, such as GC –MS, GC –ﬂame ionization detection (GC –FID), and GC –atomic absorption spectrometry (GC –AAS).

583

Y. Cai

19.3.2 Aqueous derivatization A great number of extraction procedures have been developed for the determination of metals/metalloids and organometallic compounds using hydride generation and aqueous derivatization [1,3,5,16– 18,20,33–36]. Figure 19.2 presents a representative ﬂow diagram for the determination of organotin compounds in sediments using a cryogenic trapping technique following hydride generation or ethylation [34,35]. A similar procedure can be used for methylmercury analysis of ﬁsh tissue (using methanolic KOH for digestion instead of methanolic HCl) [37]. In contrast to the Grignard reactions, ionic compounds of interest do not need to be extracted into an organic solvent. These compounds are released from the matrices simply under acidic or basic conditions. The extraction conditions for releasing target compounds should be optimized for different matrices [36,38]. An aliquot of the extract can be analyzed using the sensitive cryogenic trapping technique. Several important parameters, such as reaction time, purge time, and the amount of NaBEt4 added to the reaction vessel, can signiﬁcantly affect the ﬁnal results. Commonly, in hydride generation, cryogenic trapping and GC –AAS experiments, the reaction and purging are performed simultaneously and completed within about 4.5 min [39]. The reaction conditions are modiﬁed for aqueous phase ethylation using NaBEt4 by adding a single reaction step without helium ﬂow through the reactor to ensure efﬁcient ethylation of organotins. Thereafter, ethylation products are purged from the reactor with helium. This extra step is designed to compensate for the ethylation reaction rate being slower than that for hydride generation [35]. A maximum purging time (usually less than 10 min) is set to avoid blockage of the cold trap due to condensation of water vapor. The optimization of the derivatization parameters is carried out using a fast simplex algorithm [26,35]. Instead of using cryogenic trapping, the ethylation products of mercury (Hg2þ and methylmercury) can be purged onto a Carbotrapw or Tenaxw column at room temperature. The derivatives are then desorbed onto an isothermal GC column and separated, and then analyzed with an atomic ﬂuorescence spectrometer (AFS) [40]. A procedure for methyl- and ethylmercury analysis in ﬁsh and sediment samples using phenylation with NaBPh4 is illustrated in Fig. 19.3 [25]. Phenylation can be accomplished in both aqueous and organic (CH2Cl2) phases. The derivatives are extracted into an organic phase and analyzed using different techniques. In contrast to the cryogenic technique, where all derivatives produced in an aliquot of sample are transferred to the detection system, only a small fraction of the organic phase is injected into the GC with this phenylation method. The detection limit of this method is therefore not low enough to be of use for natural water analysis. Nevertheless, it provides a reliable and sensitive technique for mercury analysis in ﬁsh and mercurypolluted sediment samples.

584

Derivatization and vapor generation methods for trace element analysis and speciation

Fig. 19.2. Flow diagram showing representative procedures for the determination of butyltin compounds in sediment samples using hydride generation and aqueous ethylation with NaBEt4 following extraction with methanolic HCl. More details are reported in Refs. [34,35]. Abbreviations: NaDDC, sodium diethyldithiocarbamate; Pe3SnCl, tripentyltin chloride; PrMgBr, propylmagnesium bromide.

19.4

APPLICATIONS

A great number of articles have been published on the determination of metals/metalloids and organometallic compounds using derivatization and vapor generation. Some book chapters and review articles have discussed the overall analytical techniques for the analysis and speciation of trace elements [1 –7]. Special efforts are made, in this chapter, to include some important studies in which method development, technical improvement, and signiﬁcant applications were addressed. These publications are categorized based on the derivatization techniques used. Applications of using

585

Y. Cai

Fig. 19.3. Flow diagram showing procedures for the determination of methylmercury in sediment and ﬁsh samples using aqueous phenylation with sodium tetraphenylborate. More details are reported in Ref. [25].

derivatization with Grignard reagents, hydride generation, and aqueous derivatization with alkyl(aryl)borate are summarized in Tables 19.1, 19.2, and 19.3, respectively. In conclusion, derivatization procedures are very important for the analysis and speciation of trace elements. In many cases, such pretreatment steps must be done in order to analyze the compounds of interest and their different chemical forms. These derivatization methods often separate trace elements from their matrices and therefore simultaneously reduce or eliminate interferences. Target compounds are frequently concentrated after the derivatization steps, subsequently improving the detection limit of the analytical technique. This is especially true for the cryogenic trapping technique. Aqueous derivatization with alkyl(aryl)borates overcomes the limitation of the Grignard reaction in which the compounds of interest must be transferred to an organic layer for derivatization. However, the use of the alkyl- and arylation reagents is limited by their commercial availability.

586

Derivatization with Grignard reactions for analysis and speciation of metals/metalloids and organometallic compounds Metal species

Sample matrix

Derivatization reagent

Detection method

Reference

TMT, DMT, MMT TBT, TPT, TCT, FBTO TeBT, TBT, DBT, MBT TBT, DBT, TPT TBT, DBT, MBT, TCyT, DCyT TPT, DPT, MPT TBT, DBT, MBT, Sn4þ TBT, DBT, MBT TeBT, TBT, DBT, MBT TMT, DMT, Sn4þ TML, DML, TEL, DEL TML, DML, TEL, DEL, Me2EtL, MeEt2L TML, TEL, DEL, Me2EtL, MeEt2L TML, DML, TEL, DEL, Pb2þ TML, DML, TEL, DEL, Me2EtL, MeEt2L MeHg, EtHg, Hg2þ MeHg, Hg2þ MeHg, Hg2þ

DI and lake waters Water, sediment Sediment, biological tissue Marine biological products Seawater, biological samples Biological tissue Sewage, sludge Sediment, ﬁsh Marine sediment Human organ samples Greenland snow sample Rainwater Snail soft tissue Sediment, biological sample Water Sediment Natural gas condensate Sediment

BuMgCl PeMgBr, MeMgI HexMgBr EtMgBr MeMgBr BuMgBr EtMgBr EtMgBr HexMgBr PeMgBr, PrMgBr PrMgBr PrMgBr BuMgCl BuMgBr BuMgBr BuMgBr BuMgBr BuMgCl

GC– AAS GC– MS, GC– AED GC– MS GC– FPD GC– MS GC– FPD GC– AAS GC– AED GC– MS GC– FPD, GC–ICP-MS GC– AED GC– AAS GC– AAS GC– AAS GC– AAS GC– AFS GC– AED GC– AED

[41] [42] [43] [44] [45] [46] [47] [48] [49] [66] [50] [51] [52] [53] [54] [15] [55] [56]

587

Abbreviations: TMT, trimethyltin; DMT, dimethyltin; MMT, monomethyltin; TBT, tributyltin; TPT, triphenyltin; TCT, tricyclohexyltin; FBTO, fenbutatin; TeBT, tetrabutyltin; DBT, dibutyltin; MBT, monobutyltin; DCT, dicyclohexyltin; TML, trimethyllead, DML, dimethyllead; TEL, triethyllead; DEL, diethyllead; Me2EtL, dimethylethyllead; MeEt2L, methyldiethyllead; GC – AAS, gas chromatography–atomic absorption spectrometry; GC– MS, gas chromatography–mass spectrometry; GC–AED, gas chromatography– atomic emission detection; GC– FPD, gas chromatography–ﬂame photometric detection; GC –AFS, gas chromatography – atomic ﬂuorescence spectrometry; GC– ICP-MS, gas chromatography– inductively coupled plasma-mass spectrometry.

Derivatization and vapor generation methods for trace element analysis and speciation

TABLE 19.1

588

TABLE 19.2 Hydride generation with sodium borohydride for analysis and speciation of metals/metalloids and organometallic compounds Sample matrix

Detection method

Reference

Se(VI), Se(IV) As(III), As(V), MMA, DMA, TMAO As, Se, Sn, Sb As, Sb Many metals and organometallic compounds MeHg MeHg MeHg TBT, DBT, MBT TBT, DBT, MBT TMT, DMT, MMT TBT, DBT, MBT, TMT, DMT, MMT TBT, DBT, MBT, TMT, DMT, MMT, MPT TBT, DBT, MBT TBT, DBT, MBT, TMT, DMT, MMT TBT, DBT

Lake water, sediment, aquatic invertebrates Natural waters Water samples Standard DI water (studied interferences) Tuna ﬁsh mussel (SRM, T-22l) Standard Sediment Sediment Sediment Simulated estuarine samples Water samples Natural waters, sediments, macro algae Estuarine sediments Oysters Sediments, microbial bioﬁlms

HG– AFS HG– CT– AAS HG– CT– AAS HG– AAS HG– LT-GC–ICP-MS HG– CT– FTIR HG– HSS–GC –AAS HG– AFS HG– AAS HG– CT– AAS HG– CT– AAS HG– CT– AAS HG– CT– AAS HG– CT– AAS HG– CT– AAS HG– GC–FPD

[23] [57] [58] [67] [19] [22] [21] [59] [39] [34] [60] [61] [62] [63] [64] [65]

Abbreviations: MMA, monomethylarsonic acid; DMA, dimethylarsinic acid; TMAO, trimethylarsine oxide; MeHg, monomethylmercury; TBT, tributyltin; DBT, dibutyltin; MBT, monobutyltin; TMT, trimethyltin; DMT, dimethyltin; MMT, monomethyltin; TPT, triphenyltin; HG–AFS, hydride generation –atomic ﬂuorescence spectrometry; HG–LTGC –ICP-MS, hydride generation– low temperature-gas chromatography –inductively induced plasma-mass spectrometry; HG–CT–AAS, hydride generation –cryogenic trapping –atomic absorption spectrometry; HG–CT–FTIR, hydride generation –cryogenic trapping –Fourier transform infrared spectrometry; HG–HSS –GC –AAS, hydride generation–head space sampling –gas chromatography –atomic absorption spectrometry; HG– GC –FPD, hydride generation–gas chromatography –ﬂame photometric detection.

Y. Cai

Metal species

Derivatization and vapor generation methods for trace element analysis and speciation TABLE 19.3 Derivatization with sodium tetraalkyl(aryl)borate for analysis and speciation of metals/metalloids and organometallic compounds Metal species

Sample matrix

Derivatization reagent

Detection method

Reference(s)

TML, DML, Pb2þ TML, DML, TEL, DEL, Pb2þ Pb2þ TBT, DBT, MBT TBT, DBT, MBT TBT, DBT, MBT

Standard Standard

NaBEt4 NaBEt4

[26] [68]

Water Water Sediments Sediments River water samples Sediments

NaBEt4 NaBEt4 NaBEt4 NaBEt4 NaBEt4

CT–AAS HPLC – ethylation –AAS GC –MS SPME–GC –MS CT–AAS GC –AAS, GC –MS GC –AED

[69] [70] [35,38] [71] [72]

NaBEt4

GC –FPD

[73]

Waters Standards Fish tissue Standards Natural waters, ﬁsh Standard, natural waters Fish, sediments Standards, biological tissues Waters Standards, marine sediments

NaBEt4 NaBEt4 NaBEt4 NaBEt4 NaBEt4

AFS H NMR, CT–AAS CT–AAS RTP-AFS RTP-CryGC –AFS

[74] [75] [37] [40] [76]

NaBEt4

ICP-IDMS

[77]

NaBPh4 NaBPh4

GC –MS, GC – AFS GC –AED

[25] [30]

NaBPh4 NaBPr4

GC –MS GC –ICP-MS

[31] [28]

TBT, DBT, MBT, TPT, DPT, MPT Cd2þ MeAsBr2, Me2AsBr MeHg, Hg2þ MeHg, Hg2þ MeHg, Hg2þ MeHg 2þ

MeHg, Hg MeHg, Hg2þ MeHg Hg, Pb, Sn, inorganic and organic species

1

Abbreviations: TML, trimethyllead, DML, dimethyllead; TEL, triethyllead; DEL, diethyllead; TBT, tributyltin; DBT, dibutyltin; MBT, monobutyltin; TPT, triphenyltin; DPT, diphenyltin; MPT, monophenyltin; CT–AAS, cryogenic trapping – atomic absorption spectrometry; HPLC – ethylation – AAS, high performance liquid chromatography –ethylation –cryogenic trapping –atomic absorption spectrometry; SPME –GC –MS, solid phase microextraction –gas chromatography –mass spectrometry, GC –AED, gas chromatography –atomic emission detection; GC –FPD, gas chromatography –ﬂame photometric detection; AFS, atomic ﬂuorescence spectrometry; RTP-AFS, room temperature preconcentration-atomic ﬂuorescence spectrometry; RTP-CryGC –AFS, room temperature preconcentration-cryogenic gas chromatography –atomic ﬂuorescence spectrometry, ICP-IDMS, inductively coupled plasma-isotope dilution mass spectrometry.

589

Y. Cai

Acknowledgement Thanks to Anita Holloway for her assistance with the preparation of this chapter. This is contribution 204 of the Southeast Environmental Research Center at FIU. REFERENCES 1 2 3 4 5 6 7 8 9 10

11 12 13

14 15 16 17 18 19 20 21 22 23 24 25 26 27

590

R. Lobinski and F.C. Adams, Spectrochim. Acta B, 52 (1997) 1865. K. Sutton, R.M.C. Sutton and J.A. Caruso, J. Chromatogr. A, 789 (1997) 85. Y. Cai, Trends Anal. Chem., 19 (2000) 62. S. Caroli (Ed.), Element Speciation in Bioinorganic Chemistry. Wiley, New York, 1996. Y. Cai, in: J. Cazes (Ed.), Encyclopedia of Chromatography. Marcel Dekker, New York, 2001, p. 518. R. Lobinski and J. Szpunar, Anal. Chim. Acta, 400 (1999) 321. C. Vandecasteele and C.B. Block (Eds.), Modern Methods for Trace Element Determination. Wiley, New York, 1993. C.M. Barshick, D.C. Duckworth and D.H. Smith, Inorganic Mass Spectrometry: Fundamentals and Applications. Marcel Dekker, New York, 2000. N.P. Vela, L.K. Olson and J.A. Caruso, Anal. Chem., 65 (1993) 585A. P. Uden, in: P. Uden (Ed.), Element-Speciﬁc Chromatographic Detection by Atomic Emission Spectroscopy, ACS Series 479, American Chemical Society, Washington, DC, 1990. F.A. Cotton, G. Wilkinson, C.A. Murillo and M. Bochmann, Advanced Inorganic Chemistry, 6th edn., Wiley, New York, 1999. T.W.G. Solomons, Organic Chemistry, 6th edn., Wiley, New York, 1996. A.G. Davies and P.J. Smith, in: G. Wilkinson, F.G.A. Stone and E.W. Abel (Eds.), Organometallic Chemistry, The Synthesis, Reactions and Structures of Organometallic Compounds, Tin, Pergamon Press, Oxford, 1982, Ch. 11. Y.K. Chau, P.T.S. Wong, G.A. Bengert and J.L. Dunn, Anal. Chem., 56 (1984) 271. Y. Cai, R. Jaffe´ and R.D. Jones, Environ. Sci. Technol., 31 (1997) 302. S. Monsalud, M.S. Thesis, Florida International University, 1999. Y. Cai, in: R.A. Meyers (Ed.), Encyclopedia of Analytical Chemistry. Wiley, Chichester, 2000, p. 2270. J. Szpunar-Lobinska, C. Witte, R. Lobinski and F. Adams, Fresenius J. Anal. Chem., 351 (1995) 351. U.M. Gru¨ter, M. Hitzke, J. Kresimon and A.V. Hirner, J. Chromatogr. A, 938 (2001) 225. O.F.X. Donard and F.M. Martin, Trends Anal. Chem., 11 (1992) 17. J.P. Craig, D. Mennie, N. Ostah, O.F.X. Donard and F. Martin, Analyst, 117 (1992) 823. M. Fillppelli, F. Baldi, E.F. Brinckman and J.G. Olson, Environ. Sci. Technol., 26 (1992) 1457. R. Irizarry, J. Moore and Y. Cai, Int. J. Environ. Anal. Chem., 79 (2001) 97. S. Rapsomanikis, Analyst, 119 (1994) 1429. Y. Cai, S. Monsalud and K.G. Furton, Chromatographia, 52 (2000) 82. S. Rapsomanikis, O.F.X. Donard and J.H. Weber, Anal. Chem., 58 (1986) 38. H. Hintelmann, M. Hemple and R.D. Wilken, Environ. Sci. Technol., 29 (1995) 1845.

Derivatization and vapor generation methods for trace element analysis and speciation 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64

T. De Smaele, L. Moens, R. Dams, P. Sandra, J. Van der Eycken and J. Vandyck, J. Chromatogr. A, 793 (1998) 99. V. Lu¨ckow and H.A. Russel, J. Chromatogr., 150 (1978) 187. V. Minganti, R. Capelli and R. De Pellegrini, Fresenius J. Anal. Chem., 351 (1995) 471. G. Hu, X. Wang, Y. Wang, X. Chen and L. Jia, Anal. Lett., 30 (1997) 2579. Y.K. Chau and P.T.S. Wong, in: G.G. Leppard (Ed.), Trace Element Speciation in Surface Waters and Its Ecological Implications. Plenum Press, New York, 1983. Y.K. Chau, R.J. Maguire, M. Brown, F. Yang and S.P. Batchelor, Water Qual. Res. J. Canada, 32 (1997) 453. Y. Cai, S. Rapsomanikis and M.O. Andreae, Anal. Chim. Acta, 274 (1993) 243. Y. Cai, S. Rapsomanikis and M.O. Andreae, J. Anal. At. Spectrom., 8 (1993) 119. Y. Cai, S. Rapsomanikis and M.O. Andreae, Talanta, 41 (1994) 589. R. Fischer, S. Rapsomanikis and M.O. Andreae, Anal. Chem., 65 (1993) 563. A. Astruc, R. Lavigne, V. Desauziers, R. Pinel and M. Astruc, Appl. Organomet. Chem., 3 (1989) 267. Y. Cai, S. Rapsomanikis and M.O. Andreae, Microchim. Acta, 109 (1992) 67. L. Liang, M. Horvat and N.S. Bloom, Talanta, 41 (1994) 371. Y.K. Chau, P.T.S. Wong and G.A. Bengert, Anal. Chem., 54 (1982) 246. J.A. Stab, W.P. Coﬁno, B. van Hattum and U.A.T. Brinkman, Fresenius J. Anal. Chem., 347 (1993) 247. C.A. Krone, D.W. Brown, D.G. Burrows, R.G. Bogar, S.L. Chan and U. Varanasi, Mar. Environ. Res., 27 (1989) 1. T. Ishizaka, S. Nemoto, K. Sakasi, T. Suzuki and Y. Saito, J. Agric. Food Chem., 37 (1989) 1523. W.R. Cullen, G.K. Eigendorf, B.U. Nwata and A. Takatsu, Appl. Organomet. Chem., 4 (1990) 581. S. Ohhira and H. Matsui, Bull. Environ. Contam. Toxicol., 44 (1990) 294 –301. Y.K. Chau, S. Zhang and R.J. Maguire, Analyst, 117 (1992) 1161. B.F. Scott, Y.K. Chau and A. Rais-Firouz, Appl. Organomet. Chem., 5 (1991) 151. C.A. Krone, D.W. Brown, D.G. Burrows, R.G. Bogar, S.L. Chan and U. Varanasi, Mar. Pollut. Bull., 20 (1989) 528. R. Lobinski, C.F. Boutron, J.P. Candelone, S. Hong and J. Szpunar-Lobinski, Anal. Chem., 65 (1993) 2510. R.J.A. van Cleuvenbergen and F.C. Adams, Environ. Sci. Technol., 26 (1992) 1354. K. Krishnan, W.D. Marshall and W.I. Hatch, Environ. Sci. Technol., 22 (1988) 806. Y.K. Chau, P.T.S. Wong, G.A. Bengert and J.L. Dunn, Anal. Chem., 56 (1984) 271. Y.K. Chau, P.T.S. Wong, G.A. Bengert and O. Kramer, Anal. Chem., 51 (1979) 186. J.P. Snell, W. Frech and Y. Thomaseen, Analyst, 121 (1996) 1055. Ph. Quevauviller, G.U. Fortunati, M. Filippelli, F. Baldi, M. Bianchi and H. Muntau, Appl. Organomet. Chem., 10 (1996) 537. M.O. Andreae, Anal. Chem., 49 (1977) 820. S.H. Vien and R.C. Fry, Anal. Chem., 60 (1988) 465. M.A. Morrison and J.H. Weber, Appl. Organomet. Chem., 11 (1997) 761. O.F.X. Donard and J.H. Weber, Environ. Sci. Technol., 19 (1985) 1104. O.F.X. Donard, S. Rapsomanikis and J.H. Weber, Anal. Chem., 58 (1986) 772. V.F. Hodge, S.L. Seidel and E.D. Goldberg, Anal. Chem., 51 (1979) 1256. J.J. Cooney, A.T. Kronick, G.J. Olson, W.R. Blair and F.E. Brinckman, Chemosphere, 17 (1988) 1795. J.S. Han and J.H. Weber, Anal. Chem., 60 (1988) 316.

591

Y. Cai 65 66 67 68 69 70 71 72 73 74 75 76 77 78

592

C.L. Matthias, J.M. Bellama, G.J. Olson and F.E. Brinckman, Int. J. Environ. Anal. Chem., 35 (1989) 61. J.G. Bin, Z.Q. Fang and H. Bin, Environ. Sci. Technol., 34 (2000) 2697. ¨ . Ay and E. Henden, Spectrochim. Acta B, 55 (2000) 951. U J.S. Blais and W.D. Marshall, J. Anal. At. Spectrom., 4 (1989) 641. B.J. Feldman, H. Mogadeddi and J.D. Osterloh, J. Chromatogr., 594 (1992) 275. T. Gorecki and J. Pawliszyn, Anal. Chem., 68 (1996) 3008. J.R. Ashby and P.J. Craig, Sci. Tot. Environ., 78 (1989) 219. M. Ceulemans, J. Szpunar-Lobinski, W.M.R. Dirkx, R. Lobinski and F.C. Adams, Int. J. Environ. Anal. Chem., 52 (1993) 113. C. Carlier-Pinasseau, G. Pespes and M. Astruc, Talanta, 44 (1997) 1163. A. D’Ulivo and Y. Chen, J. Anal. At. Spectrom., 4 (1989) 319. V. Clamagirand, I.L. Marr and J.L. Wardell, Appl. Organomet. Chem., 7 (1993) 577. N. Bloom, Can. J. Fish Aquat. Sci., 46 (1989) 1131. N. Demuth and K.G. Heumann, Anal. Chem., 73 (2001) 4020. X. Yu and J. Pawliszyn, Anal. Chem., 72 (2000) 1788.

Chapter 20

Laser ablation sampling Richard E. Russo and David P. Baldwin

20.1

INTRODUCTION

Laser ablation sampling is the generation of a vapor-phase aerosol using a pulsed high-power laser beam focused on a sample surface. Any sample (solid, liquid, organic, inorganic, opaque, transparent, conducting, insulator, etc.) can be laser ablated. There are no sample-size or shape requirements; bulk samples as well as individual micrometer-sized particles can be directly analyzed. Depending on the analytical measurement system sensitivity, microgram to femtogram sample quantities may be sufﬁcient for analysis, rendering laser ablation sampling nearly non-destructive. Finally, a focused laser beam permits spatial characterization of heterogeneity in samples with micrometer lateral resolution and sub-micrometer depth resolution. A major attraction of laser ablation is that little or no sample preparation is required; a small portion of the sample is directly converted into vapor-phase aerosol constituents by the laser beam. The simplest application is bulk analysis when the sample is homogenous. If the sample is inhomogeneous, it can be pulverized to a ﬁne powder and pressed or fused into a solid form. On the other hand, the spatial properties of the laser beam allow the identiﬁcation of heterogeneity in the sample. For example, a single laser pulse on a sample can be used to study surface contamination or it can be used to clean the surface in order to study underlying bulk composition. The primary emphasis of laser ablation research is not on sample preparation, but studying the ablation processes [1 –6]. Choosing the correct laser properties for the particular sample deﬁnes the ability to use laser ablation sampling for accurate chemical analysis. The properties of the laser beam that inﬂuence ablation sampling are pulse duration, energy, wavelength, and spatial energy proﬁle. The irradiance (energy per unit time and area) plays the dominant role in deﬁning the quantity and chemistry of the ablated aerosol. This chapter emphasizes research studies in laser ablation without emphasis on the numerous applications. Current research in laser ablation sampling addresses the optimum choice of laser properties and quantitative analysis. The search for optimum Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

593

R.E. Russo and D.P. Baldwin

laser properties is based on the need to ablate samples without matrix dependence and without fractionation (ablated mass vapor is not chemically equal to the original sample). Fractionation can be described in two ways, nonstoichiometric ablation from a single laser pulse or the time-dependent change in elemental ratio as a crater is formed during repetitive laser pulsing at one sample location. A goal of laser ablation sampling has been to either eliminate fractionation or ensure that it is the same when ablating the sample and standards. Fractionation would not be problematic if the sample ablated the same way as the standards, or if the entire sample were ablated (particle). External or internal calibration is required for quantitative analysis because the amount of mass ablated can depend on the sample matrix (chemical, physical, and optical properties). The quantity of mass ablated must be measured or it must be equal to that ablated from appropriate standards. A goal of laser ablation research has been to identify experimental conditions under which the matrix dependence is eliminated. In this case, non-matrix matched standards could be utilized for calibration (i.e. general classes of materials that have similar but not identical ablation behavior under appropriate laser conditions). 20.2

EXPERIMENTAL SYSTEM

A typical laser ablation sampling system consists of a laser, an ablation chamber, and a detection system. Pulsed lasers are used almost exclusively for achieving ablative (explosive) conditions. Samples are placed in a chamber that is attached to an adjustable positioning stage. The laser beam is focused onto the sample surface, converting a small portion of mass (solid or liquid) into a vapor-phase aerosol. Detection is based on either measuring ionic/atomic emission in the vapor plume (plasma) at the sample surface (LIBS) (Fig. 20.1) or by transporting the ablated material to an ICP (MS or OES) (Fig. 20.2). Argon or helium gas typically carries the ablated sample to the ICP. An example of craters in metal samples is shown in Fig. 20.3. Almost all commercial laser ablation systems for chemical analysis utilize nanosecond-pulsed Nd:YAG or excimer lasers [7 –12]. Although the fundamental wavelength is at 1064 nm, optical second, third, fourth, and ﬁfth harmonics (wavelengths at 532, 355, 266, and 213 nm) of Nd:YAG lasers have been investigated for laser ablation sampling. The pulse duration from these lasers is generally 3–6 ns. Excimer lasers produce wavelengths of 308, 248, 193, or 157 nm, using XeCl, KrF, ArF, or F2, respectively. Pulse durations from excimer lasers are generally in the range 15–35 ns. The quantity of laser energy absorbed by the sample is inﬂuenced by laser wavelength. However, a fundamental correlation between laser wavelength and optical absorption has not been established because of the high peak powers (short pulse duration)

594

Laser ablation sampling

Fig. 20.1. LIBS experimental system conﬁguration.

595

596 R.E. Russo and D.P. Baldwin

Fig. 20.2. ICP (OES, MS) experimental system conﬁguration.

Laser ablation sampling

Fig. 20.3. Crater images.

597

R.E. Russo and D.P. Baldwin

and resultant non-linear processes underlying ablation mechanisms. Another process that depends on laser wavelength is shielding by the laser-induced plasma near the sample surface [13–18]. The laser-beam energy can be absorbed or reﬂected by this plasma. Some of the plasma energy may subsequently be transferred to the surface through thermal or radiative processes. Laser – plasma interactions are typically stronger at longer wavelengths. In general, greater ablation efﬁciency (amount of mass removed per unit energy), reduced plasma shielding, and reduced fractionation are realized by using short laser wavelengths (UV) and short laser-pulse durations. A current research trend is the investigation of picosecond and femtosecond pulses for laser ablation sampling [19–28]. The most common ultrashort pulsed laser is the solid-state Ti:Sapphire; the pulse duration can be compressed (fs) and stretched (ps) using an optical grating pair. Ablation mechanisms are governed by the amount of energy deposited in the sample per unit time and volume. From a classical point of view, a material system can accommodate an energy impulse through thermodynamic pathways: absorption, diffusion, melting, boiling, and vaporization. For picosecond and femtosecond lasers, the pulse duration can be comparable to or shorter than the phonon relaxation time; i.e., the laser energy can be deposited into the material before it can thermally equilibrate. Ideally, this type of interaction will lead to more of a photo-physical bond breaking process instead of classical melting, boiling, and vaporization. The heat-affected zone in a sample for a short laser pulse is small. The thermal diffusion length is on the order of nanometers for picosecond and femtosecond laser pulses compared to about a micrometer for nanosecond pulses. By using short (ps, fs) laser pulses with UV wavelengths, the optical and thermal penetration depth will be on the order of nanometers. A goal of using short laser pulses is to ablate (remove) the entire optical and heat-affected volumes so that elemental migration and fractionation are negligible. In addition, the short pulse regime may be less susceptible to the material properties, thereby providing matrix-independent sampling. The short pulse regime provides additional beneﬁts for laser ablation sampling. The reduced heat-affected zone and lower energy threshold for ablation lead to shallower crater depths per pulse. Therefore, depth resolution can be much ﬁner using fs versus ns pulses. The crater may not possess a raised rim and the surrounding area on the sample surface can have signiﬁcantly less debris using femtosecond versus nanosecond pulsed lasers (cf. Fig. 20.3). The absence of a raised rim is related to reduced melting and a different plasma mechanism for short laser-pulse ablation. The interaction of the vapor plasma with the laser-beam energy is signiﬁcantly reduced for picosecond pulses and eliminated for femtosecond pulses. The laser-beam energy proﬁle also inﬂuences spatial and depth resolution. Generally, solid-state lasers (Nd:YAG, Ti:Sapphire) have Gaussian beam proﬁles, whereas gas lasers (excimers) possess more of a ﬂat-top energy proﬁle.

598

Laser ablation sampling

With a Gaussian laser-beam energy proﬁle, the shape of ablation craters in the sample can be cone-like, reﬂecting the energy proﬁle. Flat-top energy proﬁles are better suited for producing straight-wall craters with ﬂat bottoms. However, by using appropriate optics, all of these lasers can produce such craters. For depth proﬁling of multilayers, a ﬂat-top proﬁle is preferred to ensure the integrity of ablation through individual layers. Depth resolution is governed by the laser parameters (energy, ﬂuence, wavelength, and pulse duration) and sample properties (optical and thermal), and can range from nanometers to micrometers. For lateral spatial resolution, laser beams are either focused or imaged to diameters on the order of several micrometers. In theory, a Gaussian laser beam can be focused to diffraction-limited conditions producing a spot diameter somewhat larger than the wavelength. In practice, most systems deliver a beam with a diameter ranging from several micrometers to hundreds of micrometers. In most cases, the crater diameter will equal the laser-beam diameter. Several studies have investigated the use of near-ﬁeld scanning optical microscopy (NSOM) and atomic force microscopy (AFM) for ablating samples with nanometer-diameter resolution [29– 31]. The pulse repetition rate from the above-mentioned lasers can be varied from single pulse to kHz. For spatially resolved depth analysis of inhomogeneous samples or for particle analysis, a single laser pulse may be preferred for ablation sampling. In this case, the ablated mass will be produced as a temporal transient. For LIBS, the properties of the plasma will deﬁne the signal temporal proﬁle. For the ICP-MS, the temporal proﬁle will be characteristic of the chamber volume, transport-tube length and diameter, and carrier gas composition and ﬂow rate. For homogeneous samples, a repetitively pulsed laser allows the sampled material from successive ablations to mix, providing a continuous signal. Sampling and analytical precision are improved signiﬁcantly with repetitively pulsed laser ablation compared to single-pulse ablation. Repetitive sampling and continuous aerosol production simplify optimization of analytical detection systems. 20.3

ABLATION DETECTION SYSTEMS

There are two primary conﬁgurations for detecting laser-ablated sample: directly in the laser-induced plasma near the sample surface (LIBS) (cf. Fig. 20.1) or by transporting the ablated sample to a secondary excitation source, such as an ICP (cf. Fig. 20.2). The LIBS technique offers the advantage of remote analysis whereas ICP methods offer signiﬁcantly lower detection limits. LIBS is a simple technique to implement [32–40]. The focused laser beam creates a luminous plasma near the sample surface. The sample does not need to be in an enclosed chamber; LIBS can be performed for remote sampling applications. A single lens or optical ﬁber collects the optical emission from this plasma, which is imaged into a spectrometer. The analytical capabilities of

599

R.E. Russo and D.P. Baldwin

LIBS are determined by the plasma properties, which depend on experimental parameters, including the laser pulse (energy, duration, repetition rate, and wavelength), the sample (physical and optical), and ambient atmosphere (gas, pressure). Laser-induced plasma emission is transient in time and consists of atomic and ionic spectral lines superimposed on a broadband continuum. Both integrated and time-resolved measurements have been utilized for chemical analysis in laser-induced plasmas. Due to the in situ nature of these measurements, care must be taken to avoid self-absorption of the analyte optical lines by the plasma or surrounding vapor. Most LIBS research and applications have been based on the use of nanosecond pulsed lasers. Plasma background and ionic emission exist several hundred nanoseconds after the laser pulse, whereas atomic emission can last for several microseconds. Time delayed gating is generally required for analytical detection to eliminate the initial broadband background radiation. Alternatively, a high-resolution spectrometer may be used to resolve the narrow analytical emission features from the broad background interferences [41]. Several studies have explored the use of ultrashort (ps and fs) pulsed lasers for LIBS [42 –47]. The spatial extent, growth, and decay of ionic and atomic emission lines are signiﬁcantly changed. The emission lifetime of spectral lines is at least an order of magnitude shorter and the continuum background is signiﬁcantly lower than that arising from nanosecond induced plasmas. Nongated detection has been demonstrated because of low background with respect to well-deﬁned atomic lines. Surprisingly, the analytical ﬁgures of merit reported to date are similar for nanosecond and femtosecond LIBS measurements. Research is only beginning in this area and improved analytical ﬁgures of merit are expected. The ICP is the most widely used detection system for chemical analysis using laser ablation sampling [48–55]. A typical ablation system using the ICP consists of a sample chamber with a window (transparent to the laser wavelength), a CCD camera to view the sample, and an adjustable positioning platform. An inert gas is used for entrainment and transport of the ablated sample vapor to the ICP. The effect of transport gas on laser ablation has been investigated to improve analytical performance for ICP-MS (and LIBS) [56–60]. Argon and helium have been employed with the latter gas providing improved ablation and transport efﬁciencies. The degree of enhancement is related to the wavelength, pulse duration, and irradiance. The gas enhancement was found to be laser-irradiance dependent. The largest enhancements are about a factor of approximately two- to threefold using nanosecond lasers. In support of the enhanced ICP data, crater volumes generated by ablation in the He atmosphere were larger. There was also less deposition of particles around the ablation crater rim using He. Larger enhancements (6– 10-fold) were observed using picosecond laser ablation in He versus Ar. Plasma shielding, which reduces the laser energy coupled into the sample, provides a possible mechanism to explain gas effects. The degree of plasma shielding

600

Laser ablation sampling

depends on the ionization potential of the gas species; a plasma can form more readily in easily ionized gases. Other than noble gases, nitrogen and oxygen were also reported to improve the sensitivity during LA-ICP-MS analysis. In general, chamber characteristics and transport properties need to be further studied and optimized for improved laser ablation sampling. The ablated aerosol is entrained in the transport gas and transported through plastic tubing to the inductively coupled plasma. The majority of particles are vaporized very low in the ICP and the vapor is atomized, ionized, and electronically excited. Atomic emission in this source may be used to analyze the composition of the ablated material from the weight-percent to ppm range [4]. Mass spectrometry of the ions created in the ICP provides sensitivity from weight-percent to as low as the ppb to ppt ranges [4]. Although not an in situ technique, ICP provides a source that is less prone to matrix effects than LIBS and provides signiﬁcantly improved sensitivity and reduced interferences when combined with mass spectrometry. Selection of an appropriate analytical detection system will often depend on factors unrelated to the factors discussed above. Presumably, laser ablation sampling is chosen as opposed to physical sample collection, followed by digestion and analysis, due to factors such as the speed afforded with no sample preparation, need for real-time analyses, the hazardous nature of the samples, the small size of samples, elimination of secondary wastes, or the desire for minimally destructive sampling. These same factors may dictate the appropriate detection system. LIBS may be performed in situ and therefore may be desirable due to the difﬁculty in extracting a sample for analysis or the hazards or contamination issues associated with collecting even a minute amount of material and introducing it into an ICP (e.g. if the sample is highly radioactive). On the other hand, survivability of the laser, optics, detector, and optical ﬁbers for a LIBS system in a high radioactivity environment might dictate the need to transport the ablated material to a remote location for analysis, despite contamination risks. The size, cost, and portability of the system (including the laser) may be important considerations, and a mass spectrometer is signiﬁcantly more expensive than the optical spectrometers used in LIBS or ICP-OES analyses. Availability of utilities for the laser system as well as the detection system may play a role. An ICP requires noble gases, cooling water, kilowatts of electrical power, and an exhaust system. If isotopic information is required, ICP-MS may be the only solution except with extremely high-resolution optical spectrometers for a few speciﬁc isotopes. Finally, in the absence of matrixmatched standards, LIBS is much more difﬁcult to calibrate than ICP. 20.4

CALIBRATION

For quantitative analysis, there are two general approaches for calibrating the detection system, utilizing external calibration with standards [61–78]

601

R.E. Russo and D.P. Baldwin

or internal calibration of the sample [79,80]. For the most part, external calibration requires that the standards match the sample in ablation behavior. Elements used for internal standardization need to be present with known concentration, homogeneously distributed, and behave similarly to the majority of other elements during the laser ablation process (similar or no fractionation). In the ideal case when matrix-matched standards are available, calibration is straightforward. A suite of standards is ablated to generate a plot of intensity versus concentration. The unknown sample is ablated using the same laser and detection-instrument conditions. Matrix matching is necessary because the ablation rate (quantity of mass ablated per laser pulse) and fractionation can vary with the sample matrix (matrix dependence). External calibration with matrix-matched standards has been used successfully for quantiﬁcation in a number of applications [4]. Several groups have worked to generate universal calibration curves. By optimizing the laser ablation process, it may be possible to use non-matrix matched standards. Differences in ablation rate between the sample and the non-matched standard may be corrected using a number of methods. The signal levels for minor constituents may be normalized to the signal level for a major element whose concentration is known for both the sample and the standard (e.g. use the gold elemental signal level in a gold sample or the known iron level in a steel sample). If matrix-matched standards do not exist or cannot be fabricated, external calibration for ICP analyses can be performed using other sample introduction technologies. One strategy is to use two sample introduction systems, one for the ablated sample and another for nebulized aqueous solution standards [72–76]. Direct liquid ablation also has been demonstrated as a calibration strategy [77,78]. However, the ICP with laser ablation sampling is “dry” relative to that produced using liquid sample introduction. The introduction of water vapor or droplets to the ICP can result in elevated oxide and other molecular interferences in mass spectrometry. Nebulization and ablation may also result in different particle size distributions and morphology that could result in different behavior in the ICP. Therefore, analyte response from the two introduction methods can differ to some extent. If the solution aerosols are desolvated before they enter the ICP, the response from the calibration aerosol will be closer to that of laser ablation sampling. In any case, it is critical to ensure that the ICP conditions are matched when calibrating against secondary technologies. Surveys of calibration methods are available in several recent reviews [1–5]. Many approaches have been demonstrated to monitor and compensate for changes in the ablated mass quantity [1 –5]. They include light scattering from the ablated plume, acoustic monitoring in the sample, monitoring acoustic and optical emission in the atmosphere above the sample, weighing the sample before and after ablation, or by using a mass monitor to collect

602

Laser ablation sampling

a portion of the vapor during the ablation sampling process. However, none of these schemes for using non-matched standards can correct for fractionation. A relative measure of mass ablated can be achieved by using an internal standard, simultaneously measuring emission from the analyte element and a common matrix element [79,80]. The spatial distribution of the internal standard must be homogeneous and the internal standard and analyte must be equally affected by ablation sampling (no fractionation). With this approach, a ratio of elements accounts for the change in quantity of mass ablated per laser shot. For homogeneous samples, internal standardization has been used to improve measurement precision to better than 1%. In some instances, more than one of these measures may be combined to address different aspects of laser ablation calibration. For example, mass monitoring may be employed to correct for differences between ablation rates for multiple repetitions or between samples and standards. A solution standard may be nebulized and dried prior to mixing with the ablated material in order to calibrate instrumental response and correct for matrix effects in the ICP, similar to the use of isotope dilution in solution analyses. Finally, an internal standard may be used to correct for variations in ablation rates between mass measurements or over the course of an analysis using many laser shots, whether the variation occurs due to the evolution of a crater at a single spot or due to changes encountered while raster scanning across a heterogeneous sample. 20.5

FRACTIONATION

Understanding and eliminating elemental fractionation has been one of the most signiﬁcant research agendas in laser ablation sampling [81–95]. It is important to point out that the occurrence of fractionation does not preclude the use of laser ablation sampling for accurate chemical analysis. Numerous successful applications have been performed utilizing external and internal calibration procedures to correct for fractionation [4]. Fractionation is a function of laser-beam properties (irradiance, pulse duration, wavelength). Ablation mechanisms and fractionation are not directly related to the beam proﬁle; i.e., fractionation is not eliminated using a ﬂat-top beam proﬁle. Fundamental studies have attempted to correlate fractionation with elemental properties such as melting and boiling points, vapor pressure, atomic or ionic radius, charge, and speciation. However, no unifying theory exists to describe or predict ablation processes. The wavelength effect on fractionation is currently of great interest [87–90]. In general, shorter wavelengths reduce fractionation. However, fractionation can be increased or reduced using all wavelengths if the respective lasers have enough variability of parameters (energy, irradiance). From ﬁrst principles, ablation seems related to the optical

603

R.E. Russo and D.P. Baldwin

penetration depth in a material—the shorter the wavelength, the shallower the optical penetration depth. Therefore, more energy is available for ablation per unit volume in the sample. This description assumes linear one-photon absorption, which is not strictly the case when using high peak power laser beams. Fractionation is especially noticed as a crater develops using a repetitively pulsed laser at a single sample location. An exact mechanism for crater effects on fractionation is not established. One hypothesis is that the plasma conﬁned inside a deep crater may contribute to the sampling process, leading to fractionation. Another possibility is that actual irradiance decreases as the crater deepens due to changes of effective area exposed to the laser beam. However, fractionation can occur for a single laser pulse without crater formation. Fractionation during transport involves selective vapor condensation on tubing walls, or selective nucleation of species on different sized particles. Transport efﬁciency and chemical composition are particle-size dependent. Fractionation can be signiﬁcantly reduced simply by placing bends in the transport tubing, packing the tubing with plugs of glass wool, or using in-line impactors. In addition to atomic and molecular vapor (the basis of LIBS), laser ablation sampling produces particles with a size distribution ranging from nanometers to micrometers. The size distribution depends on the laser irradiance, wavelength, ambient gas and pressure, and properties of the sample [91– 95]. Sample entrainment, transport, and detection system (ICP-MS) response are inﬂuenced by the particle size distribution. A general belief is that particles should be less than 1 mm in diameter for efﬁcient transport and complete decomposition/dissociation in the ICP. The particle inﬂuence on fractionation and analytical performance is a critical area that needs to be thoroughly investigated. 20.6

CONCLUSION

Laser ablation sampling is a powerful analytical technology. The beneﬁt of a universal sampling technology for non-destructive analysis is driving its development. Experimental parameters along with the challenges related to calibration and fractionation have been discussed. The issue emphasized throughout this chapter has been that ablation sampling is reliant on the laser properties. In fact, laser preparation is signiﬁcantly more important than sample preparation. In spite of all the issues discussed, good calibration of analytical detection systems and accurate chemical analyses are achieved when the sample and standards are ablated using the same laser parameters (Table 20.1) (cf. Refs. [1 –5]). There will always be advantages and disadvantages when comparing ablation systems due to the different speciﬁcations of

604

Laser ablation sampling TABLE 20.1 Examples of analytical laser ablation applications Matrix

Laser

Detection

Reference

Biological In vivo rat tissue near implants Tree rings

266-nm Nd:YAG 266-nm Nd:YAG

MS MS

[99] [67]

Environmental Recyclable thermoplastic polymers Road sediments Soil Stack emissions Stack emissions

266-nm Nd:YAG 266-nm Nd:YAG 532-nm Nd:YAG 532-nm Nd:YAG 1064-nm Nd:YAG

LIBS MS OES LIBS LIBS

[96] [100] [104] [105] [110]

Forensic Paint pigments Cannabis Gold Steel and glass Ink

355-nm Nd:YAG 1064-nm (NQ) Nd:YAG 1064-nm (NQ) Nd:YAG 1064-nm (NQ) Nd:YAG 266-nm Nd:YAG

LIBS MS MS MS MS

[108] [113] [114] [115] [116]

Geochemical Minerals Sulﬁde and silicate inclusions Copper ore deposits Iron in meteorites Garnets Coal

213-nm Nd:YAG 193-nm excimer 193-nm excimer 266-nm Nd:YAG 266-nm Nd:YAG 1064-nm Nd:YAG

MS MS MS MS MS LIBS

[90] [97] [98] [101] [102] [107]

Process monitoring Molten aluminum Coatings Glass Fire suppressants and refrigerants Glass

1064-nm Nd:YAG 532-nm Nd:YAG 532-nm Nd:YAG 1064-nm Nd:YAG 1064-nm Nd:YAG

LIBS LIBS LIBS LIBS LIBS

[103] [106] [109] [111] [112]

MS ¼ LA-ICP-MS, OES ¼ LA-ICP-OES(AES), NQ ¼ non-Q-switched.

the selected laser. For fundamental scientiﬁc studies, it is critical to ensure that all parameters (except the parameter of interest) are equal before attempting models or descriptions of ablation processes. Such research studies will lead to improvements in existing applications and pave the way for new opportunities in laser ablation sampling.

605

R.E. Russo and D.P. Baldwin

Acknowledgements We thank the authors, whose work was referenced in this chapter, for contributing to this exciting technology. RER acknowledges support by the US Department of Energy, Ofﬁce of Basic Energy Sciences, Division of Chemical Sciences, and the Ofﬁce of Nonproliferation and National Security at the Lawrence Berkeley National Laboratory under Contract No. DE-AC0376SF00098. DPB acknowledges support of the Ofﬁce of Nonproliferation and National Security at the Ames Laboratory under Contract No. W-7405-ENG-82. REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22

23 24 25 26

606

R.E. Russo, X.L. Mao and S.S. Mao, Anal. Chem., 74 (2002) 70A. K. Niemax, Fresenius J. Anal. Chem., 370 (2001) 332. I. Horn, M. Guillong and D. Gunther, Appl. Surf. Sci., 182 (2001) 91. J.D. Winefordner, I.B. Gornushkin, D. Pappas, O.I. Matveev and B.W. Smith, J. Anal. At. Spectrom., 15 (2000) 1161. J.F. Durrant, J. Anal. At. Spectrom., 12 (1999) 1385. W. Demtroder, Laser Spectroscopy. Springer, New York, 1998. R.E. Russo, X.L. Mao, H.C. Liu, J. Gonzalez and S.S. Mao, Talanta, 57 (2002) 425. C. Latkoczy and D. Gu¨nther, J. Anal. At. Spectrom., 17 (2002) 1264. M. Ohata, Y. Iwasaki, N. Furuta and I. Brenner, Spectrochim. Acta B, 57 (2002) 1713. V. Kanicky, V. Otruba and J.M. Mermet, Fresenius J. Anal. Chem., 371 (2001) 934. D. Gunther, I. Horn and B. Hattendorf, Fresenius J. Anal. Chem., 368 (2000) 4. R.E. Russo, X.L. Mao and O.V. Borisov, Trends Anal. Chem., 17 (1998) 461. S.S. Mao, X.L. Mao, R. Greif and R.E. Russo, Appl. Phys. Lett., 77 (2000) 2464. M.H. Hong, Y.F. Lu and S.K. Bong, Appl. Surf. Sci., 154 (2000) 196. J.M. Vadillo, J.M.F. Romero, C. Rodriguez and J.J. Laserna, Surf. Interface Anal., 27 (1999) 1009. H.C. Liu, X.L. Mao, J.H. Yoo and R.E. Russo, Spectrochim. Acta B, 54 (1999) 1607. X.L. Mao and R.E. Russo, Appl. Phys. A, 64 (1997) 1. C.R. Phipps and R.W. Dreyfus, in: A. Vertes, R. Gijbels and F. Adams (Eds.), Laser Ionization Mass Analysis. Wiley, New York, 1993. R.E. Russo, X.L. Mao, J.J. Gonzalez and S.S. Mao, J. Anal. At. Spectrom., 17 (2002) 1072. M.K. Kim, T. Takao, Y. Oki and M. Maeda, Jpn. J. Appl. Phys., 39 (2000) 6277. V. Margetic, M. Bolshov, A. Stockhaus, K. Niemax and R. Hergenroder, J. Anal. At. Spectrom., 16 (2001) 616. A. Semerok, C. Chaleard, V. Detalle, J.L. Lacour, P. Mauchien, P. Meynadier, C. Nouvellon, B. Salle, P. Palianov, M. Perdrix and G. Petite, Appl. Surf. Sci., 138– 139 (1999) 311. J.G. Mank and P.R.D. Mason, J. Anal. At. Spectrom., 14 (1999) 1143. A. Cavalleri, K. Sokolowski-Tinten and D. von der Linde, Appl. Phys. Lett., 72 (1998) 2385. X.L. Mao, O.V. Borisov and R.E. Russo, Spectrochim. Acta B, 53 (1998) 731. S. Nolte, C. Momma, H. Jacobs, A. Tunnermann, B.N. Chichkov, B. Wellegehausen and H. Welling, J. Opt. Soc. Am. B, 14 (1997) 2716.

Laser ablation sampling 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41

42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57

B.N. Chichkov, C. Momma, S. Nolte, F. von Alvensleben and A. Tunnermann, Appl. Phys. A, 63 (1996) 109. V. Kanicky, J. Musil, I. Novotny and J. Mermet, Appl. Spectrosc., 51 (1997) 1042. Y. Ding, R. Micheletto, H. Hanada, T. Nagamura and S. Okazaki, Rev. Sci. Instrum., 73 (2002) 3227. D. Kossakovski and J.L. Beauchamp, Anal. Chem., 72 (2000) 4731. B. Dutoit, D. Zeisel, V. Deckert and R. Zenobi, J. Phys. Chem. B, 101 (1997) 6955. L.J. Radziemski, Spectrochim. Acta B, 57 (2002) 1109. A. Fisher, M.W. Hinds, S.N. Nelms, D.M. Penny and P. Goodall, J. Anal. At. Spectrom., 17 (2002) 1624. K. Song, Y.I. Lee and J. Sneddon, Appl. Spectrosc. Rev., 37 (2002) 89. G.W. Rieger, M. Taschuk, Y.Y. Tsui and R. Fedosejevs, Appl. Spectrosc., 56 (2002) 689. D. Anglos, K. Melesanaki, V. Zaﬁropulos, M.J. Gresalﬁ and J.C. Miller, Appl. Spectrosc., 56 (2002) 423. B. Fairman, M.W. Hinds, S.M. Nelms, D.M. Penny and P. Goodall, J. Anal. At. Spectrom., 16 (2001) 1446. K.L. Eland, D.N. Stratis, T. Lai, M.A. Berg, S.R. Goode and S.M. Angel, Appl. Spectrosc., 55 (2001) 279. M. Capitelli, G. Colonna, M. Catella, F. Capitelli and A. Eletskii, Chem. Phys. Lett., 316 (2000) 517. D.A. Rusak, B.C. Castle, B.W. Smith and J.D. Winefordner, Crit. Rev. Anal. Chem., 27 (1997) 257. D.P. Baldwin, D.S. Zamzow, D.K. Ottesen and H.A. Johnsen, Testing of an Echelle Spectrometer as a LIBS Detector at Sandia, Ames Laboratory Report IS-5148, 2001. V. Margetic, A. Pakulev, A. Stockhaus, M. Bolshov, K. Niemax and R. Hergenroder, Spectrochim. Acta B, 55 (2002) 1771. A. Semerok, B. Salle, J.F. Wagner and G. Petite, Laser Particle Beams, 20 (2002) 67. V. Margetic, K. Niemax and R. Hergenroder, Spectrochim. Acta B, 56 (2001) 1003. B. Le Drogoff, J. Margot, M. Chaker, M. Sabsabi, O. Barthelemy, T.W. Johnston, S. Laville, F. Vidal and Y. von Kaenel, Spectrochim. Acta B, 56 (2001) 987. K.L. Eland, D.N. Stratis, D.M. Gold, S.R. Goode and S.M. Angel, Appl. Spectrosc., 55 (2001) 286. S.M. Angel, D.N. Stratis, K.L. Eland, T. Lai, M.A. Berg and D.M. Gold, Fresenius J. Anal. Chem., 369 (2001) 320. T. Hirata, Anal. Chem., 75 (2003) 228. I. Rodushkin, M.D. Axelsson, D. Malinovsky and D.C. Baxter, J. Anal. At. Spectrom., 17 (2002) 1223. I. Rodushkin, M.D. Axelsson, D. Malinovsky and D.C. Baxter, J. Anal. At. Spectrom., 17 (2002) 1231. J.S. Becker, J. Anal. At. Spectrom., 17 (2002) 1172. A.M. Leach and G.M. Hieftje, Appl. Spectrosc., 56 (2002) 62. R.E. Russo, X.L. Mao, O.V. Borisov and H.C. Liu, in: R.A. Meyers (Ed.), Encyclopedia of Analytical Chemistry. Wiley, Chichester, 2000, p. 9485. V. Kanicky, V. Otruba and J.M. Mermet, Spectrochim. Acta B, 55 (2000) 575. D. Gunther, I. Horn and B. Hattendorf, Fresenius J. Anal. Chem., 368 (2000) 4. J. Kosler, H.P. Longerich and M.N. Tubrett, Anal. Bioanal. Chem., 374 (2002) 251. S.M. Eggins, L.P. Kinsley and J.M. Shelley, Appl. Surf. Sci., 129 (1998) 278.

607

R.E. Russo and D.P. Baldwin 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94

608

A.P. Leung, W.T. Chan, X.L. Mao and R.E. Russo, Anal. Chem., 70 (1998) 4709. W.T. Chan, A.P. Leung, X.L. Mao and R.E. Russo, Appl. Surf. Sci., 129 (1998) 269. H.F. Falk, B. Hattendorf, K. Krengelrothensee, N. Wieberneit and S.L. Dannen, Fresenius J. Anal. Chem., 362 (1998) 468. C. Pickardt, J.S. Becker and H.J. Dietze, Fresenius J. Anal. Chem., 368 (2002) 173. V.R. Bellotto and N. Miekeley, Fresenius J. Anal. Chem, 367 (2000) 635. V. Kanicky and J.M. Mermet, Fresenius J. Anal. Chem., 363 (1999) 294. O.V. Borisov, X.L. Mao, A. Fernandez, M. Caetano and R.E. Russo, Spectrochim. Acta B, 54 (1999) 1351. Z.X. Chen, J. Anal. At. Spectrom., 14 (1999) 1823. V. Kanicky, V. Otruba and J.M. Mermet, Appl. Spectrosc., 52 (1998) 638. S.A. Watmough, T.C. Hutchinson and R.D. Evans, Environ. Sci. Technol., 32 (1998) 2185. M.D. Norman, W.L. Grifﬁn, N.J. Pearson, M.O. Garcia and S.Y. O’Reilly, J. Anal. At. Spectrom., 13 (1998) 477. D. Gunther, H. Cousin, B. Magyar and I. Leopold, J. Anal. At. Spectrom., 12 (1997) 165. J. Nolte, F. Schefﬂer, S. Mann and M. Paul, J. Anal. At. Spectrom., 14 (1999) 597. M. Gagean and J.M. Mermet, Spectrochim. Acta B, 53 (1998) 581. V. Huxter, J. Hamier and E.D. Salin, J. Anal. At. Spectrom., 18 (2003) 71. I. Horn, R.L. Rudnick and F.M. McDonough, Chem. Geol., 164 (2000) 281. M. Bi, A.M. Ruiz, B.W. Smith and J.D. Winefordner, Appl. Spectrosc., 54 (2000) 639. H.F. Falk, B. Hattendorf, K. Krengelrothensee, N. Wieberneit and S.L. Dannen, Fresenius J. Anal. Chem., 362 (1998) 468. X.L. Mao and R.E. Russo, J. Anal. At. Spectrom., 12 (1997) 177. F. Boue-Bigne, B.J. Master, J.S. Crighton and L. Sharp, J. Anal. At. Spectrom., 14 (1999) 1665. D. Gunther, R. Frischknecht, H.J. Muschenborn and C.A. Heinrich, Fresenius J. Anal. Chem., 359 (1997) 390. M. Ohata, H. Yasuda, Y. Namai and N. Furuta, Anal. Sci., 18 (2002) 1105. C. Craig, K.E. Jarvis and L. Clarke, J. Anal. At. Spectrom., 15 (2000) 1001. P.R. Manson and A.J. Mank, J. Analytical Atomic Spectrometry, 16 (2001) 1381. D. Gunther, S.E. Jackson and H.P. Longerich, Spectrochim. Acta B, 54 (1999) 381. D. Figg, J.B. Cross and C. Brink, Appl. Surf. Sci., 127 (1998) 287. X.L. Mao, A.C. Ciocan and R.E. Russo, Appl. Spectrosc., 52 (1998) 913. D. Figg and M.S. Kahr, Appl. Spectrosc., 51 (1997) 1185. P.M. Outridge, W. Doherty and D.C. Gregoire, Spectrochim. Acta B, 52 (1997) 2093. J.J. Gonzalez, X.L. Mao, J. Roy, S.S. Mao and R.E. Russo, J. Anal. At. Spectrom., 17 (2002) 1108. R.E. Russo, X.L. Mao, O.V. Borisov and H.C. Liu, J. Anal. At. Spectrom., 15 (2000) 1115. D. Gunther and C.A. Heinrich, J. Anal. At. Spectrom., 14 (1999) 1369. T.E. Jeffries, S.E. Jackson and H.P. Longerich, J. Anal. At. Spectrom., 13 (1998) 935. M. Guillong and D. Gunther, J. Anal. At. Spectrom., 17 (2002) 831. J. Koch, I. Feldmann, N. Jakubowski and K. Niemax, Spectrochim. Acta B, 57 (2002) 975. R. Jaworski, E. Hoffmann and H. Stephanowitz, Int. J. Mass Spec., 219 (2002) 373. J.E. Carranza and D.W. Hahn, Anal. Chem., 74 (2002) 5450.

Laser ablation sampling 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109 110 111 112 113 114 115 116

S.H. Jeong, O.V. Borisov, J.H. Yoo, X.L. Mao and R.E. Russo, Anal. Chem., 71 (1999) 5123. H. Fink, U. Panne and R. Niessner, Anal. Chem., 74 (2002) 4334–4342. W.E. Halter, T. Pettke and C.A. Heinrich, Science, 296(5574) (2002) 1844–1846. J.R. Ballard, J.M. Palin, I.S. Williams, I.H. Campbell and A. Faunes, Geology, 29(5) (2001) 383–386. J.C. Wataha, N.L. O’Dell, B.B. Singh, M. Ghazi, G.M. Whitford and P.E. Lockwood, J. Biomed. Mat. Res., 58(5) (2001) 537– 544. S. Rauch, G.M. Morrison, M. Motelica-Heino, O.F.X. Donard and M. Muris, Environ. Sci. Technol., 34(15) (2000) 3119–3123. A.J. Campbell and M. Humayun, Anal. Chem., 71(5) (1999) 939–946. J.E. Otamendi, J.D. de la Rosa, A.E. Patino Douce and A. Castro, Geology, 30(2) (2002) 159– 162. K. Rai, F.-Y. Yueh, J.P. Singh and H. Zhang, Rev. Sci. Instrum., 73(10) (2002) 3589– 3599. D.S. Zamzow, D.P. Baldwin, S.J. Weeks, S.J. Bajic and A.P. D’Silva, Environ. Sci. Technol., 28(2) (1994) 352– 358. H. Zhang, F.-Y. Yueh and J.P. Singh, J. Air Waste Manage. Assoc., 51(5) (1995) 681– 687. L.M. Cabalin and J.J. Laserna, Anal. Chem., 73(6) (2001) 1120– 1125. D. Body and B.L. Chadwick, Rev. Sci. Instrum., 72(3) (2001) 1625–1629. L. Burgio, K. Melessanaki, M. Doulgeridis, R.J.H. Clark and D. Anglos, Spectrochim. Acta B, 56 (2001) 905– 913. C.F. Su, S. Feng, J.P. Singh, F.-Y. Yueh, J.T. Rigby III, D.L. Monts and R.L. Cook, Glass Technol., 41(1) (2000) 16– 21. S.G. Buckley, H.A. Johnsen, K.R. Hencken and D.W. Hahn, Waste Manage., 20 (2000) 455–462. E.D. Lancaster, K.L. McNesby, R.G. Daniel and A.W. Miziolek, Appl. Opt., 38(9) (1999) 1476– 1480. U. Panne, M. Clara, C. Haisch and R. Neissner, Spectrochim. Acta B, 53 (1998) 1957– 1968. see also Part II, 1969–1981. R.J. Watling, J. Anal. At. Spectrom., 13 (1998) 917– 926. R.J. Watling, H.K. Herbert, D. Delev and I.D. Abell, Spectrochim. Acta B, 49(2) (1994) 205–219. R.J. Watling, B.F. Lynch and D.D. Herring, J. Anal. At. Spectrom., 12 (1997) 195–203. T. Howe, J. Shkolnik and R. Thomas, Spectroscopy, 16(2) (2001) 54– 66.

609

This page intentionally left blank

Chapter 21

Flow injection techniques for sample pretreatment Zhao-Lun Fang

21.1

INTRODUCTION

21.1.1 General Flow injection (FI) methods are based on the reproducible timing of controlled dispersion processes in the liquid phase [1]. The term “injection”, originally understood as an operation similar to that of an intravenous injection in the pioneering works of Ruzicka and Hansen, has now evolved into a much broader sense, particularly when involving pretreatment procedures. This includes a multitude of different modes of operations, such as volume-based sample loop injection or loading, hydrodynamic injection, intermittent pump sample introduction, time-based sample loading, etc. [2]. In recent years, sequential injection (SI) has emerged as a subdiscipline of FI, undergoing fast development [3]. SI allows more reproducible manipulation of solutions under computer controlled conditions, and has contributed signiﬁcantly to enhanced performance of FI procedures, albeit usually with some sacriﬁce in throughput. However, the basic principles of reproducible timing and dispersion control have not changed under different modes of operation. In this chapter, features related to FI refer to the technique in general, i.e., including SI, unless mentioned otherwise. No attempt is made to present a comprehensive coverage of FI pretreatment techniques for trace analysis within the limited space of this chapter. Instead, the reader will be provided with information and those experiences considered being most relevant and useful for exploiting the technique for trace analysis. Detailed information on related techniques may be found in several monographs [2,4 –6].

Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

611

Z.-L. Fang

21.1.2 General features of ﬂow injection on-line sample pretreatment systems When compared to batchwise operations for sample pretreatment, FI methods generally demonstrate some extremely favorable features in their performance, including: † † † † † † †

signiﬁcantly higher throughput, with short operation times, typically in the few tens of seconds to few minutes range. lower sample consumption by one to two orders of magnitude, a feature particularly important for valuable samples, such as blood, or for large numbers of samples collected from distant sampling districts; considerable savings in reagent consumption, often with .90% reduction; improved reproducibility, typically in the 1–3% r.s.d. range; more readily automated operation; reduced risks of contamination owing to the closed and inert ﬂow processing systems used—a feature particularly important for trace analysis; Reduced laboratory bench space.

A “drawback” of FI pretreatment procedures, which is usually unacceptable in batch methods, is the incompleteness of mass transfer between phases, which in some cases may be less than 50%. The lack of equilibrium in physical and/or chemical processes is an inherent feature of all operations based on these principles. This does not deteriorate the precision or sensitivity of a properly designed and calibrated FI system, including those used for separation and preconcentration or other pretreatments. However, one ought to be careful in designing and optimizing an FI system for such purposes, since extreme deviations from equilibrium may cause considerable errors in the analysis of real samples when identical equilibrium conditions cannot be achieved during calibration. On the other hand, one may exploit the highly reproducible non-equilibrium conditions in FI operations for improving selectivity through kinetic discrimination. 21.1.3 Classiﬁcation of FI sample pretreatment systems In addition to sample dissolution, sample preparation schemes for trace analysis are generally composed of various separation procedures for removing interferences and/or preconcentrating the trace analytes. Sample dissolution constitutes an important part of sample preparation, however, the need for elevated temperatures and complete breakdown of the sample matrix pose challenges to the FI principle which are difﬁcult to overcome (see Section 21.6). Sometimes, sample digestion methods considered as FI procedures, albeit effective and using similar hardware, are better identiﬁed as continuous ﬂow procedures. Those procedures which best reﬂect the basic principles of FI have been chosen for discussion.

612

Flow injection techniques for sample pretreatment TABLE 21.1 Phase separation characteristics Phase transfer

Separation mechanism

liquid– liquid

Solvent extraction Dialysis Liquid membrane extraction

liquid– gas

Gas diffusion Vapor generation

liquid– solid

Ion-exchange Adsorption Sorbent extraction Precipitation Coprecipitation

FI techniques for sample pretreatment in trace analysis usually involve online mass transfer of the analyte from one phase to another. The characteristics of the methods may be most conveniently identiﬁed and classiﬁed according to the separation mechanism or type of interface across which mass transfer takes place. The frequently used mechanisms are summarized in Table 21.1. FI is also a powerful technique for achieving sample dilution during sample preparations, however, this is of limited importance in trace analysis, as are methods based on dialysis, and precipitation; those readers who are interested in these may consult other sources where the topics have been fully elaborated [2,5,6]. Liquid membrane extraction is a powerful technique for sample pretreatment that is also employed in FI systems [7]. Since the topic is separately dealt with in Chapter 18, it will not be treated here. 21.1.4 Principles and general guidelines for the development of FI systems As mentioned earlier, FI systems are based on the principle of reproducible manipulation of solutions, both in terms of time (timing) and space (dispersion) under non-equilibrated conditions. Therefore, the precise timing of events, as well as stringent control of dispersion, which are often not important or critical in batch operations based on physical and/or chemical equilibrium, become important and key factors for the design and operation of a successful FI system. Some general guidelines to follow are listed below, particularly related to the development of FI pretreatment systems: (a) Although FI or SI systems may be operated manually (including the go–stop intervals of pumps, turning of valves, etc.), these cannot be

613

Z.-L. Fang

recommended for routine practice or for achieving optimum performance, owing to difﬁculties in precise timing. A computer-controlled system for automated programming of events, using software such as LabView, is deﬁnitely required for FI sample pretreatment systems, where operations are often more complicated than simple serial assays. (b) Hitherto, peristaltic pumps have been used as a principal means for solution delivery in FI systems; however, they are not the best choice for achieving precise timing, particularly in pretreatment systems, where high pressures develop within the ﬂow manifold (e.g., with on-line sorption column connected). Pulsation, pump tube aging, as well as factors associated with proper usage of such pumps [2] are the main factors affecting the reproducibility of timing. Stepper motor driven syringe pumps and low pressure (30– 60 bar) reciprocating piston pumps often used by Japanese workers usually provide better performance. (c) Although dispersion in FI systems may be exploited either for dilution or for preconcentration purposes, the former is seldom used in FI pretreatment systems for trace analysis. Thus, techniques for limiting dispersion (dilution) are of great concern. The most relevant rules for achieving such goals are summarized as follows: † Dispersion of a deﬁned ﬂuid zone into neighboring zones in a ﬁxed conduit (usually with 0.5 –1 mm i.d.) increases with the length of transport, particularly when the volume is in the 10 –102 ml range, but is relatively independent of the ﬂow rate and transport time. † Dispersion increases with internal diameter of the conduits. (d) Although a reasonable degree of equilibration is required for achieving sufﬁcient sensitivity and reproducibility in FI systems, steady state conditions are not required, and even deleterious, for achieving optimum performance. This feature can be fully exploited for achieving high throughput in FI pretreatment systems. 21.1.5 Practical hints for manipulation of FI equipment Some practical hints are listed here, based on principles cited in Section 21.1.4 and on the author’s personal experience: † † †

In order to reduce dispersion, all conduits should be kept as short as possible. Although i.d.s of 0.5 –1.0 mm are often used for preventing excessive backpressure, occasionally diameters as small as 0.25 mm are used to reduce dispersion when long transport lines are unavoidable. Redundant dead volumes along the conduits cause cumulative increases in dispersion that may seriously deteriorate the performance of, for example, an on-line preconcentration. Such voids are often produced by carelessly push-ﬁtted connections, loosely packed columns, and unsatisfactory valves.

614

Flow injection techniques for sample pretreatment

† †

† †

†

†

†

It is recommended that all components within a ﬂow line be inspected for unnecessary voids before running an FI system. Mixing of two solutions in neighboring ﬂuid zones may be achieved by mutual dispersion during transport. However, efﬁcient mixing is best achieved by merging two streams. Dispersion of a ﬂuid zone in a long transport line can be mostly avoided by introducing a small air segment at both the ends of the zone. However, such measures are limited to cases where detection systems are not affected by the air segments (e.g., in AAS applications) Tying the conduit into loosely interlaced knots (see Section 21.3.2.2) can signiﬁcantly reduce dispersion of a ﬂuid zone in a long transport line. Spatial position of conduits affects dispersion of ﬂuid zones through the creation of secondary ﬂows. Therefore, loosely suspended conduits should always be avoided. Conduit tubing should be ﬁxed to a rigid support, and long conduits used in a short distance should be securely coiled on the support. Connections along a ﬂow line should always be kept to a minimum. A single long conduit should substitute a long transport line with several connections, often formed because of frequent modiﬁcations during method development. Every connection, even properly made with zero dead volume between the ends, forms an extra site for potential leakages, and a multitude of connections can become a nuisance in routine practice. In general, push-ﬁtted connections using straight tubes, though convenient, are not sufﬁciently reliable, particularly in FI pretreatment systems where relatively large back pressures are developed within the system. These should be replaced by threaded ﬁtting connectors, or at least slip-proof pushﬁt connectors, whenever possible. Using a small piece of sandpaper wrapped around the tubings to increase friction may facilitate push ﬁtting of connecting tubes. Air bubbles formed spontaneously within the ﬂow lines affect the ﬂow-rate (and therefore, the precise timing of an FI system) as well as the dispersion proﬁle. Air bubbles entering a ﬂow-through detector may also cause serious interferences in the output signal (e.g. in spectrophotometry). Therefore, it is advisable to always de-gas all solutions before their introduction into the ﬂow system.

21.2

FI LIQUID–LIQUID EXTRACTION SYSTEMS

21.2.1 Introduction 21.2.1.1 General Liquid –liquid extraction is one of the most widely employed techniques in sample pretreatment for trace analysis. However, the demerits of the technique are also well known, i.e., the procedures, even automated, are usually rather tedious; and, particularly in trace analysis, susceptible to contaminations from

615

Z.-L. Fang

labware and laboratory environment; and fumes from organic solvents are often toxic. Such shortcomings may be largely avoided by employing FI techniques. FI liquid –liquid extraction systems are easily automated, and the closed extraction system signiﬁcantly reduces contamination as well as release of solvent vapors. Karlberg and Thelander [8], and Bergamin et al. [9] independently reported on FI liquid –liquid extraction systems in 1978. Since then, the technique has undergone considerable reﬁnement to improve its efﬁciency, reliability and robustness. Some typical FI liquid –liquid extraction systems are chosen to illustrate the performance of such systems. Liquid – liquid extraction procedures are usually composed of three successive steps: (a) dispensing of aqueous sample and organic solvent in deﬁned volumetric ratios into an extraction vessel; (b) creating intimate contact between the two phases by agitation; and (c) separation of the two phases. In an FI liquid –liquid extraction system, these are, respectively, executed through the use of (a) a phase segmentor, in which alternating segments of two immiscible phases are introduced to form a single continuous ﬂow; (b) an extraction coil in which mass transfer from one phase to the other is effected; and (c) a phase separator where the segmented phases are separated into independent ones. Occasionally, in some systems, determinations are made within the organic phase without phase separation, in which case the last step can be omitted. 21.2.1.2 Theoretical aspects In conventional batchwise liquid –liquid extractions, maximum exposure of the mass transfer interface between two immiscible phases is created by vigorous agitation, resulting in the formation of highly dispersed droplets of one phase in the other. Although in FI liquid –liquid extraction systems such conditions are not achievable, 80– 95% phase transfer can nevertheless be achieved in extraction coils, usually in less than a minute. The high efﬁciency of the mass transfer was explained by Nord and Karlberg [10] with the aid of an ingenious photographic technique. They provided evidence that a 0.01–0.05 mm thin ﬁlm was formed on the tube walls of the extraction coil by the phase that wetted the tube material (Fig. 21.1). When hydrophobic PTFE tubes are used, organic solvent ﬁlms are formed on the tube walls, while aqueous ﬁlms are formed on the hydrophilic walls of glass tubes. The relatively large ﬁlm area is responsible for the high efﬁciency of phase transfer in liquid –liquid extraction systems. Nord and Karlberg [10] derived an equation relating the ﬁlm thickness (df ; cm) to the inner diameter of the tube (R; cm), the viscosity (h; poise) of the ﬁlm forming phase, the ﬂow-rate (u; cm s21) and the surface tension (t; dyne cm22): df ¼ const £ Rðuh=tÞK where K is an empirical constant equal to 1/2 or 2/3.

616

ð21:1Þ

Flow injection techniques for sample pretreatment

Fig. 21.1. Schematic diagram showing ﬁlm formation in a liquid–liquid extraction coil tube. (a) static conditions; (b) ﬂow conditions. AQ, aqueous phase; OR, organic phase; F, organic ﬁlm [10].

From Eq. (21.1), it follows that the ﬁlm thickness increases at higher ﬂowrates, and disappears when the ﬂow stops; increases with an increase in viscosity of the ﬁlm forming phase and a decrease in the interfacial surface tension. Thinner ﬁlms are formed with solvents having lower viscosity and higher surface tension. Although ﬁlm thickness does not contribute signiﬁcantly to the efﬁciency of phase transfer, thicker ﬁlms are less stable, and tend to develop into small segments, resulting in irregular segmentation, which may deteriorate precision. 21.2.2 Apparatus for FI liquid– liquid extraction The various components of FI liquid –liquid extraction systems were stipulated in Section 21.2.1.2. These will be detailed in this section, both separately and as a functional component of a complete system. 21.2.2.1 Phase segmentors Phase segmentors are used to uniformly segment two immiscible phases in a ﬂowing stream. Regularity of the segmentation is important for improving precision and the efﬁciency of the phase separator. Phase segmentor designs can be as simple as a T- or Y-conﬁgured tube where the two phases enter from two branches and leave from the third as a segmented ﬂow. Further sophistication in design serves the purpose of controlling the length and regularity of the segments. Phase segmentors with a T-design, shown schematically in Fig. 21.2a, were the ﬁrst to be used for phase segmentation in FI liquid – liquid extraction systems [8]. The segmentor consisted of a glass capillary inlet for the aqueous phase and a glass tube outlet for the segmented stream. The organic phase was introduced through a platinum capillary, which merged with the inlet and outlet tubes at right angles. The inserted length of a PTFE tube within the outlet tube was adjusted to vary the segment size.

617

Z.-L. Fang

Fig. 21.2. Schematic diagram of (a) merging-tube segmentor; (b) coaxial falling-drop segmentor; OR, organic phase; AQ, aqueous phase; SP, segmented phase; Pt, platinum capillary [8].

However, such segmentors are relatively sensitive to ﬂow-rate variations, with organic phase segments decreasing at higher ﬂow-rates of the aqueous phase. Excessively small segments tend to rejoin later, forming irregular larger segments in the extraction coil. The coaxial segmentor design (originally named “falling drop” design) [11,12], introduced later, surmounted the principal shortcomings of mergingtube segmentors, and should be particularly useful when large phase ratios with high aqueous ﬂows are required for producing large preconcentration effects. This segmentor is based on droplet formation of one phase within the other, producing more reproducible and controllable segmentation than the merging-tube designs. Segmentation behavior is largely predictable, and segment lengths can be varied over a wide range of 2–50 mm for the organic phase and 3–300 mm for the aqueous phase. The segmentor is shown schematically in Fig. 21.2b, and is composed of a small Perspex or glass

618

Flow injection techniques for sample pretreatment

chamber with an inlet and a conical outlet, and a glass capillary, which is inserted into the conical part of the chamber. The organic solvent is introduced from the capillary (0.05 –0.35 mm i.d.) forming droplets that are released into the outﬂow channel of the chamber. The surrounding aqueous stream transports the droplets to the outlet, where phase segmentation occurs. Among different materials used for the construction of the segmentor chamber, all-glass chambers produced the best reproducibility of segment lengths over a wide range of ﬂow-rates for both phases. The organic solvent segment lengths are practically unaffected by the aqueous phase ﬂow-rate within a wide range of 0.25–10.0 ml min21. This implies that the aq/org phase ratio may be increased linearly with increases in aqueous phase ﬂowrate when the organic phase ﬂow-rate is kept constant. Despite the obvious advantages of the coaxial segmentor, which were later validated by the authors’ own experiences, implementation of the design in later studies of liquid –liquid extraction has been surprisingly few. This is probably due to the extra work involved in fabricating such a device, and/or one may be less demanding on the precision of trace measurements, being content with using a simple T-tube as a segmentor. However, the design offers not only improvements in precision, but also allows smoother operation, with fewer adjustments, and therefore deserves much broader acceptance.

21.2.2.2 Liquid– liquid extraction coil In an FI liquid – liquid extraction system, mass transfer of the analyte or interferents between two phases takes place in extraction coils. For transfer of analyte from an aqueous sample into an organic solvent, these are simply 0.5 – 1.0-mm i.d. PTFE tubes, which are coiled into a helix, but tubes of hydrophilic materials (e.g. glass) are required for reversed transfers. The dead volume of the coil and the ﬂow-rate of the segmented stream determine the extraction time. Provided the extraction system is chemically well optimized, and sufﬁcient extraction time is allowed, the phase transfer can be as complete as 80–99% in less than a minute.

21.2.2.3 Phase separators Phase separators are used to separate two immiscible segmented phases following extraction, and may be classiﬁed according to the mechanisms of phase separation. Gravity separators are based on differences in densities of the immiscible phases; membrane phase separators are based on differences of the two phases in afﬁnity to a porous separation membrane; whereas column separators, which are used exclusively for cloud point extraction, are based on ﬁltration of larger agglomerates of the surfactant-rich phase.

619

Z.-L. Fang

Gravity separators An earlier design of the gravity separator, produced from a glass T-tube, is shown in Fig. 21.3 [8]. The segmented ﬂow enters the separator from the horizontal side arm, and the separated phases ﬂow out in opposite directions through the two vertical arms, with the phase of higher density ﬂowing downwards. A PTFE strip is inserted into the segmented ﬂow inlet arm of the separator, extending into the organic phase outlet arm, acting as a guide to facilitate its smooth outﬂow. Although such separators are easy to produce, considerable care and experience are required for achieving good separation, particularly when two phases with similar densities are involved. We designed a more robust version of a gravity phase separator [13] which, following several modiﬁcations, has shown satisfactory performance in subsequent applications [14,15]. The basic structure of the separator shown schematically in Fig. 21.4 is suitable for the separation of an organic phase, with lower density than the aqueous phase. The separator is composed of two half blocks combined to form a conical cavity separation chamber. The upper block is produced from PTFE, containing the conical recess and outﬂow channel, and the lower block is of stainless steel, furnished with an inlet and outlet for the segmented and waste ﬂows, respectively. With this design, phase separation is aided by the difference in wetting properties of the two blocks in addition to differences in density.

Fig. 21.3. T-tube gravity phase separator for liquid–liquid extraction. 1, PTFE tube; 2, glass T-tube; 3, PTFE strip; SP, segmented phase; OR, organic phase; W, waste for aqueous phase containing residual organic phase [8].

620

Flow injection techniques for sample pretreatment

Fig. 21.4. Schematic diagram of a PTFE-stainless-steel gravity phase separator with a conical cavity. CC, conical cavity; SP, segmented phase; ORG, organic phase; AQ, aqueous phase; F, threaded ﬁttings [13].

Membrane separators Membrane separators were used most frequently for FI liquid – liquid extraction since this type of separator was ﬁrst described by Kawase et al. [16] in 1979. The separators were usually composed of two separate organic solvent resistant plastic blocks, furnished with grooves or cavities, with a microporous membrane sandwiched in between. Occasionally, the membranes are backed by supporting screens to improve their durability. A separator design, used successfully in the author’s laboratory [2,17], is shown in Fig. 21.5. The separator was produced from PTFE. The groove for the segmented ﬂow, connected to an inlet and outlet, was fabricated on the lower block; and a

Fig. 21.5. Schematic diagram of a sandwich-type membrane phase separator for liquid– liquid extraction. A, B, PTFE blocks; M, microporous PTFE membrane; G, slanted groove for segmented phase; P, metal plate for fastening A, B blocks by four screws (not shown) [17].

621

Z.-L. Fang

corresponding groove and outlet for the separated organic phase were produced in the upper block. The bottom of the groove for the segmented ﬂow was slightly slanted to create an impinging angle. The microporous membrane (18-mm2 usable area) was sandwiched between the two blocks, which were securely joined with two metal plates screwed tightly together. An impedance coil was used with the separator to adjust the pressure drop across the membrane. Various factors affect the phase separation efﬁciency of sandwich-type membrane separators, including membrane area and porosity; volume and structure of the mini-chambers housing the membrane and the pressure drop across the membrane, etc. Therefore, “ﬁne-tuning” of the separator is usually required for achieving optimum performance. This is further complicated by the relatively short lifetime of often a few working hours of the membranes, since re-tuning of the separator might be required following each membrane renewal, and re-calibration is indispensable. Despite the relatively large number of publications on the use of membrane separators, their reported multiple advantages, and the marketing of an early commercial version [18], these separators have not found broad acceptance in routine practice. One may ﬁnd an explanation to this situation from the limitations discussed above.

Column separator A packed column has been used as a phase separator in liquid –liquid extraction involving cloud point extraction. The cloud point phenomenon occurs in aqueous solutions of certain surfactants above the critical micelle concentration by an increase in temperature or by the introduction of saltingout agents. Under such conditions, the aqueous surfactant solution suddenly forms two phases and becomes turbid, owing to a decrease in the solubility of the surfactant in water [19]. Procedures based on this phenomenon provide an attractive alternative to sample pretreatment approaches with liquid –liquid extraction using organic solvents, and recently have been successfully adapted for application in an FI system [20]. The separator was simply a 3-mm i.d. glass tube with an effective length of 1.3 cm packed with a suitable ﬁltering material, such as cotton, glass wool or nylon ﬁbers. Following salt-induced aggregation of the surfactant-rich phase, larger size surfactant aggregates were entrapped within the column, while smaller sized components within the aqueous medium, including the salt, surfactant monomer, smaller sized surfactant aggregates and unretained analytes, passed through the column. The retained aggregates were then eluted using a suitable solvent. This separator is more robust and easier to operate than the previously described ones. However, it is limited to use with cloud point extraction, and the separation efﬁciency is expected to be inferior to solvent extraction systems, owing to losses during collection and dilution effects from the eluent.

622

Flow injection techniques for sample pretreatment

21.2.3 Guidelines for the development of FI liquid – liquid extraction systems Knowledge of some technical details is required for the design and smooth operation of liquid –liquid extraction systems. Some general guidelines are given here, many based on the author’s own experiences. Propulsion of organic solvents is inevitable in liquid –liquid extraction applications. However, the often-used peristaltic pumps are not suitable for such tasks, owing to the use of plastic pump tubes that are non-resistant to frequently used organic solvents. The displacement bottle, used in many publications in combination with a peristaltic pump, is helpful in partly overcoming this obstacle [2]. However, smooth operation of the device requires considerable skill and experience, and this approach cannot be recommended for routine applications. It is strongly advisable to acquire a peristaltic pump equipped with PTFE pump tubes, or better still, to use a solvent resistant lowpressure reciprocating piston pump. We have successfully used PTFE tubes with 0.25–0.5 mm wall thickness and 0.5 –0.7 mm i.d. as pump tubing with a number of conventional peristaltic pumps, achieving good performance at low ﬂow-rates of below 0.5 –ml min21 [13]. The tubes were loosely knotted at positions corresponding to the collars of normal pump tubes to prevent their sliding in the compression block during operation. Commercialized peristaltic pumps equipped with PTFE pump tubes are now also available. In order to ensure a “clean” organic phase for ﬁnal measurements in a detector, the aqueous phase should be drawn-off at a slightly higher ﬂow-rate than its inﬂow rate from the outﬂow arm of the separator. By doing so, a small fraction of the organic phase is forced to leave the separator from the aqueous outﬂow arm. This “sacriﬁce” reduces the chances of entraining aqueous phase into the separated organic fraction, which is further transported to the detector, and makes the system more robust. Entrainment of aqueous phase into a photometric ﬂow cell may cause contamination that can only be removed by washing the cell with ethanol. Careful balancing of inﬂow and outﬂow rates is NOT recommended. Membrane separators generally demonstrate better performance compared to gravity separators in terms of separation efﬁciency. However, their limitations discussed in Section 21.2.2.3 should be taken into serious consideration in routine applications, in which case gravity separators might be a better choice for maintaining stable operation over extended periods, albeit with reduced separation efﬁciencies. With membrane separators, PTFE membranes of about 0.1 mm thick having pore sizes of approximately 1.0 mm are used most frequently. Larger membrane pore sizes facilitate mass transfer, but with simultaneous lowering of water intrusion pressure, the optimization of pressure drop across the membranes becomes more critical.

623

Z.-L. Fang

Fig. 21.6. Schematic diagram of a typical FI manifold for liquid– liquid extraction spectrophotometry. P, pump; C, carrier; R, reagent; S, sample; SG, phase segmentor; EC, extraction coil; SP, phase separator; RT, restrictor or impedance coil; D, detector; W, waste [2].

21.2.4 Typical manifolds for FI liquid– liquid extraction Typical FI manifolds employed for on-line liquid – liquid extraction with spectrophotometric and AAS detection systems are shown in Fig. 21.6–21.8. These manifolds can be readily adapted for sample pretreatment in an off-line mode for various systems (GC, GC –MS, HPLC, LC –MS, etc.) by collecting the separated phase in small vessels. The differences to be noted in the various designs are as follows: (a) Manifolds shown in Fig. 21.6a and b [2] are suitable for spectrophotometric systems equipped with a ﬂow-cell, where the separated phase containing the analyte species is transported directly through the ﬂow-cell to produce a transient signal. The system in a is equipped with a membrane separator, and that in b, with a gravity separator, both using merging tube segmentors.

Fig. 21.7. Schematic diagram of an FI manifold for liquid– liquid extraction FAAS. P, pump; RG, reagent; S, aqueous sample; SG, phase segmentor; E, extraction coil; PS, membrane phase separator; R, restrictor or impedance coil; W, waste; V, injector; L, sample loop; AAS, ﬂame atomic absorption spectrometer [2].

624

Flow injection techniques for sample pretreatment

Fig. 21.8. FI manifold and operation sequence for on-line liquid–liquid extraction in ETAAS system [13]. Top: sample loading and extraction; bottom: concentrate delivery. P1, P2; peristaltic pumps; V, multifunctional valve; SG, phase segmentor; EC, extraction coil; PS gravity phase separator; SC, concentrate storage conduit; GT, graphite tube; W, waste; A, air-ﬂow entrance.

(b) For the system with a gravity separator, the phase passing through the detector is pumped out at a slightly lower (10– 15% lower, depending on its solubility in the other phase) ﬂow-rate than its inﬂow. For the system with membrane separator, an impedance coil is connected to the segmented phase outﬂow downstream of the separator to increase the pressure difference across the membrane. (c) The above systems are not suitable for detectors where the separated concentrate phase outﬂows cannot be controlled directly, such as in atomic spectrometry. Although, in principle, pumping out the residual segmented

625

Z.-L. Fang

phase is also possible, such practice is not recommended because of the large phase ratios often adopted in trace analysis. Small ﬂuctuations in the segmented phase outﬂow may transform to relatively large ﬂuctuations in the concentrate outﬂow, which are much lower, affecting both the separation efﬁciency and precision. (d) In order to achieve high enrichment factors without consuming excessive sample, the extractant ﬂow-rates are usually maintained at a low level of 0.1 –1-ml min21. Such low ﬂow-rates may not be suitable for direct introduction to systems where required sample aspiration rates are signiﬁcantly higher, such as for ﬂame-AAS. The system in Fig. 21.7 circumvents this problem using a semi-ofﬂine mode by collecting the separated concentrate in a sample loop before injection into the detection system under an optimized ﬂow-rate, using airﬂow as carrier. (e) The system in Fig. 21.8 is designed for liquid –liquid extraction combined with an ET-AAS system [13], also using a ﬂow of air to introduce a deﬁned volume of the concentrate into the electrothermal atomizer. The difference in this design from the previous one is that a sample loop was not used. 21.3

FI SOLID PHASE EXTRACTION SYSTEMS

21.3.1 Introduction Methods based on solid phase extraction form the largest class of FI on-line sample pretreatment systems in trace analysis, mostly coupled to atomic spectrometric detection. The reason for such popularity lies in the relative ease of implementing such systems, as well as the signiﬁcant improvement in sensitivity and selectivity. The latter is achievable often without signiﬁcantly affecting the sample throughput of normal atomic spectrometric systems (i.e. without pretreatment procedures). Reported efﬁciencies for preconcentration range from 10-fold to as much as 100-fold per minute, including the time for sample change and on-line detection, often with less than 100 ml sample consumed for each unit-fold preconcentration. However, careful design of the system is required for achieving such performance, and some guidelines are given in this section for setting up an efﬁcient system. Based on differences in the ﬂow manifolds employed, FI solid phase extraction systems are classiﬁed in this section as those employing packed columns and those using knotted reactors (KR). Based on differences in the mode of sample loading on the sorption medium, they are further classiﬁed as time-based or volume-based sampling. Almost all FI on-line solid phase extraction systems involve two stages of operation, i.e. sample loading and elution. Loading involves the retention of the analyte or interferent species on a solid support (sorbent), separating them from the rest

626

Flow injection techniques for sample pretreatment

of the sample matrix; and elution involves the removal of the retained species from the sorption medium. For volume-based loading, a ﬁxed volume of sample, pre-ﬁlled into a sample loop of an injector valve, is pumped through the sorption medium, usually transported by a suitable carrier stream. For timebased loading, samples are passed through the sorption medium at constant ﬂow-rate for a deﬁned period. The elution stage is usually performed in timebased mode, but occasionally, the eluent is delivered in the volume-based mode from a sample loop, in cases such as when only a single channel pump is available. 21.3.2 Sorption media for FI solid phase extraction 21.3.2.1 Packed microcolumns In FI solid phase extraction systems, microcolumns of ten to a few hundred microliters capacity packed with various sorbents are used most frequently as the sorption medium. Most columns are produced from 2–4 mm i.d. glass or plastic tubes of uniform diameter, usually no more than 2– 3 cm long. However, conical shaped columns were found to give better performance when samples were introduced from the thin end during loading, and later eluted from the same end into the detection system. Conical columns of only 9 –15 ml capacity were used in systems for ET-AAS, where concentrate volumes are limited by the graphite furnace capacity [21,22]. An important factor affecting the performance of FI on-line solid phase extraction systems for trace analysis is the property of the column packing. Compared to packing material used in batch methods, the on-line approach is more demanding on properties summarized below. † † † †

Physical and chemical stability required for at least 500 cycles of operation, during which no deterioration in performance should be noticed. Negligible swelling and shrinking during a cycle of operation. These cause problems by leading to either over-tightly-packed columns or void volumes in a column during different stages of operation. Kinetic properties should allow easy retention and fast elution of sorbed analyte by an appropriate eluent. Higher capacity for retention of analytes per unit volume.

In order to provide sufﬁcient selectivity for trace analysis, the most often used packing materials are ion-exchangers having chelating functional groups, and non-polar or medium polar sorbents that retain a large range of hydrophobic metal complexes. In the latter case, the selectivity is enhanced during the complexation, since non-complexed ions freely pass through the column. Chelex 100 is a chelating ion-exchanger used frequently in earlier studies, but has gradually lost popularity owing to its strong swelling and shrinking properties. The CPG-8Q chelating ion-exchanger, with quinolin-8-ol functional groups azo-immobilized on porous glass has excellent mechanical

627

Z.-L. Fang

and kinetic properties. However, its exchange capacity is relatively low, and its usage is restricted to samples with relatively simple matrices. The Japanese product, Muromac A-1 chelating ion-exchanger is relatively free from the deﬁciencies cited above. The C18 sorbent with octadecyl functional group immobilized on a silica gel matrix is used most frequently to retain a large variety of organic metal complexes [2,23]. Ethanol and methanol are used most often as eluents. Interference from common matrix elements, such as calcium and magnesium, are much less pronounced than most chelating ion-exchangers. Other packing materials recommended by various workers and reviewed by this author [2] include activated carbon, the XAD series polymeric adsorbent sorbents, and ﬁbrous ion-exchange materials. Anthemidis et al. [24] reported a microcolumn (30/4.6 mm i.d.) packed with PTFE turnings (0.1 mm mean width), which appears to be a hybrid between KRs (see Section 21.3.2.2) and packed columns. For the retention of Cu-APDC, with subsequent elution by IBMK into a F-AAS system, an impressive enhancement factor of 340 was reported with 1-min sample loading period, achieving a throughput of 40 h21. More recently, 100 mm PTFE beads were used for the column packing [25]. The authors reported a two-fold enhancement in enrichment factor in their application for preconcentration of APDCCr(IV) with ET-AAS detection, compared to a KR having an identical surface area.

21.3.2.2 Knotted reactors KR, produced from PTFE tubing tied into overlying knots, were ﬁrst introduced by Engelhardt and Neue [26] for minimizing dispersion in HPLC systems. Its capability for retaining precipitates was ﬁrst reported by Fang et al. [27] in the coprecipitation of trace metals using an HMDTC-Fe(II) carrier with F-AAS detection. Later, their properties of retaining organic metal complexes in solution were noted accidentally in one of our experiments, when using the reactor to transfer a sample solution containing trace metals complexed with APDC to a sorption column during sample loading [28]. The retention efﬁciency of the column was much lower than expected, and ultimately the loss was tracked to the upstream knotted reactor that, in fact, acted as a pre-column. With further optimization, it was found feasible to retain as much as 90% of complexed analytes with a KR. Another variant of KR sorption is to pre-coat the reactor walls with a suitable complexing agent before loading a non-complexed sample [29]. Owing to the use of an open tube reactor, sample loading ﬂow-rates much higher than those allowable using packed columns can be adopted. The beneﬁts and limitations of the KR reactor in trace analysis with atomic and mass spectrometric detectors have been reviewed recently by Yan and Jiang [30].

628

Flow injection techniques for sample pretreatment

21.3.3 Guidelines for the development of FI solid phase extraction systems Columns †

†

†

Sorption columns should be tightly and securely packed, sealed and connected to other components in the FI system. Void volumes in columns result in extra dispersion and reduction in the enrichment factor for an online preconcentration system. Although columns equipped both with push-ﬁtted or threaded ﬁttings (when properly implemented) can give satisfactory performance, threaded ﬁttings are strongly recommended as connectors for long term reliability and better tolerance of high sample loading rates. Note that a large percentage of poor performance results from column leakage. The use of a single multichannel or multifunctional injector valve is preferred over multiple valves in the design of on-line column pretreatment systems. The former, with columns or knotted reactors nested on the injector as a special kind of sample loop, is much more versatile, and operations are more easily automated and synchronized.

Sample loading †

†

†

†

Loading of sample onto a packed column should be performed by pumping the sample INTO the column through a pump. Sample loading by pump suction downstream of the column is generally not recommended because the reduced pressure often causes gas evolution and unstable ﬂow rates that deteriorate the performance. When organic metal complexes are formed on-line by merging the sample ﬂow with an organic complexing agent before loading onto a sorbent column, the often used PTFE transport lines upstream of the column should be substituted by polymer-coated silica capillaries, in order to avoid losses due to sorption in the transport conduits. When peristaltic pumps are used for sample loading, the pump tubes should be allowed to stabilize under working conditions by pumping water through the column for about 15 min. During this warm-up period, the ﬂow-rate is likely to drift and cause poor precision, particularly for time-based sample loading, Short pump tubes are recommended for sample delivery in order to minimize carryover. Discharge of sample waste from the column into the detection system should be avoided because the efﬂuent may exert deleterious effects on the ﬂow-through detector. These include effects of high concentration of dissolved solids on ﬂame AA or ICP spectrometer nebulizer–atomization systems, and effects of constituents in the sample efﬂuent, which poison an

629

Z.-L. Fang

ion-selective electrode or contaminate an optical ﬂow-cell window. Discharge of the efﬂuent to waste is easily accomplished using suitable valve and manifold designs (see Section 21.3.4.1).

Column washing †

Washing and/or pH equilibration of on-line columns before sample loading are not required, if samples are buffered during loading. The early stage of sample loading is used as a sequence for equilibration, with the same function as equilibrating with a buffer solution. This results in a simpler and more efﬁcient column preconcentration system. Washing columns with a rinsing solution after sample loading is only required when the residual sample matrix interferes with the ﬁnal determination in the eluate; e.g. in spectrophotometric or ET-AAS applications.

Elution †

†

Column elution ﬂow-rate is an important parameter in on-line column preconcentration for achieving optimum sensitivity. In most cases, the eluent ﬂow is connected directly to the detector. Therefore, this ﬂow-rate is particularly important for detection systems, which require a certain sample delivery rate such as F-AAS or ICP-OES. Usually a compromise has to be sought between the kinetic requirements of the sorbent and uptake rate of the detector. Reversal of the ﬂow direction through the column between the loading and elution stages is highly recommended to avoid the progressive tightening of packing material, which could, in turn, affect the ﬂow-rate. With strongly sorbed species, reversed ﬂow elution is also beneﬁcial in facilitating elution and improving the enrichment factor.

21.3.4 Typical manifolds for FI solid phase extraction 21.3.4.1 Systems for F-AAS, ICP-OES and ICP-MS Quite similar manifold designs may be employed for F-AAS, ICP-OES and ICP-MS to perform on-line solid phase extractions, using either a microcolumn or a KR, and usually may be adapted between each other with minor modiﬁcations in ﬂow parameters. Thus, the column or KR elution rate for ICP-OES and ICP-MS should be lower than those applied for F-AAS to conform to the different aspiration rates of the techniques. The ﬁrst published FI on-line solid phase extraction system was combined with F-AAS using a column packed with Chelex-100, and operated using

630

Flow injection techniques for sample pretreatment

volume-based sampling, with sample waste discharged into the spray chamber and with unidirectional ﬂow through the columns. The defects of this early design, discussed in Section 21.3.3, were overcome using more efﬁcient systems, such as that shown in Fig. 21.9, featuring time-based sample loading and a single multi-channel valve [31]. All the important points for achieving optimum performance mentioned in the guidelines are reﬂected in this design, including time-based sample loading on a conical microcolumn with efﬂuent discharge into waste, reversed ﬂow elution without washing sequence, and short transport conduits. A double pump system is used to decrease eluent consumption. An on-line column system can be readily transformed into one using a knotted reactor solid phase extraction system by simply substituting the column with the reactor. However, when the KR is employed for retaining precipitates, special designs are required to avoid coagulation and retention of precipitate outside the KR, particularly upstream of the injector valve in which the KR is nested. The latter circumstance not only causes analyte losses but also causes blockages within the valve channels. This can be avoided by employing the design shown in Fig. 21.10, where the precipitant is merged within the sampling loop of the injector valve [27]. Such a design could also be used for FI solid phase extraction of organic metal complexes that remain in solution, to reduce the risk of losses from sorption outside the sampling loop. 21.3.4.2 Systems for ET-AAS ET-AAS operations pose special requirements for the design and operation of FI sample pretreatment systems which are different from those for F-AAS. This arises because

Fig. 21.9. FI manifold of on-line solid phase extraction preconcentration system with column-in-loop design [31]: left, sample loading sequence; right, column elution sequence; P1, P2, peristaltic pumps; E, eluent; S, sample; B, buffer/reagent; C, packed microcolumn; V, multifunctional valve; W, waste.

631

Z.-L. Fang

Fig. 21.10. FI on-line sorption/precipitate collection preconcentration F-AAS system using KR [27]: (a) sample loading sequence; (b) elution/dissolution sequence. P1, P2, pumps; V, multifunctional valve; S, sample; R, complexing/precipitation reagent; E, elution/dissolution agent; KR, knotted reactor; W, waste.

† † †

The electrothermal atomizer cannot be operated in a continuous ﬂow mode as for F-AAS. The maximum sample volume that can be conveniently and reproducibly processed in an electrothermal atomizer is very low, e.g. only 50 –100 ml for the graphite furnace. ET-AAS is sensitive to matrix interferences; even with background correction, its tolerance to sample matrices is very low, unless matrix modiﬁcations are made.

Published designs on FI sample pretreatment systems for ET-AAS reﬂect efforts in dealing with these speciﬁc features of ET-AAS. Typical manifolds based on FI and SI principles are shown in Figs. 21.11 and 21.12. Important features in the manifold design and operation may be summarized as (a) Column dimensions are signiﬁcantly smaller than those used in F-AAS, 10–15 ml being the normal capacity of a minicolumn, and packings are chosen to exclude matrix constituents while trapping the trace analytes.

632

Flow injection techniques for sample pretreatment

Fig. 21.11. FI manifold for column-in-loop solid phase extraction preconcentration ET-AAS [21]. Three main operation sequences are shown out of ﬁve. (a) Sample loading; (b) sample elution; (c) delivery of collected eluate into graphite tube. P1, P2 peristaltic pumps; V, multifunctional valve; C, sorption column packed with C18; L, eluate (concentrate) collector; W, waste; GF, graphite tube.

633

Z.-L. Fang

Fig. 21.12. Schematic diagram of SI –ET-AAS system for on-line preconcentration with column-in-tip design [33]. Top: SP, syringe pump; GF, graphite furnace; V1, selector valve; V2, syringe valve; HC, holding coil. Bottom: sequence of zones in the holding coil before sample loading. R, APDC reagent; Et, ethanol eluent.

(b) FI operations are usually carried out in parallel with the furnace temperature program (i.e., one sample is processed in the FI system while another is processed in the atomizer), in a semi-on-line mode, for achieving higher sample throughput. (c) A column-rinsing step is included before elution to avoid introduction of residual sample matrix into the atomizer. (d) The systems are often programmed to achieve heart cutting or zone sampling of the eluted concentrate so that only the most concentrated fraction is introduced into the atomizer, while discarding the rest. (e) An airﬂow is used to deliver the sampled concentrate zone into the atomizer. The FI system in Fig. 7.11 is based on a column-in-loop design [32], with time-based sampling, similar to that used for F-AAS, with which ﬂow reversal during elution is readily achieved. Dispersion of the sampled concentrate in the relatively long transport line to the atomizer may be minimized by using airﬂow transport through small-bore capillaries of 0.2 –0.3 mm i.d. When the column is substituted by a KR, this system is transformed into one suitable for on-line preconcentration by coprecipitation [27]. The SI system shown in Fig. 7.12 [33] adopted a column-in-tip design, which was originally proposed by Porta et al. [34], in which the microcolumn is

634

Flow injection techniques for sample pretreatment

incorporated into the tip of the sampling probe of the autosampler. The advantage of the column-in-tip design is the direct dispensing of eluate (concentrate) into the atomizer, without using a transport line. Tightening of column packing is avoided by including a sequence of air suction through the column after the concentrate delivery. Volume based sampling is achieved most conveniently in SI systems. Zones of rinsing water, eluent (discarded fraction), eluent (sampled concentrate fraction), column rinsing agent, sample and reagent are sequentially aspirated into a holding coil, with each zone separated by a plug of air. The whole string of ﬂuid zones is then pumped through the column in reverse order. An intermittent stop was made before delivery of concentrate into the atomizer, to allow insertion of column into the atomizer. Huang’s group has published a series of applications for FI on-line preconcentration with ET-AAS detection, employing a simple column-in-tip system with a microcolumn of Muromac A-1 chelating resin mounted at the tip of an autosampler arm [35]. Deﬁned volumes of samples were loaded on the column by suction, which apparently reduced the imprecision in ﬂow-rate associated with suction loading, and satisfactory results were reported for a number of applications [35,36]. 21.4

FI VAPOR GENERATION SYSTEMS

21.4.1 Introduction Vapor generation (VG) methods, including hydride and cold VG, are established sub-disciplines of atomic absorption spectrometry. In this section, VG is treated as a sample pretreatment procedure for atomic spectrometry. The most important reason for the popularity of hydride generation (HG) AAS and cold vapor (CV) AAS is their high relative sensitivity. The relative detection limits are usually 3– 5 orders of magnitude lower than F-AAS methods, and often an order of magnitude lower than ET-AAS methods. However, batch operated systems suffer from demerits such as large sample and reagent consumption. The application of FI principles and techniques to HG-AAS pioneered by Astrom’s work [37] was reﬁned to overcome such demerits by signiﬁcantly reducing sample and reagent consumption, as well as improving selectivity, precision and throughput. FI –VG-AAS was, in fact, the ﬁrst FI-AAS system to be commercialized. 21.4.2 Gas – liquid separators for FI vapor generation Although gas diffusion separators with sandwich and tubular designs have been reported, they have not achieved wide popularity. Such separators involve transport of the gaseous phase between two neighboring channels separated by a porous membrane, which has relatively short lifetime, and are less robust than gas expansion separators. Readers interested in gas-diffusion separators

635

Z.-L. Fang

may consult other sources where detailed information is available. Only gas expansion separators will be treated in this section. In a gas-expansion separator, gas–liquid separation is achieved in a gas-expansion chamber of a few milliliters capacity. The reaction mixture is drawn to waste through an outlet from below the separator, with forced outﬂow, while the expanded gases are discharged from an upper outlet. The Vijan-type glass U-tube and W-tube gas–liquid separators used in earlier applications of FI VG methods [2] are currently mostly substituted by more efﬁcient detachable designs shown in Fig. 21.13a and b, produced from plastic materials [5]. The cylindrical-shaped separation chamber is furnished with an inlet for the gas –liquid mixture, positioned about halfway down the chamber. The mixture ﬂows down the wall of the chamber to the bottom, during which the evolved gas phase leaves the chamber from an outlet on top of the separator. The waste is drawn off from an outlet located in the lower part of the chamber. Packing the separator half-full with glass beads of approximately 3 mm diameter helps to reduce aerosol formation and foaming. As a further precaution to avoid aerosol entrainment into the nebulizer, sometimes a gaspermeable microporous membrane is inserted in the gas outlet of the separator. A three-part detachable design, later proposed by Cadore and Baccan [38], involved the use of a fritted glass disc at the bottom of a cylindrical separation chamber, through which the inert stripping-gas was introduced. This design was reported to improve the separation efﬁciency of BiH3 from the liquid phase, compared to U-tube separators. However, its use seems not to have been extended to other VG species. 21.4.3 Guidelines for development of FI vapor generation systems Despite the similarity in basic conﬁgurations of FI –HG-AAS systems based on gas-expansion separation, the performance of the system may vary extensively, depending on the design of the system components and on the operational parameters adopted. Important principles for the design of FI–HG-AAS systems were given by Fang [2]. Although the principles are related mainly to systems based on gas-expansion separation, most of the principles are valid for gas-diffusion separation systems and integrated systems as well. The important points are revised and summarized as below: (a) Minimization of dead volume of the gas–liquid separator is extremely important for obtaining optimum sensitivity; however, this should not be pursued by sacriﬁcing the effectiveness of separation. (b) A carrier gas (usually argon), introduced before or within the gas–liquid separator, is required to strip and transport the evolved hydride and hydrogen gases smoothly through the separator into the quartz atomizer. It is extremely important that the carrier gas maintains a steady ﬂow, unaffected by the relatively large pressure ﬂuctuations generated by the

636

Flow injection techniques for sample pretreatment

Fig. 21.13. Detachable expansion-type gas– liquid separators: (a) two-piece detachable separator with glass beads [5]; (b) two-piece detachable separator with microporous membrane (Perkin– Elmer); (c) three-piece detachable separator with fritted glass disc [38]; G– L gas–liquid mixture; D, to AA spectrometer; W, to waste; B, glass beads; O, o-ring; M, microporous membrane; F, fritted disc. Reproduced by permission of Royal Society of Chemistry.

637

Z.-L. Fang

(c)

(d)

(e)

(f) † † † † † † † †

discontinuous release of hydrogen gas segments into the gas–liquid separator. The gas-ﬂow regulation should be capable of maintaining a ﬂow as low as 150 ml min21 with backing pressures of over 0.3 MPa to ensure good precision. Stripping gases are found not necessary in VG-AAS employing vapor trapping in an electrothermal atomizer. Tubes with large inner diameters of about 1.0 –1.2 mm are recommended for producing reaction coils and transport conduits. The larger diameter does not contribute much to the dispersion owing to the large sample volumes and gas-segmentation of the ﬂow stream following reductant addition, but reduces the back pressure and pulsations induced by hydrogen generation. Forced withdrawal of reaction waste from the gas-expansion separator at a ﬂow-rate higher than the total liquid inﬂow rate by a factor of 2–3 (depending on the viscosity of the liquid phase) is strongly recommended to ensure smooth operation. The small loss in sensitivity by doing so is usually negligible. Balancing of the inﬂow and outﬂow should NOT be attempted, since this may easily result in carriage of waste solution into the quartz tube and leading to serious contamination of the atomizer cell. Short reaction coils of no more than 10 cm length (1 mm i.d. tubing) are recommended for suppressing interferences and for enhancing throughput. Typical parameters for FI–VG or SI –VG which may be adopted or used as a starting point for further optimization include the following [5]: Sample volume: 0.5 ml Acid carrier ﬂow-rate: 10–15 ml min21 Sodium tetrahydroborate reductant concentration: 0.1– 0.5% Reductant ﬂow-rate: 2 ml min21 Waste withdrawal ﬂow-rate: 20 –30 ml min21 Carrier gas ﬂow-rate: 100 ml min21 Reaction coil length (0.7 mm i.d.): 30 cm Quartz tube atomizer dimensions: 7 mm i.d., 170 mm long

21.4.4 Typical FI manifolds for VG-AAS 21.4.4.1 FI systems for VG-AAS Despite the signiﬁcant developments made in VG-AAS systems, very little has changed in the basic conﬁguration of FI–VG-AAS systems since their ﬁrst introduction. A typical system is shown in Fig. 21.14. After loading acidiﬁed samples into the sample loop of the injector, the sample is injected into an acid carrier stream and transported to merge the downstream with a ﬂow of reductant (usually tetrahydroborate) at a convergence point. The vapor phase is generated together with hydrogen from the reaction mixture, while being transported through a reaction coil. A stabilized ﬂow of argon or nitrogen stripping gas is introduced downstream of the coil. The gas– liquid mixture is

638

Flow injection techniques for sample pretreatment

Fig. 21.14. Schematic diagram of basic manifold for FI –VG-AAS system [5]: AAS, atomic absorption spectrometer with quartz tube atomizer; Ar, argon stripping gas; C, acid carrier; P, peristaltic pump; R, tetrahydroborate reductant; a, b, reaction coils; S, sample injector; SP, gas–liquid separator; W, waste.

transported to the gas –liquid separator where the separated gas phase is carried to the atomizer cell and the liquid phase is pumped out to waste. 21.4.4.2 FI systems for vapor trapping in graphite furnaces with ET-AAS detection The trapping of hydride-forming analytes in a preheated graphite furnace, through sequestration of the volatile hydrides, has been proposed by Sturgeon et al. as a batch procedure [39]. Usually, time-based sample introduction is adopted to increase the processed sample volume and improve the detection limit. A typical system is shown in Fig. 21.15, where the sample ﬂow timing is controlled by an injection valve, and a carrier stream is used to transport the sample into the reaction coil [40]. Another noticeable feature in this system is the absence of a carrier gas. Although argon carrier gas was used for transporting the generated vapor into a heated furnace atomizer, as in FI–VG-AAS systems using quartz atomizers by some workers, Tao and Fang [40] found this to be unnecessary and even detrimental. Hydrogen evolved in the VG reaction is usually sufﬁcient for transporting the hydrides to the furnace, whereas the use of carrier gases may reduce the sequestration efﬁciency by shortening the residence time of the vapor within the atomizer. 21.4.4.3 SI systems for VG-AAS In FI–VG-AAS systems, the reductant is always delivered continuously to merge with the sample/carrier ﬂow, irrespective of whether the sample or carrier zone is merged. A saving in reagent would be possible if the reagent pump is controlled to deliver reagent only when the sample zone passed the

639

Z.-L. Fang

Fig. 21.15. Schematic diagram of time-based FI sample deposition system by hydride sequestration for ET-AAS [40]: P, pump; V, valve; S, acidiﬁed sample; R, reductant; A, hydrochloric acid; W, waste; GLS, gas–liquid separator; GT, graphite tube. Adapted from reference [40] by the permission of Royal Society of Chemistry.

merging point. However, this is rarely practiced because of complications in programming and timing, owing to ﬂow-rate drifts of peristaltic pumps. Although such difﬁculties may be readily overcome by employing a SI system for VG-AAS, the system posed other difﬁculties if the acidiﬁed sample and reagent were to be brought into contact when transported through a holding coil. The gas evolution would seriously interfere with the ﬂow control. To avoid such circumstances, Ma et al. [41] devised a double syringe pump SI system shown in Fig. 21.16. The reagent was delivered for 3 s at 2-ml min21 (i.e. only

Fig. 21.16. Schematic diagram of a SI – VG-AAS system [41]: SP1, SP2, syringe pumps, V1, V2, 2-way valves; SV, multi-position selector valve; S, acidiﬁed sample; RC, reaction coil; P, peristaltic pump; GLS, gas–liquid separator; HC, holding coil; W, waste; Ar, stripping gas; AAS, atomizer.

640

Flow injection techniques for sample pretreatment

100 ml 0.5% NaBH4 consumed) during 9 s of sample/carrier delivery, and the reagent was reduced dramatically without affecting throughput, precision and sensitivity. A peristaltic pump was used to withdraw the waste from the gas expansion separator at double the total inﬂow of liquid phase (17-ml min21). The pump was also used to speed up sample change during the VG stage, by rinsing out the previous sample through a T-piece. 21.5

FI GAS DIFFUSION SYSTEMS

21.5.1 General The volatile species of an analyte in a ﬂow stream may be separated from interferents in an ill-deﬁned sample and transferred into a liquid acceptor stream with well-deﬁned composition by FI on-line gas diffusion. Reaction conditions for the gas–liquid separation and detection of the separated species may then be optimized independently. Under non-equilibrated conditions, the phase transfer rarely exceeds 30%, and may not be suitable as a pretreatment procedure for trace analysis. However, this can be compensated, to some extent, by applying preconcentration measures, and the approach has proved to be useful for enhancing the selectivity of trace determinations. The ﬁrst FI gas diffusion separation system was reported in 1979, by Baadenhuijsen and Seuren-Jacobs [42], using a sandwich type separator with a semi-permeable silicone rubber membrane. Most subsequent developments on gas diffusion separations followed a similar approach, but used PTFE microporous membranes. The analyte is usually transformed into a volatile species by means of a suitable acid-base chemical reaction before the separation; such as for the release of carbon dioxide, sulfur dioxide, hydrogen cyanide, hydrogen sulﬁde. Some analytes, e.g., ethanol, ozone, and chlorine dioxide, are sufﬁciently volatile to be separated from liquid phase under higher temperatures. The separated species is then absorbed by a liquid acceptor stream and transported, with or without a derivatization reaction, to the detector. When the separated gaseous analyte species reacts with a reagent to form a different species on the acceptor side, transfer of the gaseous analyte through the separation interface is enhanced. Thus the transfer of ammonia is enhanced by the use of an acidic acceptor. 21.5.2 Gas-diffusion separators Two main types of gas diffusion separators may be identiﬁed for use in FI systems, i.e., the sandwich type and the tubular type. However, the latter is of limited use, and only the sandwich design will be treated in this section. A typical structure of this type of gas-diffusion separator is shown in Fig. 21.17. The separator is composed of two half blocks on which are engraved channels that are mirror images of each other, with a porous membrane which separates

641

Z.-L. Fang

Fig. 21.17. Schematic diagram of an FI gas-diffusion separation system with sandwichtype separator: CR, carrier stream; R, reagent; S, sample injector; A, acceptor stream in and out; P, pump; W, waste [2].

the donor and acceptor streams sandwiched in between. Inlet and outlet ports, extending from the channels and furnished with connectors, are provided on each block. The dimensions of the channels, which may be either straight, meandering or spiral, can vary in the range 0.1 –1.0 mm deep, 1–3 mm wide, and 3–10 cm long. 21.5.3 Typical FI manifolds for gas-diffusion separation and preconcentration 21.5.3.1 Gas diffusion separation systems A typical FI manifold for gas diffusion separation is shown in Fig. 21.17 [42,43]. The samples are injected into a carrier stream and usually merged with a reagent to transform the analyte into a volatile species in the donor channel of the gas-diffusion separator. The gaseous analyte species diffuses through the membrane into the acceptor channel and is absorbed by the acceptor stream, which is delivered to a ﬂow-through detector. In this system the donor and acceptor streams ﬂow continuously usually at equal ﬂow-rates, typically in the range 0.5 –1.5 ml min21. Enrichment effects are thus not expected. 21.5.3.2 Gas diffusion preconcentration systems Preconcentration effects can be achieved by stopping the acceptor stream for a predetermined period while the sample in the donor stream is pumped continuously through the gas diffusion separator, at the expense of a larger sample volume. This was conveniently achieved using a time-based sampling gas-diffusion preconcentration FI system with a sandwich type separator nested in the sample loop reported by Zhu and Fang [44], and shown in Fig. 21.18. The system was used successfully for the determination of total cyanide. The samples were acidiﬁed before being transported through a heated reaction coil to liberate the hydrogen cyanide. The sample stream was transported through the donor channel of the separator while the basic acceptor stream was stopped for a deﬁned period by switching the loop out of the ﬂow.

642

Flow injection techniques for sample pretreatment

Fig. 21.18. FI manifold with gas-diffusion separator nested in sample loop of the injection valve, used for preconcentration of volatile species by time-based sampling (sample loading sequence) [44]. AS, autosampler; T, heater (optional); GDS, gas-diffusion separator; V, injection valve; R1, reagent for generation of volatile species; R2, acceptor reagent stream; R3, derivatization reagent (optional); D, detector; W, waste; a, valve position in sample injection sequence. Crossed circles in valve represent blocked channels.

During this preconcentration period the hydrogen cyanide penetrating the membrane was collected in the acceptor solution, and subsequently transported to the detection system. A 3.5-fold enrichment was achieved. A similar preconcentration system was used by Schulze et al. [45] for the determination of ammonia, with 10-fold sensitivity enhancement. 21.6

FI ON-LINE SAMPLE DIGESTION

21.6.1 Introduction Sample digestion or dissolution is often the most rate-limiting factor for sample throughput. However, digestion procedures often involve operations at elevated temperatures that pose special challenges in closed ﬂow systems such as FI. Microwave heating was expected to be best suited for such applications owing to its capability of fast heating of liquids ﬂowing within nonconductive plastic (e.g. PTFE) tubings. However, several problems, which are not always addressed in related publications, are often encountered even with microwave heating. The main points may be summarized as incomplete mineralization of the sample owing to the short reaction times, evolution of gases during digestion, large percentage of non-absorbed power, and high pressure build-up [46]. In order to overcome these difﬁculties, some on-line digestion systems, claimed as FI systems, were modiﬁed to the point that the basic principles of FI

643

Z.-L. Fang

no longer existed. Such systems may be better identiﬁed as batch procedures operated with pumps. Only the most relevant systems will be described in this section. 21.6.2 FI on-line sample digestion systems for AAS The earlier FI on-line sample digestion systems coupled to F-AAS detection were all applied to the on-line digestion of blood using microwave ovens. However, the digestion conditions were not strong enough for complete breakdown of the organic constituents in the sample to allow the use of simple aqueous standards, and it was necessary to use matrix-matched standards to obtain acceptable results. Burguera et al. [47] reported an on-line microwave digestion system for the F-AAS determination of zinc and copper in whole blood, with sample uptake directly from the vein of a patient’s forearm. The blood sample was pumped from the patient at 0.5 ml min21, sequentially merged with streams of EDTA anticoagulant and an acid mixture (1 mol l21 HNO3 þ 1 mol l21 HCl), and mixed in a 500-cm coil, before being delivered into the injector loop. The sampled digest mixture was transported by a carrier stream, through a 2-m digestion coil located in the microwave oven, using a second pump. A membrane gas-diffusion separation device, installed downstream of the oven, was used to de-gas the digest before introduction into the FAAS system. Aqueous standards with 70 g l21 human albumin and 7% glycerol were required for calibration, obviously owing to incomplete digestion. 21.6.3 FI digestion systems coupled to VG-AAS Tsalev et al. [46,48] developed an on-line digestion system for liquid samples coupled to FI –CV and HG-AAS systems, shown schematically in Fig. 21.19.

Fig. 21.19. FI manifold of on-line microwave digestion system for VG-AAS [46]: C, carrier; S, sample injection valve; P, pump; MWO, microwave oven; R, reductant; DC, dummyload coil; W, waste; GLS, gas– liquid separator. Adapted from Ref. [46] by permission of Royal Society of Chemistry.

644

Flow injection techniques for sample pretreatment

A 40–50 W focused-microwave digester was used with reduced irradiated space of about 56 cm3, to improve efﬁciency, and aqueous samples were heated to 80 –908C within 3–4 s at 10-ml min21 ﬂow-rate. The system was applied to the VG-AAS determination of Hg, As, Bi, Pb, and Sn in urine and environmental waters with sample throughputs of 13–30 h21, using oxidizing mixtures composed of bromide –bromate–acid or persulphate –acid-complexing agent. The high efﬁciency of the system was demonstrated by quantitative recoveries of eight different organic and inorganic mercury species in urine using aqueous standards prepared from Hg(NO3)2 [49]. The system was further modiﬁed by Welz et al. [50], accommodating a merging-zones sample and reagent introduction system to reduce acid consumption and installing a capillary ﬂow restrictor to increase the pressure during digestion. Guo and Baasner [51] described an efﬁcient on-line microwave digestion system for FI –CV-AAS determination of mercury in blood samples. Bromatebromide oxidizing reagent was added to 1:1 diluted blood samples before injection into a KR reaction coil installed in the microwave reactor. Permanganate was then merged with the cooled sample digest. Complete recoveries for four organomercury compounds were obtained, with a sample throughput of 45 h21 in the analysis of 1:10 diluted whole blood samples. 21.6.4 FI systems for digestion of solid samples in AAS Haswell and Barclay [52] developed an on-line microwave digestion system that allowed the analysis of slurried samples with organic matrices using simple aqueous standards. Slurried samples, containing 0.005–0.5% solid samples (, 180 mm) in 5% (v/v) HNO3, were injected into a water carrier, and transported through a 20-m long 0.8 i.d. PTFE reaction coil, installed in a 525 W microwave oven operated at 90% power. Subsequent cooling of the digest in a 5-m long PTFE loop immersed in an anti-freeze cooling bath completely removed the gas segments generated in the heating process. This allowed the introduction of the sample digest directly into F-AAS system without affecting the ﬂow pattern. Burguera and Burguera [53] described an FI microwave digestion system for the ET-AAS determination of lead in solid biological samples. Slurried solid samples or blood were merged with a HCl –HNO3 mixture and transported through a PTFE coil installed inside a microwave oven. Gases formed during the digestion were removed by a gas-diffusion separator, and the digest was cooled in a bath before injection into a sample collector tube. A deﬁned volume of the mineralized sample collected in the tube was then dispensed into the graphite tube by airﬂow. Good agreement with certiﬁed values was obtained for bovine muscle and liver, pig kidney and whole blood samples, but lower results were obtained for the botanical samples, indicating insufﬁcient mineralization conditions for these samples.

645

Z.-L. Fang

21.6.5 FI pretreatment systems with on-line photo-oxidation by UV irradiation FI systems offer a convenient platform for on-line photo-oxidation pretreatment. An FI –HG-AAS system has been developed by Atallah and Kalman [54] with on-line conversion of organoarsenicals into arsenate using a photo-reactor. The injected sample and carrier streams were merged with a potassium persulphate oxidant before being transported through a 5-m PTFE reaction coil wrapped around a mercury lamp. The sample was then acidiﬁed on-line and processed as in a typical FI– HG-AAS system with sodium tetrahydroborate reduction. Most other workers in later publications adopted the reactor design reported in this work. Tsalev et al. [55] reported an FI on-line UV photo-oxidation system for organoarsenic and organotin species with HG-AAS detection. Treatment with alkaline peroxydisulfate in a 10–15 m PTFE KR for .1.5 min achieved . 90% transformation of inorganic As(III) and six organoarsenic species to arsenate. The UV photo-oxidation with acidic peroxydisulfate at 95–1008C provided recoveries of . 80% for dimethyltin, trimethyltin, triethyltin, tripropyltin, triphenyltin, monobutyltin, dibutyltin and tributyltin, but only approximately 15% for tetrabutyltin. The throughput was 20 and 12 samples per hour with 10and 15-m knotted reactors (i.d. 0.5 mm), respectively. On-line photo-oxidation was also used successfully for the digestion of organic phosphorus compounds in waters, waste waters [56], and soil leachates [57], followed by spectrophotometric determination using the molybdenum blue reaction. Although on-line UV irradiation of water samples in a mixture of perchloric acid and peroxydisulfate gave complete recoveries for most organic phosphorus compounds tested, it was found to be insufﬁcient for the digestion of condensed phosphates. A combined digestion system was, therefore, developed with a coiled photo-reactor and thermal reactor (908C) connected in series, resulting in . 85% recovery for condensed phosphates [56]. Six-metre coils produced from 1-mm i.d. PTFE tubing were used for both reactors. The out-ﬂowing digest was loaded directly into a sample loop of an injector for the subsequent photometric determination. UV irradiation alone was found to be sufﬁcient for the determination of total phosphorus in soil leachates because of the relatively low concentration of condensed phosphates in such samples [57].

REFERENCES 1 2 3

646

J. Ruzicka and E.H. Hansen, Flow Injection Analysis, 2nd edn. Wiley, New York, 1988. Z.-L. Fang, Flow Injection Separation and Preconcentration. VCH Publishers, Weinheim, 1993. J. Ruzicka and E.H. Hansen, Anal. Chem., 72 (2000) 212A.

Flow injection techniques for sample pretreatment 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44

M. Valcarcel and M.D. Luque de Castro, Non-chromatographic Continuous Separation Techniques. Royal Society of Chemistry, Cambridge, 1991. Z.-L. Fang, Flow Injection Atomic Absorption Spectrometry. Wiley, Chichester, 1995. A. Sanz-Medel (Ed.), Flow Analysis with Atomic Spectrometric Detectors. Elsevier, Amsterdam, 1999. J.A. Jonsson and L. Mathiasson, Trends Anal. Chem., 18 (1999) 318, see also 325. B. Karlberg and S. Thelander, Anal. Chim. Acta, 98 (1978) 1. H. Bergamin F8, J.X. Medeiros, B.F. Reis and E.A.G. Zagatto, Anal. Chim. Acta., 101 (1978) 9. L. Nord and B. Karlberg, Anal. Chim. Acta, 164 (1984) 233. K. Backstrom and L.-G. Danielsson, Anal. Chim. Acta, 232 (1990) 301. V. Kuban, L.-G. Danielsson and F. Ingman, Anal. Chem., 62 (1990) 2026. G.-H. Tao and Z.-L. Fang, Spectrochim. Acta B, 50 (1995) 1747. Q. Fang, Y.-Q. Sun and Z.-L. Fang, Fresenius J. Anal. Chem., 364 (1999) 347. S.C. Nielsen, S. Sturup, H. Spliid and H. Hansen, Talanta, 49 (1999) 1027. J. Kawase, A. Nakae and M. Yamanaka, Anal. Chem., 51 (1979) 1640. Z.-L. Fang, Z.-H. Zhu, S.-C. Zhang, S.-K. Xu, L. Guo and L.-J. Sun, Anal. Chim. Acta, 214 (1988) 41. Y. Sahlestrom and B. Karlberg, Anal. Chim. Acta, 185 (1986) 259. B. Moreno Cordero, J.L. Perez Pavon, C. Garcia Pinto and M.E. Fernandez Laespada, Talanta, 40 (1993) 1703. Q. Fang, M. Du and C.W. Huie, Anal. Chem., 73 (2001) 3502. Z.-L. Fang, M. Sperling and B. Welz, J. Anal. At. Spectrom., 5 (1990) 639. B. Welz, M. Sperling and X.-J. Sun, Fresenius J. Anal. Chem., 346 (1993) 550. J. Ruzicka and A. Arndal, Anal. Chim. Acta, 216 (1989) 243. A.N. Anthemidis, G.A. Zachariadis and J.A. Stratis, Talanta, 54 (2001) 935. W. Som-Aum, S. Liawruangrath and E.H. Hansen, Anal. Chim. Acta, 463 (2002) 99. H. Engelhardt and U.D. Neue, Chromatographia, 15 (1982) 403. Z.-L. Fang, M. Sperling and B. Welz, J. Anal. At. Spectrom., 6 (1991) 301. Z.-L. Fang, S.-K. Xu, L.-P. Dong and W.-Q. Li, Talanta, 41 (1994) 2165. K. Benkhedda, H.G. Infante, E. Ivanova and F. Adams, J. Anal. At. Spectrom., 15 (2000) 429. X.-P. Yan and Y. Jiang, Trends Anal. Chem., 20 (2001) 552. Z.-L. Fang and B. Welz, J. Anal. At. Spectrom., 4 (1989) 543. Z.-L. Fang, M. Sperling and B. Welz, J. Anal. At. Spectrom., 5 (1990) 639. Z.-R. Xu, H.-Y. Pan, S.-K. Xu and Z.-L. Fang, Spectrochim. Acta B, 55 (2000) 213. V. Porta, O. Abollino, E. Mentasti and C. Sarzanini, J. Anal. At. Spectrom., 6 (1991) 119. Y.-H. Sung, Z.-S. Liu and S.-D. Huang, Spectrochim. Acta B, 52B (1997) 755– 764. H.-J. Chang, Y.-H. Sung and S.-D. Huang, Analyst, 124 (1999) 1695. O. Astrom, Anal. Chem., 54 (1982) 190. S. Cadore and N. Baccan, J. Anal. At. Spectrom., 12 (1997) 637. R.E. Sturgeon, S.N. Willie and S.S. Berman, J. Anal. At. Spectrom., 1 (1986) 115. G.-H. Tao and Z.-L. Fang, J. Anal. At. Spectrom., 8 (1993) 577. H.-B. Ma, S.-K. Xu, H.-Y. Zhou, S.-L. Wang and Z.-L. Fang, Guangpuxue yu Guangpu Fenxi, 20 (2000) 529. H. Baadenhuijsen and H.E.H. Seuren-Jacobs, Clin. Chem., 25 (1979) 443. S.-H. Fan, J.-X. Li and Z.-L. Fang, Fenxi Shiyanshi, 10(2) (1991) 39. Z.-H. Zhu and Z.-L. Fang, Anal. Chim. Acta, 198 (1987) 25.

647

Z.-L. Fang 45 46 47 48 49 50 51 52 53 54 55 56 57

648

G. Schulze, C.Y. Liu, M. Brodowski, O. Elsholz, W. Frenzel and J. Moller, Anal. Chim. Acta, 214 (1988) 121. D.L. Tsalev, M. Sperling and B. Welz, Analyst, 117 (1992) 1729. J.L. Burguera, M. Burguera and M.R. Brunetto, At. Spectrosc., 14 (1993) 90. D.L. Tsalev, M. Sperling and B. Welz, Analyst, 117 (1992) 1735. B. Welz, D.L. Tsalev and M. Sperling, Anal. Chim. Acta, 261 (1992) 91. B. Welz, Y.-Z. He and M. Sperling, Talanta, 40 (1993) 1917. T.-Z. Guo and J. Baasner, Anal. Chim. Acta, 278 (1993) 189. S.J. Haswell and D. Barclay, Analyst, 117 (1992) 117. J.L. Burguera and M. Burguera, J. Anal. At. Spectrom., 8 (1993) 235. R.H. Atallah and D.A. Kalman, Talanta, 38 (1991) 167. D.L. Tsalev, M. Sperling and B. Welz, Spectrochim. Acta B, 55 (2000) 339. R.L. Benson, I.D. McKelvie, B.T. Hart, Y.B. Truong and I.C. Hamilton, Anal. Chim. Acta, 326 (1996) 29. D.M.W. Peat, I.D. McKelvie, G.P. Matthews, P.M. Haygarth and P.J. Worsfold, Talanta, 45 (1997) 47.

Chapter 22

Automation of sample preparation M.D. Luque de Castro and J.L. Luque Garcı´a

22.1

INTRODUCTION

22.1.1 Generalities Automation of the equipment used in daily human activities has experienced massive development in recent years. Chemical analysis has required growing automation as a consequence of the exponential increment in the number of analyses that all areas of social interest—particularly clinical, environmental, industrial and food areas—presently demand [1 –3]. Full automation of the analytical process is sometimes the best solution to laboratory overload as 24 h working days are possible without increasing personnel costs. Nevertheless, fully automated analysers have—as a sometimes unsurpassable shortcoming—high acquisition costs with medium-tohigh maintenance costs, mainly caused by the consumable material these analysers use. For economic reasons, partial automation is, in most cases, the accepted solution. The simplest and least expensive partial automation affects the last steps of the analytical process, that is, measurement –transduction of the analytical signal and data collection and treatment. At present, commercial analytical instruments are automated, so these two aspects are addressed most times. Sample preparation (SP), particularly when the sample is a solid, is the one step in the analytical process more difﬁcult to be automated and, of course, the least developed as compared with the subsequent two. The reason for this delay is the different physical states of the samples, which also call for very different devices for their treatment depending on whether the sample under study is a gas, a liquid or a solid. Even within a given physical state, the equipment required for treatment can be very different (e.g. a liquid sample may be corrosive, thus requiring a special material for walls in contact with the sample; it may contain or form volatile compounds during the treatment, making the use of hermetic devices mandatory, etc.). When dealing with solid samples, the variability increases: devices for treatment of a sample of steel, Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

649

M.D. Castro and J.L. Luque

a stone, a food, a soil, etc. must be very different, thus complicating potential design and commercialisation of automated devices for general uses [4]. 22.1.2 Principal shortcomings in automating sample preparation Despite progress, SP remains the bottleneck of the analytical process; it is in the past 10– 15 years only that automation has been actively addressed. By contrast, advances in automated chromatographic and spectroscopic equipment started one or two decades before. The reasons why SP remains the bottleneck of the analytical process vary among experts [5], but the most widely endorsed are as follows: (a) As a rule, SP has always been viewed as a manual technology, often regarded as “low tech”, and usually assigned to the least trained staff. (b) The addition of instrumentation to a traditionally non-instrumental step requires chemists’ conversion to new techniques and chemists are somewhat reluctant to quickly accept new technologies. (c) SP has little academic appeal and is rarely recognised as an analytical chemical area by academe. As a result, many universities have shied away from investigating it. (d) In industry, the chemist’s role is thought to be more “high tech”: to support the use of the analytical instruments as well as analyte detection. Industrial chemists spend most of their time evaluating and implementing new analytical technologies and generating and interpreting data. In addition, analytical techniques are more generic in nature; by contrast, SP can be done in many ways, a number of which are sample speciﬁc. (e) There are not many new concepts for the automation of SP, and early attempts tended to mimic manual operations. These approaches were unsuccessful and expensive as many of the manual procedures were not amenable to automation. (f) Many of the major instrument manufacturers do not identify SP automation as important and it is mainly the smaller ﬁrms that have become niche players and provided the required instrumentation. For this reason, most of the early instruments were stand-alone SP modules that could not be integrated with analytical equipment as parts of the analytical process. The large ﬁrms, however, are slowly beginning to introduce automated equipment for SP. (g) Robotics has been a revolutionary development in the context of SP automation; the implementation of robotic technology, however, is quite difﬁcult and poses strong technological demands. Clearly, robotics and laboratory unit operations constitute the ﬁrst insight into ﬂexible automation of SP. (h) The acceptance time of new technologies is a major constraint in SP. Widespread acceptance of a new technology usually requires a long time

650

Automation of sample preparation

(i)

(often 10 years or more). Also, the time needed to validate the ensuing methods further restricts acceptance of the new technology. Regulatory agencies (particularly in the United States) are very slow to adopt new technologies, mainly as a result of most regulated methods being prescription, rather than performance based. The inertia of most workers, usually reluctant to change and new training, is one other reason for slow acceptance. Financial payback is another major factor in the acceptance of the newer technologies, which are usually expensive. Whether the purchase of laboratory automation equipment is justiﬁed is largely dependent on the speciﬁc intended application and is actually a combination of both ﬁnancial and strategic beneﬁts.

All these constraints have restricted the development of automated SP devices. 22.1.3 Batch versus serial approaches to automated sample preparation Automated SP can be accomplished by using a batch or a serial approach. In the batch mode, multiple samples are prepared and then transferred to the analytical instrument for measurement. In the serial mode, samples are prepared one at a time and the SP device is connected (integrated) with the analytical instrument only. From a temporal viewpoint, it is assumed that, when the SP time is equivalent to or shorter than the analysis time, then a serial method is preferable. Serial methods are also more useful with samples where the assays are time dependent or when there is a stability question of the prepared analyte (e.g. with labile samples or matrices). Examples of better application of serial techniques are OPA-amino acid derivatives for HPLC analysis, where some derivatives have very short half-lives, and for the determination of moisture, where batching of assays could expose the samples to moisture in the environment and give rise to spurious results. When the sampling cycle time is very long, then serial SP makes more sense. Two extreme examples are environmental early warning analysis of contaminated water streams, where samples are collected every 4–6 h, and fast SP procedures, such as microwave digestion. In the latter case, collecting the sample directly from the microwave vessel online makes a great deal of sense—some microwave digesters, however, are integrated with the analytical instrument to perform that task. From the viewpoint of sample availability, if limited sample is available, the user may feel more conﬁdent in preparing one sample at a time; by contrast, batch preparation is to be preferred for abundant, complex samples. Regarding the equipment subsequently used, ofﬂine SP makes sense when the instrument is very expensive (e.g. a GC –MS, which one would not want to

651

M.D. Castro and J.L. Luque

“tie up” during routine use). In general, the serial mode will require the development of low-cost analysers. Where this mode has its greatest potential is in feedback control led systems, which enable real-time (or pseudo real-time) process monitoring. Batch systems are more easily implemented in laboratories with routine loads and less so in laboratories that require a high ﬂexibility or mixing and matching SP and instruments. One other situation where the batch mode might make sense is when a wide variety of SP techniques feed a wide range of analytical instruments. The type of SP procedure involved dictates whether batch or serial approach is required. For example, centrifugation and microwave-assisted digestion tend to require batch processes. On the other hand, new technologies, such as those of axial centrifuges and the new ﬂow-through microwave digesters, have revealed that even traditionally batch steps can be made serial. The batch environment is often an accommodation of the human operator in the laboratory rather than the most appropriate way for a given chemical – instrumental system. Frequently, batch methods are not science based but developed by the constraint of an 8 h working day, so phrases such as “let stand overnight to settle” or “allow digesting for 1 h” are common in these procedures. Automated systems are not so strongly constrained and should not be penalised with human limitations. The human element also dictates the use of the batch mode because of our limited ability to simultaneously keep track of too many events. In short, automated SP is changing most SP protocols and giving a new face to this step of the analytical process, which was formerly envisaged as a clumsy, unproductive step. 22.1.4 Bar codes: a necessary tool in automating routine analyses One of the primary purposes for analytical automation is the possibility of analysing a huge number of samples, either simultaneously or sequentially, and without any human intervention. When large amounts of samples are analysed, mainly in routine analyses, the importance of the correct identiﬁcation of each sample in order to obtain the appropriate correlation between the sample analysed and the results obtained is obvious. At present, the best way to identify a sample, among a number of them, from collection to result, is by using bar codes. A bar code can best be deﬁned as an “optical Morse code”. Series of black bars and white spaces of varying widths are printed on labels to uniquely identify items. The bar code labels are read with a scanner, which measures reﬂected light and interprets the code into number and letters that are passed on to a computer. All bar codes have start/stop characters that allow the bar code to be read from both left to right and right to left. Unique characters placed at both the beginning and end of each bar code, the stop/start characters provide timing references, symbol identiﬁcation, and direction of read

652

Automation of sample preparation

information to the scanner. By convention, the unique character on the left of the bar code is considered the “start” and the character on the right of the bar code is considered the “stop”. Besides the linear bar code, today’s new bar codes are two dimensional, electronic or small computer chips, which sort data in inconspicuous places like a credit card. Traditionally, this technology has been included in clinical and bioanalytical analysers where large amounts of samples are analysed daily and when a correct assignation of the results obtained is mandatory. Trace element analysis is not the most common area of use of this technology. Only some commercial analysers for trace element analysis include this identiﬁcation system, as is the case of the equipment commercialised by Analytik Jena AG, which incorporates a bar codes reader guaranteeing correct sample identiﬁcation. 22.2

AUTOMATION OF LIQUID SAMPLE PREPARATION

The approaches for automation of liquid SP can be classiﬁed into two groups, depending on their operational mode, namely continuous and discontinuous systems. 22.2.1 Continuous systems Continuous automatic approaches to liquid SP are characterised by the fact that the transport of samples along the system is effected by establishing a gas or liquid stream ﬂowing through the straight and coiled tubes making up a typical manifold. The sample can be mixed with reagent(s) in a number of ways, and a variety of intermediate operations from the mere halting of the ﬂow to the incorporation of continuous separation units (such as dialysers, extractors, etc.) can be involved in the operation of this type of continuous approach. Typically, continuous detection systems are used featuring a ﬂow-cell through which the stream carrying the reaction mixture is passed. Some techniques, such as atomic spectroscopy, which do not require a ﬂow-cell, can also be used. The transient signals provided by the sensing system, whose time dependence and shape are obviously a function of the operational mode used, are related to the analyte concentration. Functionally, these continuous systems resemble liquid and gas chromatographs, although their foundation is markedly different. They also differ from discontinuous automatic systems in various respects, namely the fashion in which samples are transported and mixed with diluents and reagents, the manner in which carry-over between samples and reagents is avoided and the type of detection used. These differences obviously result in others such as design, cost and versatility [2]. Automatic continuous segmented analysers were the earliest to be developed in the ﬁeld of automatic methods of analysis. They originated from

653

M.D. Castro and J.L. Luque

the transcendental contribution of Skeegs in 1957 [6], materialised in the ﬁrst continuous dynamic measuring system with sequential introduction of samples and the use of a ﬂow-cell. Sample carry-over was prevented by segmentation with air bubbles introduced between successively aspirated samples. Skeegs’s original idea was consolidated in the development and massive commercialisation of Technicon autoanalyzers and later Skalar assemblies and others. For many years, these were the only alternative available for the automation of high-throughput control laboratories. However, the development of other modes of non-segmented automatic continuous analysis, such as ﬂow injection (FI) [1,7] initially known as ﬂow injection analysis (FIA) and later shortened to FI (both because of protest from ﬂuoro immuno assay practitioners and the fact that in very few of the methods thus developed total analysis actually takes place), completely continuous ﬂow analysis (CCFA) [8], and more recently, sequential injection analysis (SIA) [9], has demonstrated the invalidity of Skeegs’s exclusivist approach. These latter approaches clearly excel over segmented methods in performance (rapidity, reagent saving, cost, ﬂexibility). These facts led Technicon to substitute the commercialisation of the segmented analysers with discontinuous systems, which are discussed in Section 22.2.2. From the very beginning of FI [1,7,10– 12], the technique illustrated two key characteristics, namely versatility and availability of fast and simple construction and modiﬁcation of the manifolds in order to fulﬁl the necessities in each case (see, e.g. Fang’s treatment of this subject in Chapter 21). Neophytes in the technique were delighted in including in the FI manifold multiple conﬂuence points, valves and several separation units as well as detectors, thus demonstrating the possibility of materialising any design, solving almost all problems related to liquid sample handling and treatment and showing their imagination for these tasks. Other salient features of FI are the absence of air bubbles; the insertion of injection of the sample into the ﬂow, rather than being aspirated into it; the potential inclusion of chemical reactions or separation steps, such as dialysis, liquid –liquid extraction, etc.; the control of dispersion of the injected sample; transient detection and the nonequilibrium conditions under which detection is performed (ﬁxed-time kinetic methods). As noted earlier, the main feature of FI is its versatility. Users have traditionally made use of this feature to develop methods of SP for the analysis of trace metals in liquid samples involving different separation units, such as solid-phase extraction columns [13,14], sorption/preconcentration knotted reactors [15,16], etc., or incorporating several reactors to the FI manifold when several treatments assisted by auxiliary energies such as UV radiation [17], ultrasound energy [18] (cf. Chapter 12) or microwaves [19] (cf. Chapter 8) must occur. FI methods are normally developed using laboratory-made approaches. However, there are some commercially available equipment, such as the OI Analytical’s Flow Solutionw IV (FS-IV), used for the analysis of ionic species.

654

Automation of sample preparation

This system provides precise, high volume, and rapid results via simultaneous multichannel operation. The FS-IV features a choice of FI or even segmented ﬂow analysis (SFA), offering the ﬂexibility and advantages of both. This ﬂexible system allows the performance of complex SP steps, such as online distillation, UV digestion, online liquid –liquid extraction and membrane separation. In recent years, FI has been supplemented by SIA [20]. Whereas in FI the samples are injected into a continuous ﬂow of carrier solution, in SIA the sample and reagent zones are sequentially aspirated into a channel to subsequently reverse the ﬂow and transport the stacked zones to the detector. During the course of these operations, the zones undergo some mutual dispersion, and the analyte is converted into a detectable species. Mutual penetration of stacked zones and dispersion of the sample are key parameters that must be studied and adjusted according to the intended measurement [9]. SIA has important advantages over conventional FI: as the manifold does not have to change if the ﬂow parameters or injection volumes are modiﬁed [21,22]; the consumption of sample and reagent solutions is lower; it is very versatile because a single conﬁguration can be adapted for multicomponent determinations using several chemical reactions and multidetection systems. One of the main drawbacks of FI, the difﬁculty of determining more than one or two components with the same set up, can be circumvented using the SIA approach. As in FI approaches, several liquid samples’ treatments have been performed by introducing the appropriate device into the SI manifold. In this way, liquid – liquid extraction for the determination of Fe(III) and total Fe in wines has been performed, using an online device for gravimetric analysis as extraction module [23]; tangential ﬁltration allowing the determination of Fe(II) and 2 Fe(III); or NO2 3 and NO2 in aquatic suspensions containing particulate matter and sediments [24]; solid-phase extraction using a renewable microcolumn with ion exchange beads for the determination of nickel in waste waters and urine [25] and adsorptive stripping voltammetry for the simultaneous determination of copper, lead, cadmium and cobalt, among other elements, in aqueous solutions using dimethylglyoxime and Ni oxime as complexing agent for producing adsorption of the metals on an electrode surface [26]. Most recently, the so-called third generation of FI has emerged. This is the lab-on-valve (LOV) approach, the conceptual basis of which is to incorporate the entire necessary unit operational manipulations required, and when possible, even the detection device, into a single, small integrated microconduit, or “laboratory”, placed atop a selection valve. Particular focus will be placed on its use as a vehicle for pretreatment of complex matrices for determination of trace level concentrations of metal ions by electrothermal atomic absorption spectrometry (ET-AAS) and inductively coupled plasma mass spectrometry (ICP-MS) via exploitation of the renewable microcolumn concept. Despite their excellent analytical chemical capabilities, ET-AAS and ICP-MS often required that the samples be subjected to suitable pretreatment in order to obtain the necessary sensitivity and selectivity [27].

655

M.D. Castro and J.L. Luque

Fig. 22.1. Scheme of the Millennium Merlin System used for mercury analysis. (Reproduced with permission of PS Analytical.)

CCFA [8] is a mode of automatic analysis characterised by (a) unsegmented ﬂow and (b) continuous introduction of the sample as a ﬂowing stream at low ﬂow-rate. This is, therefore, a different mode from that of segmented (SF) or FI modes, both of which have discontinuities of some sort or another. A commercial example of this type of analyser is the Millennium Merlin system by PS Analytical (Fig. 22.1). This system is used for mercury analysis, allowing mixture of the liquid sample with a reductant reagent stream, then gas/liquid separation before detection of the mercury by atomic ﬂuorescence. The exchange of the lamp and ﬁlters gives rise to the Millennium Excalibur system, also commercialised by PS Analytical and used for the determination of As, Se, Sb, Te and Bi. Another example of completely continuous ﬂow systems is that commercialised by Applikon, which allows online determination of Cr(VI), Ni and Cu by standard colorimetric methods adding automatically the appropriate reagent(s) in each case. 22.2.2 Discontinuous approaches Discontinuous sequential or batch automated approaches do not differ markedly from robotic conﬁgurations: both feature mechanical transport, although in discontinuous automated approaches this is accomplished by a conveyor belt or a turntable, whereas with robotic stations it is performed by a robotic arm mimicking the actions of a human operator. The earliest discontinuous automatic systems were introduced after Technicon autoanalyzers (SFA), in response to the need for enhancing

656

Automation of sample preparation

performance in some ﬁelds, particularly in clinical chemistry. In fact, the discontinuous approaches feature a number of advantages over continuous ones: 1. They allow the rapid treatment of a large number of samples. 2. They are more versatile, i.e., more readily adapted to different needs without much alteration. 3. Because of their inherent characteristics (samples are physically isolated from one another) the risk of sample carry-over is much lower and arises from the sampling system. 4. They allow the use of corrosive ﬂuids (e.g. organic solvents, concentrated acids) impossible with continuous systems on account of the risk of the system connections and walls being attacked. However, the discontinuous approaches have some disadvantages or shortcomings that should not be underestimated: 1. (They are generally complex mechanically. In fact, they are cumbersome to maintain as they require skilled workers even for minor repair. Moreover, commercially available discontinuous systems are highly computerised, which adds to the aforesaid complexity. 2. Despite their versatility, their basic conﬁgurations are difﬁcult to change. Thus, in FI and SIA systems, the replacement of some component or the incorporation of another module (from a straightforward to a sophisticated continuous separation unit) is a relatively easy task. Hence, research on, and the development of, automated methodologies is easier using continuous approaches for laboratories without the infrastructure required to incorporate the latest advances in micromechanics and microelectronics. 3. They are generally much more expensive than continuous systems. 4. Because of their conﬁguration, they do not allow the online incorporation of analytical separation techniques, unless they are implemented on selfcontained automated units independent of the analyser. 5. SP using discontinuous systems is similar to that carried out manually. Hence, they allow ready adaptation of manual methods than do continuous approaches insofar as the latter require stricter optimisation of the different chemical and physico-chemical variables involved. Most of the discontinuous automated approaches used for liquid SP are liquid handlers, which are mainly designed and massively used in biochemical laboratories and in a much lesser extension in environmental analysis of metals. These systems vary from the most simple, an automatic dispenser, to very complicated automated systems that allow several operations, such as dilution, liquid reagents dispensing, powder dispensing, vortex shaking, weighing, liquid transfer, autosampling for AA, NMR, IR or UV/Vis spectrophotometers, etc., incorporating, moreover, optional devices such as barcode readers, plate and tube labellers, plate sealers, etc. One of the newest of these systems is the Gilson’s 215 liquid handler, which is a ﬂexible and robust

657

M.D. Castro and J.L. Luque

large-bed device (not a complete instrument as it lacks a detector) designed to accomplish everything from the simplest to very complex liquid handling tasks (Fig. 22.2). It offers 100 standard racks and a large array of custom racks to allow the use of a variety of sample vessels, such as microplates, microcentrifuge vials, test tubes, scintillation vials, bottles and so forth. A Z-arm enables the piercing of thick septa while the liquid level detection system coupled with the inside/outside probe rinse station reduces cross-contamination. Solid-phase extraction steps are also automated in a huge number of liquid sample analyses. The commercialisation of this type of system is widely extended. An example of such an automated solid-phase extraction system is the Gilson’s Aspec XL4. This system processes four samples in parallel with identical treatment of each sample, allowing processing up to 50 samples per hour. It also operates in sequential or batch modes and can be conﬁgured to collect single or multiple fractions. This system is mainly used for pharmaceutical bioanalysis; however, it can also be used for trace metal preconcentration, as the system is compatible with either standard solid-phase extraction cartridges. A similar system, but one specially designed for trace metal preconcentration, is commercialised by Dionex and uses chelation ion chromatography to provide metal preconcentration in minutes rather than hours. Several methods have been developed using preconcentration chelating columns, allowing determination of trace transition metals in reagent grade acids, bases, salts [28] and water miscible organic solvents [29], as well as elimination of iron and aluminium interferences in sample matrices [30]. A non-commercial example of a hybrid discontinuous –continuous manifold for special SP is that shown in Fig. 22.3. It is a portable approach, which can be

Fig. 22.2. Gilson’s 215 liquid handler. (Reproduced with permission of Gilson.)

658

Automation of sample preparation

Fig. 22.3. Hybrid discontinuous–continuous manifold for monitoring trace metals in sea water. (Reproduced with permission of Elsevier.)

installed aboard a ship for monitoring trace metals in seawater. As can be seen in the ﬁgure, the two previous treatments of the liquid sample, ﬁltration and UV digestion, are performed in a continuous manner; meanwhile, preconcentration of the metals on the electrode surface is performed in an automated batch-mode [31]. Aliquots of 10 ml can be processed at a rate of one every 10–20 min. The features of the calibration software result in high-quality data, as required for biogeochemical and pollution studies [32]. 22.3

AUTOMATION OF SOLID SAMPLE PREPARATION

The various approaches enabling automation of solid SP to variable degrees can be classiﬁed into two main types: (1) one-step automation; (2) direct sampling. Robotics, which allows either several steps or the overall analytical process to be automated, is considered in a separate section due to its special features. 22.3.1 One-step approaches to automation and acceleration of solid sample preparation Most commercial approaches for accelerating either sample dissolution or leaching—the latter usually known as “extraction”—have appeared within

659

M.D. Castro and J.L. Luque

the last two decades and have been the subject matter of previous chapters. Thus, processes such as microwave-assisted extraction or dissolution and supercritical ﬂuid extraction are discussed in Chapters 8 and 10, respectively; pressurised solvent extraction, better known as accelerated solvent extraction (ASE, a name coined by Dionex) is described in Chapter 11. Sonication, which can assist a wide spectrum of processes, including leaching and digestion, without requirement for special sample cells but only the use of an ultrasound probe of the appropriate frequency and pulse amplitude, is the subject of Chapter 12. All these approaches require previous assistance of the user for weighing the sample, locating it inside the chamber and initiating the extractor. After the leaching or extraction step, usually drastically accelerated, user actuation is mandatory; at least the extractor is connected online with equipment for the subsequent step, namely, a chromatograph or a highly discriminating detector. In short, these approaches reduce the time for putting in solution the target analytes as compared with conventional, manual approaches. 22.3.2 Direct solid sampling Direct solid sampling refers to equipment used wherein either a part of, or the overall sample, is treated prior to or at the point of detection, for proper monitoring. One of these approaches, such as laser ablation (see Chapter 20) can involve either removal of a portion of the sample—the ablated material— with subsequent transport to the detector (MS, ICP-emission or ICP-MS) or in situ excitation and monitoring of the emission (laser induced breakdown spectrometry or LIBS). Similar in situ sampling, excitation and detection take place with glow-discharge spectrometry. With electrothermal atomisation (and excitation, if required) the sample is placed in the atomiser and is subject to the steps required for the analysis—see Chapter 5. One controversy over terminology arises from the ordinary usage of “solid sampling” in relation to solid samples delivered to an electrothermal device, acting as electrodes in glow-discharge applications or being “ablated” by a laser. It is not clear if it is correct to use this designation when stating that, “the sample is weighed and placed in the graphite tube or on the L’vov platform”; “the sample is placed in the glow-discharge chamber”, or “the sample is positioned on the XYZ translation stage for laser irradiation”. When sampling takes place with these techniques, before of after the sample is introduced into the speciﬁc device used, is a matter for discussion by users.

22.4

ROBOTICS

Although robotics constitutes a discontinuous approach to automation, its special characteristics lead most authors to consider it as a separate means of automation.

660

Automation of sample preparation

A robot can be deﬁned as “an automatically controlled, reprogrammable, multipurpose, manipulative machine with several degrees of freedom, which may be either ﬁxed in place or mobile for use in automation applications” [33] or, more loosely, as “a multipurpose machine which, like a human, can perform a variety of different tasks under conditions that may be unknown a priori” [34]. The different types of robots currently available can be classiﬁed according to physical features, such as hardware construction, degrees of freedom, coordinate system or level of sophistication and technology [35]. Obviously, robotic techniques are more complex than other automated laboratory analytical techniques. Laboratory robots were originally marketed as do-it-yourself technology. Some of the ﬁrst practitioners were chemists who were not skilled in the engineering and software aspects, so highly trained, dedicated personnel were needed. In many applications, the overall lead times to implementation became excessive and chemists were often sidelined for months awaiting the tools they needed to do their job. Although robotic technology is claimed to provide ﬂexible automation, in fact it possesses limited ﬂexibility and applicability. One other reason for the slow acceptance of robotics has been its presentation in the game doing whimsical or trivial tasks, which caused potential users to overlook serious applications. Fortunately, robot-based drug candidate screening systems represent a success story; for time-to-market and labour- and time-saving reasons, these systems are making a comeback in this very speciﬁc pharmaceutical ﬁeld. The proprietary nature of robotic communications and software has also restricted their acceptance by laboratories. The lack of standardisation among robot suppliers made it difﬁcult to connect the robots to analytical instruments and other peripherals in a straightforward manner. The initially envisaged uses of a robot in the laboratory, which led several renown manufacturers to failure in the past, have changed with time. The early robotic units tended to mimic human manipulations, which made interfacing cumbersome and resulted in haphazard implementation of many applications. In general, robots were not really optimised for task-oriented work; their most general use was as sample-transport devices that moved samples or surrogates among a series of dedicated workstations, which were optimised to do speciﬁc tasks, such as solid-phase extraction or ﬁltration. At present, robots are devoted to things that users could hardly ﬁgure out how to do easily in any other way. These unique needs range from massive robotic systems for handling thousands of samples produced in high-throughput drug discovery and biochemical study programmes to stand-alone devices that drone away at repetitive tasks. Depending on both the number of tasks to be performed and their complexity, users can currently choose between robotic stations and workstations. The former automate the entire processes; they often include facilities for sample insertion into analytical instruments and use the typical arm and an array of modules and peripherals to make tasks easier for the robot. The original

661

M.D. Castro and J.L. Luque

idea of laboratory robotics revolved around a human-like arm capable of handling many analytical tasks. Today, manufacturers design arms to function as “movers of stuff”, and operations such as transferring or ﬁltering solutions are handled in modules around them. Workstations represent a newer vision of robotics. They are custom automation tools that assist workers in performing their job better and more efﬁciently. These stand-alone automated units are used for speciﬁc purposes, such as handling liquids or preparing solid samples; they are growing in popularity as they can perform one to three tasks with reduced costs and sophistication. Some workstations can be interconnected or operated with robotic arms, while others use their own, dedicated arms (particularly to handle solid samples). One of the main differences between robotic arm systems and workstations is cost. Thus, a system built around a robotic arm costs more than $100,000 whereas workstations can run from $10,000 to 100,000, depending on their level of sophistication. However, it is the intended use, as much as cost, which often dictates the choice of design. Because robotic technology continues to have some magical connotations in relation to laboratory automation, a number of manufacturers and users still use the words “robot” and “robotic” indifferently to refer to both robotic stations and workstations. In addition, any instance of automation is also indiscriminately associated with robotics by many. One case in point is the Internet page http://www.lab-robotics.org/manufact.htm, wherein about 100 companies summarise the highlights of their products, many of which bear little or no relation to robotics. Both suppliers and users consider most dispensers and liquid handlers highlighted in Section 22.2.2 as robotic approaches. 22.4.1 Workstations, robots, modules and peripherals The most salient difference between robotic stations and workstations is that, whereas a workstation can only be used for the tasks (all or some) for which it was constructed, robotic stations can be modiﬁed by changing software, modules or peripherals as required to undertake one or more speciﬁc tasks, or even a whole analytical process. As a result, describing a workstation is as simple as listing its intended functions, whereas characterising a robotic station includes stating the type of arm it uses and the equipment that helps the arm perform its tasks. Figure 22.4 depicts the scope of application of each option in terms of task complexity and throughput. 22.4.1.1 Workstations The most simple and common workstations are those for dilution and/or reagent addition to a number of samples in a simultaneous manner, either to all samples in a rack or to a line with a slide z-axis (as in the Biomexw 2000 model from Beckman). Most workstations are designed to operate with liquid samples, such as with those from Cyberlab, Gilson, Zymark, SciLog, Sagian,

662

Automation of sample preparation

Fig. 22.4. Fields of application of workstations according to task complexity and throughput.

Beckman and Hamilton, which manufacture speciﬁc equipment for liquid handling, solid-phase extraction and preparation of liquid samples for insertion into chromatographs. Some ﬁrms (e.g. Zymark, Bohdan Automation, Source for Automation) also manufacture workstations for automatic dissolution of, mainly, pharmaceutical products; however, they can also be used to weigh and dissolve (or leach) other, speciﬁc types of solids. The diversity of equipment with which manufacturers have ﬂooded the market is exempliﬁed by Zymark corporation. In the last few years, this ﬁrm has launched six different types of workstations for handling both liquid and solid samples—particularly the former, whose characteristics and external aspects can be found on the Web site of the company. Figure 22.5A and B depict the prelude and the tablet processing Zymark workstations, both of which were designed for treating solid (general and speciﬁc, respectively) samples. 22.4.1.2 Robots A laboratory robot constitutes the most ﬂexible tool for automation as it can reproduce, with minimal adaptation, almost every task performed by an analyst. A laboratory robot is essentially a machine consisting of various parts, namely: (a) The manipulator, a mechanism usually comprising various segments, whether jointed or sliding relative to one another, for the purpose of grasping and/or moving objects, usually in several degrees of freedom. The essential parts of the manipulator are the body, the arm and a hand or endeffector. Robots can usually be furnished with various types of hand in order to grasp objects of different size or shape. Based on the co-ordinate system they use, manipulators can be classiﬁed into Cartesian or gantry (deﬁned by the x, y, z three-dimensional system, with translation motion

663

M.D. Castro and J.L. Luque

Fig. 22.5. Zymark workstation for solid sample treatment. (A) Preludee model. (B) Tablet Processing II model. (Reproduced with permission of Zymark Corporation.)

but no radial motion), cylindrical (deﬁned by the x, z two-dimensional system and the f angle, with a radial motion and two translation motions), spherical [deﬁned by the x one-dimensional system and two (f and w) angles, with one translation and two radial motions] and revoluting or anthropomorphic (with all motions of the radial type). (b) The controller, an information processing device whose inputs are both desired and measured position, velocity and other pertinent variables of the process. The robot can be controlled by the user, in a point-to-point manner in the case of demonstration robots and by a computer otherwise. Computer-controlled robots are much more common and beneﬁt from continuous improvement derived from the development of new software programming languages. (c) The power supply, which provides the energy for locomotion of the different manipulator parts. Laboratory robots are usually equipped with electrical or pneumatic energy sources.

664

Automation of sample preparation

(d) Sensors, which are used to boost performance in intelligent robots. These devices can be of various types (e.g. stereoceptive sensors) and differ in their physical (e.g. the way variables are transduced into analogue or digital signals) and computational foundation (e.g. the way sensory data are processed to obtain sensory information). Sensors can be ﬁxed or mobile and of the contact or non-contact type, depending on whether the target variables are sensed with or without physical contact with some body contained in the world. Force and tactile sensors are of the contact type and vision, proximity and range sensors, of the non-contact type. One of the most important features of a robot, which determines its work envelope (i.e., the maximum extent and reach of the robot) is its position in the robotic station, which can be ﬁxed or variable. The former is used in circular robotic stations, which are typical of Zymark’s Py technology. In this conﬁguration, the robotic arm is in the centre of a circle and the different peripherals are included in a work envelope radius as removable pieces of a pie. The work envelope of the arm in this case is 3608 and the radius equal to, or shorter than, the extent of the arm (see Fig. 22.6A). A mobile arm in the robotic station is supported in a track (Fig. 22.6B), which allows displacement of the arm in a length that varies, depending on the particular station, from 1 to 2 m. The capacity of the robotic station for locating peripherals in the work envelope of the arm can be expanded by using a more complex arm capable of operating on both sides of the track. Robotic stations surpass circular ones in throughput but the two are similarly accurate. 22.4.1.3 Modules and peripherals The modules of a robotic station are the devices (apparatus, instruments, racks) used by the arm to perform its tasks. In circular conﬁgurations, the modules are referred to as “peripherals”. The number of modules present in the work envelope of a robot arm varies with the number of tasks it is to perform in a given process. Some modules can be as complex as self-contained workstations. Although most of the modules required for the different steps of the process can be provided by either the arms or an alternative manufacturer, some users design and construct their own modules, either because of the high speciﬁcity of the task or with a view to reducing costs. A detailed discussion of such custom modules is obviously beyond the scope of this section. What follows is thus a brief description of the most general of these devices. The balance is the most important module of a robotic station in dealing with solid samples. The only way of automating weighing is by using a robot or workstation. In addition to being connected to the power and event controller (PEC), or central computer, the balance must be accessible to the robot arm, which should be furnished with a special end-effector or hand (a gripper hand) and a powder-pouring device (usually vibration based). Segregation of sample

665

M.D. Castro and J.L. Luque

Fig. 22.6. Robotic stations with ﬁxed and mobile arms. (A) Circular, PyTechnology from Zymark. (B) Linear track from Hudson. (Reproduced with permission of Zymark Corporation and Hudson Control Group, respectively.)

particles in terms of size can be caused by the programmed vibration of the source tube during the transfer and sample weighing step, which causes a bias in subsequent data. In this case, the pouring routine can be altered by rewriting the software subroutine to eliminate vibration and transfer the sample material in a single step [39]. Weight-based measurements of liquids are frequently used in robotic stations as they provide more accurate data than do volume measurements. Zymark’s new automated ﬁlter weighing system is an excellent example of the use of the balance with gaseous environmental samples. It provides automated and exact repetition of the analytical steps for compliance with EPA PM2.5 methods and uses no external air source in order to avoid introducing contaminants; also, ﬁlters are located before and after each weighing, and also static ion-discharged before weighing.

666

Automation of sample preparation

A dissolve and dilute module is a dual peripheral that includes a vortex mixer to facilitate dissolution and the achievement of a homogeneous solution, respectively. A master laboratory station (MLS) is a module consisting of three syringes for dispensing liquids in conjunction with the dissolve and dilute module. However, the MLS can be used for additional purposes, such as aspirating the phases involved in a liquid – liquid extraction (using a different syringe for each phase) [40]. A centrifuge is a key module when solid samples are to be leached or a precipitate forms during the process. The use of sensors to ensure correct positioning of the tubes with respect to the robot before and after the centrifugation step is mandatory here. Computer vision and neural networks have been used to detect errors in a robot system performing the automatic loading and unloading of a centrifuge [41]. All-purpose hands and syringe hands, available in a variety of designs, are also required elements of a robotic station. Different sized objects (e.g. sample ﬂasks, test tubes, probes, hold and press push-buttons) also call for different types of hand. A syringe hand facilitates the withdrawal of liquids from vessels. Hand design has beneﬁted from innovations devised by academic research groups [42]. Liquid – liquid, solid– liquid and liquid– solid extraction modules are also required devices for implementation of a number of methods. Racks can hold a variable number of test tubes, the positions of which are numbered so that the robot can distinguish them. In addition to holding the tubes, racks can accommodate devices such as the aspirator assay probe, or even the sample probe if one is used [40]. Additional modules not always required in a robotic station include a capping– uncapping module, used to remove and replace screw caps; a bar code reader, which is usually a laser bar code scanner combined with a turntable assembly capable of reading a label positioned anywhere around the circumference of a vial; and an ultrasonic bath, which is required for sonic mixing or cleaning, but also, occasionally, to facilitate dissolution or leaching. The performance of robots in some processes can be improved by using various approaches to expedite one or more steps thereof. One such approach involves adapting non-automated devices for automated use in a robotic station. Such is the case with microwave digesters, which substantially accelerate solid SP. For example, an ordinary focussed-microwave extractor can be loaded and unloaded by impulsion and aspiration, respectively, using a peristaltic pump [36,37]; subsequently, specially constructed ﬁngers can be used to handle the digestion vessels [38]. Also, the dawn of genome analysis protocols relied on automated laboratory work, including the pipetting of reaction mixtures and reagents, loading of samples onto gels and grinding of clones onto ﬁlters [39]. Some researchers have succeeded in adapting robotic-based equipment to

667

M.D. Castro and J.L. Luque

their speciﬁc needs [40,41]. Modular robotic systems have been developed to eliminate bottlenecks in different areas and to allow a choice of specimen introduction by hand, conveyor or mobile robot [42]. Robot control is one other current research topic for various university researchers. This area encompasses new hardware concepts for robot control [43], software environments for optimising the productivity of robotic laboratories [44] and dedicated software for use in environmental laboratories [45]. Finally, simulation and graphical animation of robots have proved highly useful with a view to both optimising robot work and avoiding catastrophes on the workbench [46]. Incorporating robotic stations into large-scale laboratories entails establishing appropriate links with a laboratory information management system (LIMS) to ensure precise control of the laboratory as a whole [47]. Zymark’s Clara 2000 software package exempliﬁes the incorporation of advances in computer science into commercially available robotic laboratory equipment. Its release followed the acquisition of Scitec Automation Holdings by Zymark. Based on an upgraded version of the Scitec Clara LT open architecture software, Clara 2000 provides substantial advantages in modularity, stability, Visual Basic customisation capability and upgradeability for the future. Contributions from academic research groups have also been reported. One example is the practical approach to real-time scheduling and multitasking at the computer level developed by Entzeroth [48], which covers scheduling software, handling of errors and unexpected situations, and data management. A diagram of work and data ﬂow during the screening process, and ﬁgures which show the scheduling or robotic movements of a receptor binding assay, an error handler for an exception-speciﬁc situation, and one for a general system error have also been reported. 22.4.1.4 Common and differential aspects of workstations and robotic stations Two essential facts to be considered when purchasing a robotic station are that no single ﬁrm builds everything one is bound to need to construct a robotic system tailored to their needs, and that not all commercially available equipment operates by the same set of instructions or plays by the same set of rules. Compatibility between modules from different sources should thus be carefully checked prior to purchasing. Reliability, which is one other key feature, depends on the particular components of the robotic system and its design. Because systems are linked to operate serially, even small losses of reliability at each device can be magniﬁed at the end. There is one other consideration in setting up robotic equipment that is shared by fully automated systems in general, namely the surprising number of disposable items, such as pipette tips, autosampler vials, ﬁlters and containers they go through. Even the microlitre amounts of solvents consumed can add up to large volumes in high-throughput operations. Users must bring these

668

Automation of sample preparation

ancillary materials into the laboratory and then dispose them; as a result, they may create their own small superfund site. Also, not all pipettes or autosampler vials are usable; thus, unlike a human, a robot cannot use a pipette with a crooked tip. On the other hand, disposables should be of a high quality to ensure reliability. Procurement of supplies and waste disposal are therefore two important issues in setting up and maintaining a robotic laboratory. Because workstations can be designed and dedicated to a single, speciﬁc task, they are normally simpler, mechanically and usually more reliable than are robotic arm systems. In addition, they can provide a data trail for regulatory compliance and are typically designed to be operated by nonexperts. Commercial workstations vary in their level of sophistication, which allows the automation of even the simplest procedures. Economically, automation can beneﬁt anybody running more than 150 samples a day and can justify purchasing a moderately priced workstation. More sophisticated workstations, however, are only proﬁtable with a heavier workload. As a rule, a workstation will replace a $40,000-a-year technician and will have a one- to two-year payback period. Workstations operate best when there are one to three functions being automated; more operations or less deﬁned analytical steps require the sophistication of robotic arm systems, which are more ﬂexible and work best early on in research, when the methodology is not well deﬁned. The combination of arms and modules with workstations has real advantages. A comparison of the efﬁciency of a tracked robot handling compound dissolution in collaboration with a liquid-handling workstation versus the robot alone revealed the robot – workstation team to increase productivity by 130% [37]. Where these types of systems pay off the most is in high-throughput discovery and screening programmes. The automated systems developed to handle these chores can cost millions of dollars to set up, require full-time experts in automation to design and maintain, and demand a huge ﬁnancial commitment to operate. In any case, robotics is the only way to go for these types of applications. Choosing between a robot plus peripherals and a workstation is a difﬁcult task. From the beginning, automation held the promise of freeing analysts from cumbersome, time-consuming, repetitive tasks. This is especially true with the quality control (QC) laboratory, which must routinely test products such as pharmaceuticals or foods prior to release, often with a well-deﬁned analytical procedure dictated by regulatory requirements. In these laboratories, workstations are typically the best solution as they are often more “hardwired” and are better in QC laboratories, where the analytical steps are well understood and this equipment does save laboratories time and money. The best solution for implementing the complex treatments required by some solid samples is the sequential use of two workstations; when this is impossible, a robotic station is the next best choice in most instances.

669

M.D. Castro and J.L. Luque

22.4.2 The role of robots in the analytical process Automation of the analytical process by use of robotic equipment (robotic stations and workstations included) can range from a single step to the whole analytical sequence. The number of steps that are robotised should be dictated by the user’s experience and judgement, always as a function of the target process, costs, number of samples to be processed, etc. Straightforward singletask uses of robots, robotic SP procedures and fully robotised methods are discussed below, as are more rational uses in combination with other techniques intended to ensure optimum development of each step of the analytical process. 22.4.2.1 Single-task and simple uses of robotics One of the most important reasons for failure at the beginning of the laboratory robotics era, when no workstations were available, was the use of large robots to perform a single, repetitive task, such as weighing, diluting or solid-phase extraction. One of the most common single tasks assigned to robots is weighing, which is the analytical step most difﬁcult to implement using alternative automated approaches. Automating laboratory weighing may not increase the speed of weighing, but can free the valuable time of scientists and technicians. It has been claimed that, where a balance is in constant use for 4 h or more per day, the investment in a weighing automation system will be paid back [49]. Sartorius, one of the main European analytical balance companies, has combined its Genius ME analytical balance with an advanced benchtop robot to facilitate laboratory weighing processes. The robot gripper can pick up microtubes, vials or microﬂuidic products ranging from 4 to 40 mm and transport them to the balance. Samples can be identiﬁed by one-dimensional or even two-dimensional bar codes. The widest range of tube and vial sizes can be placed on the platform, which accommodates up to 1920 microtubes (20 racks, each holding 96 tubes) or approximately 500 test tubes or bottles. The gripper and the conﬁguration of tubes or bottles on the platform can be customised to meet the speciﬁc requirements of a laboratory. The balance, with a resolution of 0.01 mg, is placed separately on its own platform to prevent mechanical interference. This reduces the time it takes for the weigh cell to stabilise and minimises the uncertainty of measurement, which has a positive effect on the total throughput of the robot. Completing the system is automation software based on Windows NT/2000, which combines simple, graphic programming of methods using widespread Windows functions, sequence control, and evaluation and storage of data. The logical, easy-to-understand operator guidance features allow the microtube or bottle layout and methods to be quickly and easily adapted to the user’s speciﬁc requirements. Simple, inexpensive workstations designed for speciﬁc tasks include the TurboVape concentration workstations and the RapidTracee workstation, which have so far found widespread use mainly in clinical analysis.

670

Automation of sample preparation

Non-commercial systems for single tasks include a microwave digestion system for dissolution of Ti(IV) oxide [50,51] and adaptations of workstations for special tasks such as the robotic-chromatographic method for the determination of glycosylated haemoglobin using a reprogrammed Hamilton Microlab 2200 pipetting Cartesian robot [52]. A robotic system was also used for the accurate, precise preparation of calibration standards, and automated, unattended multispecies preparation for both anion and cation analytical channels to support continuous online ion chromatography operations. Two robot Py-sections of a Zymark robot were designed, assembled and programmed in EASYLAB control language in command modules, the system resulting in substantial workforce savings [53]. Also, a robot was interfaced to an expert system for development of a standard addition method. In this way, preliminary input of information by an operator and subsequent control of the robot table set-up, the concentration of cation addition solution, the monitoring wavelength, sample pH adjustments and the method of reagent addition were enabled. Three standard additions were run by the robot and a calibration line was constructed and tested for ﬁt to a linear model, the system ﬁnally determining whether more additions were required [54]. 22.4.2.2 Sample preparation SP is the most general application of both workstations and robotic stations as the tasks involved in this step of the analytical process are the most timeconsuming, error-prone and difﬁcult to develop by unskilled operators; in addition, safety restrictions apply when toxic materials are to be handled. The use of a speciﬁc approach depends on the number of steps involved and their complexity. As a general rule, whereas most environmental samples subjected to a robotic treatment are solid, those dealt with by clinical laboratories are typically liquid and highly complex in nature. The type of analyte to be isolated from a given sample and the nature of the latter determine the operations to be performed by either the workstation or the robot to prepare the sample for subsequent steps, as well as the main steps following the treatment. The operations carried out after preliminary operations dictate the nature of the latter. When the step following treatment of the sample is insertion into a highly discriminating instrument such as an NMR or ES-MS instrument, the robotic station is aimed at a 24 h working day of the high-price instrument in an unattended manner, thus avoiding either the high-maintenance personnel costs of continuous work or the purchase of another instrument. This task even justiﬁes the use of a simple robotic station to insert samples with non-complex matrices into the instrument. Although workstations and even robotic stations commonly perform liquid –liquid and liquid –solid separations, these steps can be implemented in a continuous manner by using a more inexpensive set-up and with shorter development times when the number of samples is not very high and the results must be available within a short time. The unit operations that can be

671

M.D. Castro and J.L. Luque

performed failure-free by a robot include weighing, centrifugation, ﬁltration and special instances of solid and liquid transfer. On the other hand, reagent addition, dilution, heating, homogenisation, derivatisation and insertion into a measuring instrument are generally more rapidly, economically and efﬁciently performed by other automated alternatives, whether continuous or discrete. 22.4.2.3 Robotic development of the whole analytical process Most frequently, using a robotic station to develop an entire analytical process is unwarranted. In the mid-1980s, when robotic technology ﬁrst reached the analytical laboratory, robots were more of a novelty than a useful tool. At that time, conventional manual titrations and similar easy tasks were entrusted to robotic stations. At present, however, well-established criteria exist to ensure correct use of the potential of robotic technology. The use of automated titrators, whether photometric or potentiometric, to complete the work performed by the robot is not an instance of hyphenated batch methodologies but rather one of the functioning of the titrator as a module of the robotic station. Such is the case with the determination of fuel parameters including the diene value [55] and mercaptan sulphur [56]. In the former application, the robot weighs the sample, reﬂuxes it in the presence of maleic anhydride and extracts the analytes into an aqueous phase, which is poured into the titration vessel, where the robot inserts the photometric or potentiometric probe. For the determination of mercaptan sulphur, the robot also weighs the sample and removes sulphide by precipitation with a CdSO4 solution and liquid –liquid extraction. Once the fuel is sulphide-free, which is checked using a photometric probe, it is poured into the titration vessel by the robot arm, which also plunges an Ag electrode prior to starting addition of the titrant (an AgNO3 solution). In both cases, the automated titrator acts as a module of the robotic station and is operated by the robotic arm. An ICP-AES instrument has been coupled to a robotic station from a sample vial following treatment by the robot [57]. Coupling of mass spectrometric and NMR detectors used in this context has also been based on direct aspiration. 22.4.2.4 Coupling robotic and continuous systems as the most reliable development of the overall analytical process The “bridge” spanning the raw sample and reading of the analytical signal, which is a function of the analyte concentration, has traditionally been built around single batch, continuous or robotic methodologies for driving the sample to the detector. Each approach has its own advantages and disadvantages, and no single currently available choice for automation is the panacea. Combinations of a robotic station or a workstation with a liquid [58–76] or gas chromatograph [70,71,77 –81] or a capillary electrophoresis instrument [82] have been reported in which an autosampler was used as an interface, the robot arm either placing the vials in it or loading them (following positioning on the autosampler) with the treated samples.

672

Automation of sample preparation

The Zymark –Waters partnership for the development, among others, of an intelligent software interface between the Zymark TPWe platform and the Water Millennium Chromatography Data Manager is an example of the need to make robotic equipment and chromatographs compatible. Users experienced in FI methodology proposed this continuous approach [2,7,83] as the most easygoing friend of robotics in order to take advantage of both. Thus, the expeditiousness and simplicity of FI systems are offset by their inability to automate certain operations (some so essential as weighing); on the other hand, robots, which are the most powerful systems as regards the variety of processes they can develop, have an inherent relative slowness compared to ﬂowinjection systems in addition to a high purchase cost. The different aggregation states of the sample involve different degrees of difﬁculty for inserting them into an FI manifold: whereas liquid samples can be directly introduced, solid samples require one or more previous operations. The pretreatment steps can be shortened by using a more complex hydrodynamic system [84]. Thus, while the use of several automated approaches to circumvent the problems encountered in automating laboratory processes is desirable, the conservative policies of equipment manufacturers and the usual inertia of research groups to drastically change investigation guidelines have so far delayed advances in the combined use of several automated approaches. The complementary features of the FI technique and robotics warrant their joint use with a view to exploiting their respective advantages while avoiding their shortcomings in isolation. Surprisingly, however, the uses of such a promising couple have scarcely been explored so far. The two ways of combining FI systems and robotic stations are by having the two individual systems operate independently of each other, wherein the user acts as an active interface between both [85] and, the most operative, by setting up an integrated system where the two subsystems will interrelate to eliminate the need for human intervention. When the FI manifold acts as a module of the robotic station, the different operations involved in the analytical process are distributed between modules in order to optimise the performance of the overall system while avoiding the need for human intervention in communicating between the two systems physically and logically. This approach was developed by the authors’ research group. As demonstrated for more than 25 years, FI facilitates the automation of a number of separation steps—namely, solid-phase and liquid – liquid extractions, dialysis, gas diffusion, pervaporation, and so forth—as well as a large number of types of reactions developed with or without the help of auxiliary energies, such as microwave, ultrasound, heat, UV radiation, etc., easily acting on the given FI unit. Provided the sample is liquid or the analytes have been dissolved, practically all the possible steps of SP can be developed in an FI manifold in a faster, simpler and less expensive manner as compared with other automated, continuous or discontinuous, alternatives.

673

M.D. Castro and J.L. Luque

For this reason, after weighing and dissolution or leaching, as in the case of solid samples, the best way of developing subsequent steps of SP is by aspirating the extract or digestion to the dynamic manifold. The robotic station –FI coupling has provided methods, as is the case of the fully automated method for the determination of metals in soil. The use of a two-channel FI manifold to introduce robot-treated (weighed, microwavedigested, centrifuged) samples into the measuring instrument (i.e., an atomic absorption spectrometer) enables real and pseudo-dilutions with a dramatic enhancement of the calibration range. In this way, the need for a prior estimation of the analyte concentration is avoided. In addition, such an expensive peripheral as a syringe hand is replaced with an inexpensive manifold, which reduces purchase costs [37]. The overall operational set-up for solid sample pretreatment plus determination is shown in Fig. 22.7. The addition of modiﬁers to the sample solution to be introduced into the spectrometer is also facilitated by an FI manifold, either mixing them with the carrier solution, using an auxiliary injection valve [86] or an intimately coupled valve arrangement [87]. 22.4.3 Analytical scope of robotics for sample preparation Although robots can be used in virtually all analytical areas, their major applications encompass the clinical, pharmaceutical and biotechnological ﬁelds, with minor but growing interest in the environmental ﬁeld. The large number of samples typically handled for daily analysis by a clinical laboratory promoted the manufacturing of the well-known airsegmented continuous autoanalyzers, mainly from Technicon [88]. The original robot arms gradually replaced autoanalyzers and were eventually superseded by workstations for speciﬁc tasks, such as pipetting, diluting, reagent addition (i.e., sample-handling tasks) [89]. Dedicated clinical laboratory workstations are used to receive samples in capped, bar coded vials in an indexed holding rack that the robot decaps in a decapper and introduces into the analyser(s) as a function of the speciﬁc bar code [90]. The use of robots in the health care sector is not limited to the clinical laboratory; in fact, it also encompasses tasks such as assistance to surgical procedures, elimination of mundane chores and reduction of exposure of personnel to communicable diseases. The pharmaceutical ﬁeld has exploited robotic stations for the main purposes of dissolution testing and drug design. In fact, most of the procedures described in robotic manuals (e.g. Zymark’s monographs and manuals [91]) are concerned with the determination of drugs in pharmaceutical preparations. Robotic stations, as such, were initially also programmed for such simple use as dissolution testing. However, the most important pharmaceutical use of robotics at present is in combinatorial chemistry in the search for new drugs. The impact of robotic

674

Automation of sample preparation

675

Fig. 22.7. Robotic station for the fully automated determination of metals in soil. (– – –) Passive interface. (—) Tubing. Active interfaces of the computer to the robot and its peripherals are not shown. (Reproduced with permission of Elsevier Science.)

M.D. Castro and J.L. Luque

high-throughput screening on drug discovery is a consequence of its ability to screen up to 100,000 samples per day per instrument; this in turn results from the demand for new compounds, which is being met by combinatorial chemistry [92]. Biotechnology is no doubt the most rapidly expanding ﬁeld of application of robots. After years of hard work, the human genome was eventually deciphered in 2001. In fact, genomics continues to be one of the most exciting ﬁelds of research, and has been joined in this respect by proteomics—the word in the mouth of all attendants of the 2001 Pittsburgh Conference—and by metabonomics. Most robotic biotechnological research work is conducted by workstations. Agricultural laboratories are increasingly taking advantage of the use of robotics as seasonal overloads call for a 24 h working day without involvement of temporary personnel. Thus, tasks such as weighing, leaching, ﬁltering and measurement are frequently performed by robots in the determination of parameters such as pH [93], organic matter [94], conductivity [95], lime [96] and phosphorus in soil [97] or bitterness in virgin olive oil [98]. 22.5

ADVANTAGES AND DISADVANTAGES OF AUTOMATION OF SAMPLE PREPARATION

The advantages of automating SP are the result of the aims pursued in adopting this modus operandi. They have been traditionally related to: 1. Samples, which occasionally are dealt with in large numbers or too scarce or valuable to be handled manually. 2. Analytes, which are sometimes present in very dissimilar (macro components, traces) or low (ultra traces) concentrations in the sample. 3. Reagents, some of which are scarce or expensive (e.g., enzymes and biochemicals in general). 4. Rapidity, frequently essential in large laboratories, such as those in hospitals, urgently requiring the analytical results (e.g., clinical parameters in acute crises or shock treatments), and in industrial and other laboratories requiring constant availability of data for process control. 5. Economy in personnel and material expenditure. 6. Precision, closely related to the elimination of both deﬁnite and indeﬁnite errors arising from the so-called “human factor” (tiredness, mood, prejudice, pathological complaints and so forth). 7. Safety, concerning both the use of toxic reagents—particularly organic, volatile solvents—and the manipulation of toxic samples. The above outstanding advantages offered by automation of SP cannot lead to mythicising this manner of laboratory work, as a series of shortcomings derive from the partial or complete elimination of human intervention in SP, in

676

Automation of sample preparation

particular, and in the analytical process in general. These shortcomings affect both and they should be taken into account in deciding whether a given step is to be automated or not and, if yes, which alternative enables minimisation of the shortcomings. The ﬁrst shortcoming of automation is that the more automated the process or step is, the less is the contact with the chemist or worker with it. This, in turn, results in a lack of continuous control over each situation and of discrete observations increasing the knowledge of the experimental events. This fact prevents the operator from obtaining a certain type of information, which in some cases might be even more interesting than that arranged to be obtained. In addition to this separation, the analyser or automated equipment requires more frequent check-ups and there is a greater risk of sample– result mismatching. The later can be avoided using bar codes. The popularity—sometimes magic—of replacing human effort and faculties in the realisation of any task negatively affects the attitude of laboratory principals. To make matters worse, manufacturers tend to overprice their products and usually make no mention of their limitations, the knowledge of which is as important as that of their potential. As a rule, potential purchasers do not have a good knowledge of the characteristics and possibilities of the vast range of commercially available automated equipment. The ease with which results are generated and the overconﬁdence in them can also lead to non-critical evaluation of the results, the sole responsibility of the chemist. 22.6

FUTURE PROSPECTS

Where automation of SP will go in the future can be anticipated by the foreseeable evolution of the necessities in the analytical laboratory, namely: 1. Shortening of the time required for automated SP, as required by the growing number of samples to be analysed. This reduction in time can be achieved by the design and commercialisation of both, general and dedicated devices assisted by different sources of energy. 2. Integration of several steps involved in common SP procedures in a single commercial device. 3. Broadening of robotics implementation—either workstations or robotic stations, as required—for unattended development of routine analyses. 4. Extension of laser uses, which will encompass improvement of present uses and new applications to the three physical states of samples, but particularly to solid samples. 5. Miniaturisation of automated SP devices is a present trend which will grow in the future as a result of the continuously smaller volumes required in the subsequent steps of the analytical process.

677

M.D. Castro and J.L. Luque

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36

678

M. Valca´rcel and M.D. Luque de Castro, Flow Injection Analysis: Principles and Applications. Ellis Horwood, Chichester, 1987. M. Valca´rcel and M.D. Luque de Castro, Automatic Methods of Analysis. Elsevier, Amsterdam, 1988. P.B. Stockwell, Automatic Chemical Analysis. Taylor Francis, Ltd, London, 1996. M.D. Luque de Castro and J.L. Luque-Garcı´a, Acceleration and Automation of Solid Sample Treatment. Elsevier, Amsterdam, 2002. R.E. Majors, LC –GC, 13 (1995) 742. L.T. Skeegs, Am. J. Clin. Pathol., 28 (1957) 311. J. Ruzicka and E.H. Hansen, Flow Injection Analysis, 2nd edn. Wiley Interscience, Canada, 1988. D.A. Burns, Anal. Chem., 53 (1981) 1403A. V. Cerda`, A. Cerda`, A. Cladera, M.T. Oms, F. Mas, E. Go´mez, F. Bauza, M. Miro´, R. Forteza and J.M. Estela, Trends Anal. Chem., 20 (2001) 407. B. Karlberg and G.E. Pacey, Flow Injection Analysis. A Practical Guide. Elsevier, Amsterdam, 1989. K.K. Stewart, G.R. Beecher and P.E. Hare, Fed. Proc. Am. Soc. Biochem., 33 (1974) 1429. J. Ruzicka and E.H. Hansen, Anal. Chim. Acta, 78 (1975) 145. P. Liu, Q.S. Pu, Q.Y. Sun and Z.X. Su, Fresenius J. Anal. Chem., 366 (2000) 816. J.R. Baena, S. Ca´rdenas, M. Gallego and M. Valca´rcel, Anal. Chem., 72 (2000) 1510. K. Benkhedda, H.G. Infante, E. Ivanova and F.C. Adams, J. Anal. At. Spectrom., 15 (2000) 1349. S.C. Nielsen and E.H. Hansen, Anal. Chim. Acta, 422 (2000) 47. E.P. Achterberg, C.B. Braungardt, R.C. Sandford and P.J. Worsfold, Anal. Chim. Acta, 440 (2001) 27. R.Y. Wang, J.A. Jarratt, P.J. Keay, J.J. Hawkes and W.T. Coakley, Talanta, 52 (2000) 129. P. Liu, Q.S. Pu, Z. Hu and Z.X. Su, Analyst, 125 (2000) 1205. J. Ruzicka and G.D. Marshall, Anal. Chim. Acta, 237 (1990) 329. J. Ruzicka, Anal. Chim. Acta, 261 (1992) 3. A. Cladera, E. Go´mez, J.M. Estela and V. Cerda`, Talanta, 43 (1996) 1667. R.C. de Campos Costa and A.N. Araujo, Anal. Chim. Acta, 438 (2001) 227. C.X. Galhardo and J.C. Masini, Anal. Chim. Acta, 438 (2001) 39. J.H. Wang and E.H. Hansen, Anal. Chim. Acta, 435 (2001) 331. W.W. Kubiak, R.M. Latonen and A. Ivaska, Talanta, 53 (2001) 1211. J. Ruzicka, Analyst, 125 (2000) 1053. Dionex, Application Note 73, Jul 2000, p.5 Dionex, Application Note 72, Jul 2000, p.4 Dionex, Application Note 76, Jul 2000, p.4 E.P. Achterberg and C. Braungardt, Anal. Chim. Acta, 400 (1999) 381. E.P. Achterberg and C.M.G. van der Berg, Anal. Chim. Acta, 284 (1994) 463. Manipulating Industrial Robots: Vocabulary, International Organization for Standardization, ISO/TR 8373, 1988. D. Nitzan, Development of Intelligent Robots: Achievements and Issues, IEEE J. Rob. Autom., RA, 1 (1985) 3. H. Waldman, Dictionary of Robotics. MacMillan Pub. Co, New York, 1985. A. Velasco-Arjona and M.D. Luque de Castro, Anal. Chim. Acta, 333 (1996) 205.

Automation of sample preparation 37 38 39 40 41 42 43 44 45 46

47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77

P. Torres, E. Ballesteros and M.D. Luque de Castro, Anal. Chim. Acta, 308 (1995). B. Ciommer and N. Muller, GIT Fachz. Lab., 40 (1996) 389. C. Shumate, E. Mardis, L. Weinstock and D. Bruce, Lab. Rob. Autom., 7 (1995) 73. T. Londo, P. Lynch, T. Kehoe, M. Meys and N. Gordong, J. Chromatogr., 798 (1998) 73. S. Dunkerley and M.J. Adams, Lab. Autom. Inf. Manage., 33 (1997) 93. S. Graves, B. Holaman and R.A. Felder, Clin. Chem., 46 (2000) 772. J. Steinle, T.H. Connolly and F. Pteiffer, Lab. Rob. Autom., 6 (1994) 313. R. Buhlmann and J. Gil, J. Chromatogr. Sci., 32 (1994) 243. Automated Environmental Lab Combines Robotics, Software, Chemical and Engineering News, 14 Aug, 1996. R. Schafer, C. Schulz and O. Stuckert, 4th International Conference on Automation, Robotics and Artiﬁcial Intelligence Applied to Analytical Chemistry and Laboratory Medicine, Montreux, Feb 1995. N. Beasley, Lab. Equip. Dig., 33 (1995) 19. M. Entzeroth, Lab. Autom. Inf. Manage., 33 (1997) 87. J. Marklew, Lab. Update, Jul–Aug (2000) 16. J. Norris and L. Ross, Anal. Proc., 30 (1993) 164. J.D. Norris, B. Preston and L.M. Ross, Analyst, 117 (1992) 3. C.D. Herold, K. Andree and R.A. Felder, Clin. Chem., 39 (1993) 143. J.L. Chadwick, Lab. Rob. Autom., 9 (1997) 21. S.E. Eckert-Tilotta, T.L. Isenhour and J.C. Marshall, Anal. Chim. Acta, 254 (1991) 215. A. Velasco-Arjona and M.D. Luque de Castro, Analyst, 123 (1998) 1867. A. Velasco-Arjona and M.D. Luque de Castro, Fresenius J. Anal. Chem., 363 (1999) 311. A. Velasco-Arjona, J.J. Manclu´s, A. Montoya and M.D. Luque de Castro, Talanta, 45 (1997) 371. L. Pieta, Adv. Lab. Autom. Rob., 7 (1991) 303. M.D. Luque de Castro and P. Torres, Trends Anal. Chem., 14 (1995) 492. L.A. Gifford, C. Wright and J. Gilbert, Food Addit. Contam., 7 (1990) 829. J.E. Coutant, P.A. Westmark and R.A. Okerholm, J. Chromatogr., 108 (1991) 139. J.E. Franc and K.A. Pittman, J. Chromatogr., 108 (1991) 129. L.A. Brunner and R.C. Luders, J. Chromatogr. Sci., 29 (1991) 287. K. Banno and R. Takahashi, Anal. Sci., 7 (1991) 571. C.N.C. Meredith and T.J. White, Anal. Proc., 27 (1990) 182. J. Hempenius, J. Wieling and D.A. Doornbos, J. Pharm. Biomed. Anal., 8 (1990) 313. D. Crahan, Anal. Proc., 27 (1990) 23. T.L. Lloyd, S.K. Gupta and J.R. Alianti, J. Chromatogr., 678 (1996) 261. T.L. Lloyd, T.B. Perschy and J.J. Tomlinson, Biomed. Chromatogr., 6 (1992) 311. J. Hempenius, G. Hendriks and C.C. Lin, J. Pharm. Biomed. Anal., 12 (1994) 1443. K. Pakulniewicz and J. Town, LC –GC, 13 (1995) 116. J.Y.K. Hsieh, C. Lin and M.R. Dobrinska, J. Pharm. Biomed. Anal., 12 (1994) 1555. E.I. Minder and D.J. Vonderschmitt, Chemom. Intell. Lab. Syst., 17 (1992) 119. Y. Tsina and B. Wong, J. Chromatogr., 675 (1996) 119. L. Ga´miz-Gracia, A. Velasco-Arjona and M.D. Luque de Castro, Analyst, 124 (1999) 801. M.M. Walstead, E. Yapchanyk and Y. Haroon, J. AOAC Int., 81 (1998) 9312. W.C. Koskinen, L.J. Jarvis and D.D. Buhler, Soil Sci. Soc. Am. J., 55 (1991) 551.

679

M.D. Castro and J.L. Luque 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98

680

A. Millier and G. Wallet, Chemom. Intell. Lab. Syst., 17 (1992) 153. P. Torres and M.D. Luque de Castro, Instrum. Sci. Technol., 24 (1996) 57. J.R. Ormand, D.A. McNett and M.J. Barttels, J. Anal. Toxicol., 23 (1999) 35. R.W. Taylor and S.D. Le, J. Anal. Toxicol., 15 (1991) 276. M. Arhoun, J.L. Martı´n-Herrera, E. Barreno and G. Ramis-Ramos, Lab. Rob. Autom., 11 (1999) 121. J.A. Garcı´a-Mesa, M.D. Luque de Castro and M. Valca´rcel, J. Flow Injection Anal., 10 (1993) 262. M. Valca´rcel and M.D. Luque de Castro, Non-Chromatographic Continuous Separation Techniques. Royal Society of Chemistry, Cambridge, 1991. M.P. Granchi, J.A. Biggerstaff and P. Grey, Spectrochim. Acta, 42 (1987) 169. M.S. Jime´nez and J.R. Castillo, J. Anal. At. Spectrom., 12 (1997) 1397. A. Rı´os, M.D. Luque de Castro and M. Valca´rcel, Anal. Chem., 58 (1986) 663. B.P. Ram, P. Singh and D. Allison, J. Assoc. Off. Anal. Chem., 74 (1991) 43. J.C. Boyd, R.A. Felder and J. Savory, Clin. Chem., 42 (1996) 1901. R.A. Felder, J.C. Boyd, K.S. Margrey, W. Holman, J. Roberts and J. Savory, Anal. Chem., 63 (1991) 741A. G.L. Hawk and J. Strimaitis, Advances in Laboratory Automation Robotics. Zymark Co, Hopkinton, MA, USA, 1984–1991. S. Unger, Biotech. Lab. Int., 5 (2000) 6. P. Torres, J.A. Garcı´a-Mesa and M.D. Luque de Castro, Fresenius J. Anal. Chem., 346 (1993) 704. P. Torres, J.A. Garcı´a-Mesa and M.D. Luque de Castro, J. Autom. Chem., 16 (1994) 183. P. Torres, J.A. Garcı´a-Mesa, M.D. Luque de Castro and M. Valca´rcel, Fresenius J. Anal. 346 (1993) 704. M.F. Joyce, P.A. Langan and W.S. Reid, Commun. Soil Sci. Plant Anal., 26 (1995) 19. P. Torres, J.A. Garcı´a-Mesa and M.D. Luque de Castro, Lab. Rob. Autom., 6 (1994) 233. J.A. Garcı´a-Mesa, M.D. Luque de Castro and M. Valca´rcel, Lab. Rob. Autom., 5 (1993) 29.

This page intentionally left blank

SECTION - 2

Chapter 23

Sample preparation for crude oil, petroleum products and polymers Robert I. Botto

23.1

INTRODUCTION

I was standing in Joe Brenner’s laboratory at the Israel Geological Survey in Jerusalem. I was curious and excited about the crude oil analysis project Joe had described prior to my visit. Now Joe handed me the pint bottle containing the actual sample to be analyzed. “This is the crude oil we need a bulk composition of.” “Be careful not to open it except under the hood.” “It’s pretty foul.” I turned the bottle over in my hands. “You need a bulk composition of what?” “I see water droplets clinging to the side of the bottle, some water on top, a gray layer that looks like an emulsion, a black, oily sludge,…and the bottom of the bottle is covered with ﬁne dark solids.” “Which part of this is the sample?” “All of it” replied Joe. “We need a bulk composition—that means we need to analyze the entire sample to reﬂect the total production from that well.” “Joe we have at least three phases here, well, four if you count the vapor above the liquid.” “I don’t see how can this can even be representatively sampled let alone analyzed as a bulk material!” “We need to devise a special sample preparation procedure” Joe replied.“I agree Joe” “Lets think about this…” 23.1.1 Nature of petroleum crude, products and polymers What oil producers call crude oil is a highly complex material. Wellhead crudes often contain several distinct phases that may or may not be visible to the beholder. Crudes often consist of natural materials from the reservoir mixed Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

683

R.I. Botto

with man-made materials such as oil ﬁeld chemicals used to assist in production and drilling muds. The natural part of the crude usually consists of organic compounds in liquid or semi-liquid form (although some very heavy crudes can be solids at ambient temperatures), brine and solids. The brine usually represents aqueous reservoir ﬂuid produced with the crude but it may represent ﬂuid injected into the oilﬁeld to enhance production. It may be in droplets associated with the oil or emulsiﬁed with it. The solids may represent organic materials that have come out of solution from the oil or inorganic materials from the reservoir (or both). Most crudes contain dissolved inorganic gases such as hydrogen sulﬁde and carbon dioxide, organic sulfur and nitrogen compounds, some volatile enough to produce intense odor. Elemental sulfur and elemental mercury have been found in signiﬁcant quantities in some crudes. Crudes containing low quantities of sulfur and sulfur compounds are termed “sweet”. Crudes containing high levels of sulfur compounds, especially hydrogen sulﬁde and mercaptans, are termed “sour”. The molecular weight range, physical properties and volatility of the organic portion of the crude classiﬁes it roughly as “light”, “medium” or “heavy”. The organic portion contains mainly hydrocarbons (which may be further broken down as to type) but will also contain compounds of oxygen, nitrogen, sulfur and the halogens. Reﬁner’s terminology can be confusing as approved organic nomenclature is generally replaced by a profusion of historic terms that have become the language of the industry (See Table 23.1). Terms related to what is done to the crude after it leaves the production ﬁeld constitute the language of petroleum reﬁning. Table 23.2 contains terms for a progression of reﬁnery processes. In essence, one ignores the IUPAC chemical dictionary when talking to a reﬁnery corrosion engineer about atmospheric pipestill overhead corrosion caused by a light gas oil cut of Arab heavy that contains high levels of naphthenic acids and organic chlorides! Even simple hydrocarbons are referred to by their traditional names—ethylene, butylene, isoamylene, isoprene, etc. Modern reﬁneries produce a diverse product slate. Direct products from reﬁneries include fuel gas, liqueﬁed petroleum gas, motor gasoline, diesel fuel, aviation fuel, petroleum solvents, lubricating oil basestocks, fuel oil, asphalt, sulfur and petroleum coke. Most reﬁneries are set up to supply feedstocks to chemical plants for manufacturing of bulk petrochemicals, specialty chemicals and polymers. The product slate from the modern petrochemical complex, such as the ExxonMobil Baytown Complex, is extremely diverse—everything from pure bulk hydrocarbons, such as benzene, to highly specialized copolymer products for the packaging and rubber industries. Gary and Handwerk [1] have published a good overview of the petrochemical industry. Over the past 27 years, the author has analyzed almost every type of fossil fuel [2], reﬁnery and petrochemical product (methane to polymers) together with the materials of the equipment used in production, the process chemicals and catalysts and, of course, the unwanted byproducts of production and

684

Sample preparation for crude oil, petroleum products and polymers TABLE 23.1 Petrochemical terms related to composition or physical properties Term

Meaning

Alkylate

High octane product from alkylation reaction used for gasoline blending Residual material from crude distillation containing high molecular weight hydrocarbons together with polar compounds containing oxygen and nitrogen referred to as “asphaltenes” Residue after distillation Synthetic rubber made from isobutylene and isoprene, trademark A segregated fraction of a distillation Several grades of fuel blended from virgin gas oil and various distillates An alkene having two double bonds such as butadiene Crude oil or fraction to be “charged” to a unit for processing Products in storage A petroleum liquid heavier than naphtha having an initial boiling point .4008F A reﬁned petroleum naphtha for motor fuel The highest boiling fraction of a distillation A highly aromatic product of the hydroforming process Initial boiling point—the ﬁrst drop of distillate A reﬁned petroleum distillate boiling 375 –5758F and having a ﬂash point .1158F The lower boiling hydrocarbons—normally those boiling at or below ambient temperature Liquiﬁed petroleum gases—normally C3 –C4 hydrocarbons Petroleum fractions suitable for making lubricants—heavier than gas oils generally Distillates between kerosene and lube stock fractions Light petroleum distillate boiling 60–4308F—may be “virgin” obtained from crude distillation or “thermal/cracked” in furnace process from heavier feed Cyclic saturated hydrocarbons (cycloalkanes) A value assigned to gasoline relative to its motor anti-knock properties—higher numbers have less knocking tendency. Scale is based on isooctane ¼ 100 and heptane ¼ 0

Asphalt

Bottoms Butyl Cut Diesel Dioleﬁn Feed stock Finished stock Gas oil Gasoline Heavy ends or heavies Hydroformate I.B.P. Kerosene Light ends LPG Lube stock Middle distillates Naphtha

Naphthenes Octane number

continued

685

R.I. Botto

TABLE 23.1 (continuation) Term

Meaning

Oleﬁn Overhead Parafﬁn Rafﬁnate

Unsaturated hydrocarbon Product removed in distillation as vapor Alkane In solvent reﬁning the portion of oil undissolved and unremoved by the solvent Product of thermal or catalytic cracking with improved volatility and octane quality Petroleum resid blended with lighter oil for fuel Heavy oil left after crude distillation—heaviest components of the crude Hydrocarbons formed naturally and not by processing Virgin naphtha from oil ﬁeld—sometimes called natural gasoline

Reformate Residual fuel oil Resid Virgin Wellhead condensate

manufacturing—corrosion scales, fouling deposits, sludges, wastes, etc. It would be impossible to characterize all of these sample types in this chapter. Instead, the precursor crude oil and the mainly hydrocarbon products made from crude will be highlighted, beginning by considering the elemental species in these materials. 23.1.2 Element context and species in petroleum crude, products and polymers Crude oil no doubt contains all of the natural elements at some concentration. Because it is a complex mixture, the elements it contains are distributed among the various phases that make up the whole. In addition to what is natural, contaminants have often been added in the drilling, production and transportation process. Table 23.3 is an attempt to model the composition of a crude oil on an elemental basis, indicating their distribution between three phases: organic liquid, solid and aqueous liquid (brine). Some crudes may contain organic solids composed of colloidal dispersed natural polymers or asphaltenes. These are likely to be rich in nickel, vanadium and other metals. Only a little progress has been made in identifying speciﬁc organic molecular species hosting various elements in crudes. It is well known that Ni and V are hosted mainly by porphyrin structures. Metal porphyrins are supposed to have been derived from the original chlorophyll and hemoglobin structures contained in plants and animals decomposed in the paleoenvironment of the source formation [4]. Relic molecules such as these are known as biological markers. Other metals may be hosted by porphyrins or chelated by

686

TABLE 23.2 Petrochemical processes and trace element sensitivity Description

Contaminant or trace element sensitivity

Resultant problem

Desalting

Heated crude is mixed with water and separated to remove salts Desalted crude is distilled producing fuel gases, naphtha, gasoline, kerosene, gas oil, lube and heavy bottoms

Surfactants from oil ﬁeld production Halides (inefﬁcient desalting or halohydrocarbons from oil ﬁeld chemicals or slop), sour gases, oxygen (from crude or ingress) Halides, sour gases, oxygen (from ingress)

Emulsion formation

Atmospheric distillation

Vacuum distillation Thermal cracking (Visbreaking) Catalytic (“Cat”) cracking Coking Hydrotreating/ Hydroprocessing/ Hydrodesulfurization

Heavier atmospheric fractions are distilled at low pressure to increase volatilization and separation Heat and pressure are used to break large hydrocarbon molecules into smaller ones Heavy oil is reacted on a ﬂuidized bed of silica/alumina catalyst in hot pressurized system to produce lighter products Heavy oils are thermally treated to produce light products and petroleum coke Catalytic hydrogenation to remove sulfur, nitrogen, oleﬁns, upgrade quality and stability

Overhead corrosion, Tray fouling (salts)

Overhead corrosion

Sour gas products Corrosion, fouling, coking including HCN, halides, metals Ni, Fe, V, sour gas Catalyst deactivation, products (including HCN) corrosion Metals, sour gas products Possible utilization/disposal issues, corrosion Metals, nitrogen, sour Catalyst deactivation, gas products corrosion

continued

Sample preparation for crude oil, petroleum products and polymers

Process

687

688

TABLE 23.2 (continuation) Description

Contaminant or trace element sensitivity

Resultant problem

Alkylation

Produces high octane blend stock by reacting isoparafﬁns and oleﬁns with acid (H2SO4 or HF) catalyst Molecular rearrangement over a noble metal catalyst for octane improvement—conversion of normal parafﬁns to branched Conversion of low octane naphthas to high octane products by aromatization over noble metal catalyst Solvent extraction to remove aromatics from lube feed stocks Removal of sour, acid gases from process gas stream by scrubbing with amine which is regenerated by ﬂash distillation Caustic (NaOH) wash to remove H2S and mercaptans Removal of parafﬁn wax from lube base stocks to improve low temperature pour and cloud points—may use ketone solvent or propane and dewaxing aids or selective hydrocracking Liquid propane is blended with oil from which asphaltenes precipitate and are separated

Water

Corrosion

Sulfur, water, oxygen

Catalyst deactivation, corrosion

Sulfur, nitrogen, arsenic, metals

Catalyst deactivation

Sour gases, oxygen HCN, heat stable amine salts

Solvent degradation, corrosion Cyanide corrosion, loss of amine

Ca, salts buildup

Fouling, loss of caustic

Metals (hydrocracking catalyst)

Catalyst deactivation

None

Ni, V content of deasphalted oil used to monitor process efﬁciency

Isomerization

Catalytic reforming Lube extraction Sour gas treatment

Caustic sweetening Dewaxing

Deasphalting

R.I. Botto

Process

Process

Description

Contaminant or trace element sensitivity

Resultant problem

Sulfur recovery

Sulfur plant produces elemental sulfur from H2S from sour gas treatment—example Claus process Major process for producing ethylene from naphthas, oils by reaction with steam in high temperature furnace

Salts

High ash content

Steam cracking

Catalytic polymerization

Chlorides, Al, As, Pb, Hg, P, Na, K, V, Zn even at part per billion levels Process for making polybutene for butyl rubber, Some polymer processes polyethylene, polypropylene, etc. are highly sensitive to metals, oxygenates, sulfur, halides, etc.

Furnace tube coking/corrosion and corrosion of downstream equipment [3] Catalyst deactivation, product malformation, contamination

Sample preparation for crude oil, petroleum products and polymers

TABLE 23.2 (continuation)

689

R.I. Botto TABLE 23.3 Element speciation in crude oil model Element(s) or group

Organic phase

Solids

Aqueous (brine) phase

H, C

Hydrocarbon, etc.

Carbonates, soot or carbon

N

Amines, amides, pyridines, quinolines, pyrroles, indoles, nitriles, porphyrins, etc. Acids, alcohols, aldehydes, ketones, ethers, esters, peroxides, etc.

Colloidal asphaltenes

Water, CO2, carbonates, organic acids Nitrate, cyanate

O

Halides

Alkalis Alkaline earths Al Si P S

Organic halides, freons (some may be from production additives) Possible chelates (?) Possible chelates (?)

Silicone contamination (defoaming agents, etc.) Phosphines, phosphates Elemental, H2S, mercaptans, disulﬁdes, alkyl/aryl sulﬁdes, thiophenes, etc. Chelates Porphyrins, chelates Porphyrins, chelates

Halide anions

Solid salts Solid salts

Cations in solution Cations in solution

Solid salts Silica

Possible silicates (?)

Phosphates Metal sulﬁdes and sulfates

Ti V, Ni Cr, Fe, Mn, Co, Ni, Cu, Zn Ge, Mo Carboxylates As, Sb Arsines, stibines Se Organic selenides, etc.

Oxide (contaminant) Asphaltenes Asphaltenes, corrosion product

Hg

Asphaltenes, sulﬁdes

Pb

690

Porphyrins, chelates, alkyls, elemental Chelates, alkyls from contamination

Water, CO2, SO2, organic acids, oxyanions

Carbonates, metal oxide/ hydroxides, silica, etc. Solid halide salts

Phosphate Sulﬁde, polysulﬁde, sulﬁte, sulfate, thiosulfate, thiocyanate

Selenate, selenite anions Halides

Sample preparation for crude oil, petroleum products and polymers

various polar ligands. Arsenic and Sb appear to be present partly as low molecular weight alkyl or aryl arsines and stibines, whereas Mo and Ge are present as carboxylic acid salts [5]. Crude contamination can obscure the origin of certain elements in the crude or in the products resulting from it. Halohydrocarbons are sometimes used in crude production and, together with surfactants and defoaming agents, can contaminate crude. Transport and storage of crude involves pipelines, shipping and tankage. Materials of construction can corrode and contaminate crude with corrosion products such as iron sulﬁdes and oxides. Corrosion inhibitors may also have been used to protect crude handling equipment. Pump oils, grease and oil additive packages containing various metals can leak into crude from pump seals and other equipment. Water and oxygen leaking into storage containers tend to oxidize some components of crude over time. Hydrogen sulﬁde in crude may begin to oxidize to elemental sulfur and acids that aggravate corrosion. “Rob we need to consider ﬁrst of all what we have here”, Joe said. “Look closely here.” “We have a heavy crude oil ﬁrst of all.” “The water phase is probably some brine from the formation.” “In addition, we have colloidal solids that may carry metals we will be interested in measuring—it’s probably a natural part of the crude…” “But some of those solids might be corrosion products from production, Joe.” “True”, Joe replied. “This crude is really sour!” “It’s loaded with H2S and mercaptans.” “It’s bound to corrode any steel it comes in contact with.” “We may have oilﬁeld chemicals like corrosion inhibitors in there too.” “Yeah it smells right through all this packaging!” “Yuck!” “So we have oil, solids, brine, possibly corrosion contaminants and oil ﬁeld chemicals plus dissolved noxious gases—and we need to analyze all of it.” “Basically, yes” “We better put this stuff back in the hood while do some more thinking Joe.”

23.1.3 Sample preparation challenges for trace element analysis Performing accurate, reliable trace element analysis is difﬁcult for any sample material. The usual hurdles must be overcome—representative sampling, sample handling and preparation without loss or contamination, precise and accurate analyte measurement free from bias and the accurate reporting of the analytical results with relevant quality information. Sample preparation for petroleum crudes and products offers some special challenges in all of these areas and in the safe handling of potentially ﬂammable, toxic materials.

691

R.I. Botto

Representative sampling for petroleum crudes and products is not always straightforward. The material sampled may be stored in large tanks, pressurized cylinders, barrels or even holds of ships. Often the sample must come directly from an operating unit through a sampling point; a ﬂuid access system built into the unit. Crudes and some liquid products may be multiphase materials, as noted above. In storage they tend to separate and settle. Large tanks and barrels are not easy to homogenize! Tank draws may be available at several levels (top, middle, and bottom) but the samples obtained may be quite different. Solids and water may even predominate in samples drawn from tank bottoms. Corrosion products could be present as well. Samples taken in pressurized cylinders have vapor and liquid volumes. Sampling the liquid layer may not represent the entire sample. Bleeding the sample cylinder for “outage”, though usually necessary for safe handling, releases part of the sample leaving a somewhat non-representative residue. Material taken through a sampling point can be fractionated or contaminated by the sampling equipment. Representative sampling of petroleum products by manual and automatic methods is discussed in two widely referenced standards [6,7]. Even solid polymers are often difﬁcult to sample in a representative manner due to inhomogeneities from poorly dispersed additives or wear/corrosion contaminants. In plant/ﬁeld sampling it is too often the case that inappropriate or dirty containers are used to collect valuable materials for trace element analysis. It is recommended that appropriate, pre-cleaned sample containers be provided to those obtaining samples for trace element studies. In one memorable case, clean glass sample bottles were provided for a naphtha trace metal study but the analytical results revealed high levels of Pb and Sn in the naphtha samples. It was later determined that the persons collecting the samples found it more convenient to collect them in ﬁve gallon tin cans (with Pb base solder joints) and later pour them up into the clean bottles provided! A discussion of techniques for equipment/container cleanliness is included in Section 23.3 below. Trace element species in the sample can interact with the walls of the container and be removed or can even cause some of the container wall to be incorporated into the sample by corroding it. Mercury and sulfur are examples of reactive elements that can “disappear” from a sample by attaching to the container wall. If the vapor space above the sample contains air, the sample can be gradually oxidized, causing precipitation of resins carrying trace elements to the container walls. Sample cylinders and containers lined with inert substances such as ﬂuorocarbons or silica are available for special samples. Acid cleaned glass containers are usually best for liquids that do not develop signiﬁcant vapor pressure. Polyethylene or polypropylene bottles and bags are appropriate for solids. It should be noted that organic vapors and oxygen might be able to penetrate the walls of plastic containers and organic liquids might leach plasticizer from plastic containers. Volatile organic materials should not be sampled or stored in plastic bottles.

692

Sample preparation for crude oil, petroleum products and polymers

Sample preparation schemes for petroleum crudes and products must take into consideration the various potential forms of the analyte elements and, particularly, their volatility. Some elements may be partially lost upon opening the sample bottle, even before sample preparation has begun. Less volatile species can be lost in the various stages of destructive sample preparation, such as acid mineralization or furnace ashing. Various techniques are available for retaining potential volatile species when converting a petroleum sample into an aqueous or other acceptable form for analysis. The potential ﬂammability and toxicity of petroleum materials requires particular attention to protect the analyst. Chemical fume hoods rated for toxic materials should be used when handling volatile petroleum liquids along with appropriate personal protective equipment. Sources of ignition, especially electric motors such as stirring motors, should not be used in the presence of signiﬁcant quantities of petroleum liquids. Special safety considerations are included with the description of the sample preparation techniques in the following sections. 23.2

SAMPLE PREPARATION TECHNIQUES AND INSTRUMENTATION

23.2.1 Ashing techniques The most widely used sample preparation techniques for trace element analysis of petroleum and petrochemicals aim to destroy the organic sample matrix and render the trace elements without loss or contamination in aqueous solution. The obvious way to destroy the organic matrix is by combustion. Reducing petroleum samples to ash using an open ﬂame or a combustion furnace is the traditional method of obtaining an ash sample. The American Society for Testing and Materials (ASTM) maintains standard testing procedures for the petroleum industry. Nadkarni [8] has prepared a recent method summary guide. Procedures used to obtain a gravimetric ash content are of two types—dry and wet (sulfated) ashing. The ash so obtained is used to determine trace element content. 23.2.1.1 Dry ashing Open beaker furnace dry ashing In high temperature dry ashing, a petroleum or polymer sample is combusted in air, usually in an open container, and without the use of an ashing aid such as sulfuric acid to assist in coking or sequestering the trace metals. In dry ashing, samples containing volatile materials are ﬁrst “ﬂamed off” or ignited over an open gas ﬂame and the sample is allowed to burn off until a non-volatile residue remains. This residue is placed in a mufﬂe furnace for ﬁnal ashing at 700 –9008C [9,10]. Samples containing little or no volatile materials, such as coke, may be placed directly into the mufﬂe furnace without ﬁrst igniting them [11]. Ashing

693

R.I. Botto

temperatures of 10008C or more may be required for refractory or graphitic coke samples. A temperature of 1000 – 11008C is necessary to decompose all carbonates to oxides, although it is not necessary to do this for metal analysis. Ashing temperatures can be reduced to 500 or even 4008C to maximize recovery of potentially volatile elements. A dedicated chemical fume hood is needed to ignite petroleum samples over gas burners. The hood will become full of soot and will require frequent cleaning. The mufﬂe furnace will require an air bleed and should be kept free of dust by blowing it out with an air blast. The furnace itself will require an additional fume hood to exhaust heat and possible fumes. See also Chapter 7 for additional information on dry ashing procedures. Pure quartz beakers are preferred for dry ashing because of their high temperature tolerance and lack of contaminants. Cover glasses, if used, should also be made of quartz and positioned leaving a small gap to allow air to enter the beaker. Over time, even quartz equipment will become etched and should be discarded. Beakers may be removed from the furnace with tongs and placed in a glass desiccator for cooling or the furnace may be cooled off leaving the beakers inside. Complete ashing at 5408C, the temperature used in our laboratory for most dry ashing, requires several hours and is often convenient to perform overnight. The residue obtained is the high temperature ash (HTA). Microwave assisted dry ashing Microwave furnaces are available for dry ashing petroleum materials. As with conventional furnaces, the samples may have to be ignited and burned off before placing in the microwave furnace. Ceramic or quartz ﬁber crucibles are typically used for microwave ashing. The microwave furnace features a rapid heating rate and small amounts (up to a few grams) of material may be completely ashed in less than 1 h. As high temperatures similar to furnace ashing are employed, the residue obtained is an HTA. Low temperature dry ashing Low temperature ashing is accomplished in an apparatus that exposes the sample to a radio frequency excited oxygen plasma at low pressures. The technique has been used to obtain relatively unaltered mineral matter from coal [12]. Applicable sample types are heavier oils that will not evaporate readily, resids, coke and solid polymers. The sample container is typically a quartz boat. Long periods of time (up to several days) are required for complete ashing and the ash may have to be stirred several times to expose new surface. Because of the lengthy period required for complete low temperature ashing, this technique is not widely used. The residue obtained is the low temperature ash (LTA). 23.2.1.2 Sulfated (wet) ashing Sulfated (wet) ashing is the standard technique used to determine the total amount of metal organic additives in new lubricating oils by reducing them to

694

Sample preparation for crude oil, petroleum products and polymers

metal sulfates [13]. The chemistry and pitfalls of the technique for total additive measurement have been documented [14]. Metal sulfates are relatively non-volatile at furnace ashing temperatures, thus they are retained in the sulfated ash (SA). All types of crude oils and petroleum liquids ranging from volatile naphthas to resids and even solid polymers can be wet ashed. Sample size is normally scaled according to the expected level of metals in the sample, larger sample sizes for low metals content. A typical procedure for sulfated ashing of an oil sample follows from Ref. [2]: (1) Weigh up to 20 g of sample into a quartz beaker and add either 5 or 10 ml of high purity (trace metal grade) concentrated H2SO4 (5 ml for 0–5 g samples, 10 ml for 5–20 g samples). A method blank should be prepared using an identical amount of H2SO4. (2) Coke the sample rapidly under a fume hood on a hot plate, stirring constantly with a quartz rod to ensure good mixing and to prevent foaming over. A “high” hot plate (400 –5008C) is required for good coking. Continue stirring until SO3 fumes disappear and dry coke remains. The sample may ignite during this step; wear heat and acid-resistant gloves. (3) Place the beaker containing the stirring rod in a mufﬂe furnace for overnight ashing at 5408C. The residue thus obtained is the SA. 23.2.1.3 Ash dissolution The HTA, LTA and SA may be solubilized by digesting with acids to recover all but silica. Nitric and HCl are normally used, either admixed 1:3 as “aqua regia” or added separately mixed with water (generally 25% volume acid). The acid mixture is allowed to reﬂux in the beaker covered with a watch glass until solubilization of the ash is complete. Silica, if present, will remain as a ﬂocculent white material. Adding a few drops of HF will dissolve and remove small amounts of silica but HF can also cause sparingly soluble ﬂuorides to precipitate and, if added in too large an amount, can etch the beaker and any other glass the solution comes in contact with. Beware the presence of dark insoluble material that may indicate that ashing was not complete. The sample will require another calcination/dissolution in this case. Keep in mind that it is usually desirable to match the type and concentration of acids present in solutions to be analyzed with the type and concentration of acids in the calibration solutions and blanks to be used for instrumental analysis. Silica may be quantitatively determined from HTA or LTA by removing the ash or a portion of it and fusing it with lithium tetraborate or lithium metaborate [15]. The resultant melt is dissolved in dilute HCl. Chapter 9 contains detailed information on ash fusion/dissolution techniques. Alternatively, the ash can be solubilized using HF and boric acid using elevated pressure digestion, as described in Section 23.2.2.2 (oven type or microwave assisted) or at atmospheric pressure using the microwave oven, as in “Ash solubilization by atmospheric pressure microwave digestion”.

695

R.I. Botto

23.2.1.4 Element recovery from ashing techniques Studies on recovery of elements from petroleum bitumen and coal ashed at various temperatures have been performed and a few are summarized in Ref. [16]. The recovery of elements from dry and sulfated ashing of butyl rubber has been investigated by spiking samples of the rubber with commercially available organometallic standards in lube base oil. Determinations were made by inductively coupled plasma-atomic emission spectrometry (ICP-OES) and the results are shown in Table 23.4. Acceptable element recovery (. 70%) by dry or sulfated ashing was determined for the following elements: Ag, Al, Ba, Bi, Ca, Cu, Fe, Mg, Mn, Mo, Na, Ni, P and V. Acceptable recovery with either ashing technique can be inferred or has been reported for the following

TABLE 23.4 Element recovery for spiked butyl rubber from dry and sulfated ashing Element

Ag Al As B Ba Bi Ca Cd Cr Cu Fe Mg Mn Mo Na Ni P Pb Sb Sn Ti V Zn

696

Dry ashing, 100 g sample n¼3

Sulfated ashing, 10 g sample n¼5

% Recovery

% RSD

% Recovery

% RSD

85 101 12 100 88 101 102 33 87 72 100 105 95 96 96 100 90 34 56 82 92 96 96

8.1 3.5 22 6.6 1.9 5.4 10 20 3.4 2.6 15 5.9 7.9 3.5 3.7 13 2.0 53 5.0 4.3 4.7 6.9 6.9

88 91 16 Low 86 103 100 71 49 87 72 99 84 82 99 84 80 79 74 66 67 93 67

2.0 5.7 54 1.6 3.3 13 22 9.8 3.3 20 4.7 5.1 8.2 1.9 23 6.0 5.7 4.8 6.8 11 3.1 11

Sample preparation for crude oil, petroleum products and polymers

elements: Be, Co, Ge, K, Li, rare earth elements, Sc, Sr, W, Y and Zr. Elements with acceptable recoveries by dry ashing but unacceptable recoveries using sulfated ashing are the following: B, Cr, Sn, Ti and Zn (Table 23.4). Elements with acceptable recoveries by sulfated ashing but unacceptable recoveries using dry ashing are the following: Cd, Pb and Sb. Elements with unacceptable recoveries using either high temperature ashing method are: As (Table 23.4) and Hg (inferred). Phosphorous in some samples may occur as low molecular weight phosphines that may be lost during ashing. Elements not tested in this study but presumed to be partially or substantially lost during high temperature ashing are: halogens, N, S, Se, Te and Tl. As mentioned above, Si is retained during ashing but will not be recovered unless provision is made using ash fusion or HF digestion. Low temperature ashing is reported to recover the following elements: Al, As, Ba, Be, Ca, Cd, Ce, Co, Cr, Cs, Cu, Eu, Fe, Hf, K, La, Mg, Mn, Mo, Na, Ni, Pb, Rb, Sb, Sc, Se, Sr, Ta, Th, Tl, V, W, Zn and Zr [17]. The halogens, Hg, Sb, S, and Os are lost or partially lost. Si may be recovered from the ash using ash fusion or HF digestion in a ﬂuorocarbon polymer vessel. A few elements, notably Hg, S and the halogens, are not recoverable using any form of open container ashing procedure. Recovering these elements requires a closed container approach, such as oxygen bomb combustion, described in Section 23.2.3.1. “Joe, what do you say we just wet ash this nasty stuff. Wet ashing will destroy the organic part, retain the salts and digest the solids all in one step!” “We can use plenty of sample, giving us a low dilution factor for the trace elements.” “With your ICP…” “Rob you are forgetting that some of the most important trace elements we are interested in—As, Se and Hg, for example, are lost in high temperature ashing, even sulfated ashing.” “It was a good idea though.”

23.2.2 Acid mineralization techniques Acid mineralization (also referred to as wet oxidation) techniques use the oxidizing and dehydrating power of concentrated mineral acids to decompose the organic structures in petroleum materials and solubilize the inorganic residues. The following acids may be employed singly or in mixtures: nitric acid (HNO3), sulfuric acid (H2SO4), perchloric acid (HClO4), phosphoric acid (H3PO4) and hydrochloric acid (HCl). Hydrogen peroxide (H2O2) 30–35% may be added as a digestion aid. Sometimes an inorganic catalyst is used to speed digestion. Acid digestion must be thorough enough to completely mineralize the sample. Partial digestions often leave solutions containing high levels of soluble organic compounds such as organic acids. Soluble organics can seriously interfere with the measurement of the analyte elements

697

R.I. Botto

using spectrochemical methods. All acid mineralizations are potentially hazardous and can cause violent or explosive reactions. Comments on safety are included below. Acid mineralization techniques can be classiﬁed as open vessel or closed container (elevated pressure) methods. Chapter 6 contains more information on wet digestion methods. 23.2.2.1 Open vessel digestions Kjeldahl type and reﬂuxing digestions Kjeldahl digestion (since 1883) is the classic example of an open vessel acid mineralization. The technique has long been used to determine the nitrogen content of fuel oils, lube oils and crudes [18]. In Kjeldahl digestion, a 1– 1.5 g oil sample is attacked by boiling concentrated H2SO4 in the presence of potassium, mercury and copper salts, which act as catalysts to speed up the digestion. Complete digestion requires 1–2 h. Unfortunately, the catalyst salts are undesirable for trace metal analysis preparations and without them, digestions are lengthy and incomplete. Acid mineralization of petroleum without salt catalysis requires strong oxidizing acids and/or hydrogen peroxide. Acid mineralization is used extensively in the petroleum industry to determine elements partially or completely lost by ashing techniques, such as P, As, Hg and Se. The ASTM procedure for P in lubricating oils uses a 300 ml Kjeldahl apparatus to digest up to 2 g of oil [19]. The oil is heated to fumes with H2SO4 ﬁrst, then concentrated HNO3 is added dropwise or in small portions until the digest is straw yellow in color, indicating that most of the organic material has been destroyed. Care is taken not to allow the H2SO4 to evaporate completely as loss of P could occur. Final treatments using 30% H2O2 and HNO3 follow until the digest is colorless. All forms of P are retained in this digestion and the total P content of the digest may be determined by spectrophotometry or atomic spectroscopy. A detailed acid mineralization procedure for As determination using up to 10 g of petroleum crude or products was described by Hofstader [20]. The sample is gradually combined with concentrated H2SO4 and HNO3 in an open 600 ml beaker until fumes of nitrogen dioxide appear. The temperature of the digest is maintained close to 2008C and additional HNO3 is added to produce nitrogen dioxide fumes at a steady rate. Once the digest has reached a uniform dark brown color it is heated to evolve fumes of H2SO4. Additions of HNO3 are made and heating is continued until the straw yellow color is obtained. Water is added and evaporated to fumes of H2SO4 several times to minimize undigested organic residue. The ﬁnal treatment is made with 2 ml of HClO4 added to 25 ml of H2SO4 and held at 1508C for 1 h. (Higher temperature results in loss of the HClO4 by distillation of the water azeotrope at 2008C). This step is done behind a safety shield in a perchloric acid fume hood. The aqueous digest is then diluted for analysis using spectrophotometry or atomic spectroscopy. Acid mineralization can be accomplished in a reﬂux apparatus minimizing loss of reagents until oxidation is complete. Acid digestion of oils for Hg analysis

698

Sample preparation for crude oil, petroleum products and polymers

can be performed using H2SO4 and HNO3 only in a reﬂux apparatus [20]. This particular apparatus allows the volatiles formed during the oxidative digestion to be collected, where they can be returned to the digestion ﬂask periodically. Without H2O2 or HClO4 ﬁnish, the digestion does not completely destroy all organic matter so the digest is extracted with hexane and purged of residual hexane prior to analysis for Hg. Walker et al. [21] used a 300 ml Kjeldahl ﬂask ﬁtted to a condenser as a reﬂux apparatus for digesting 1 –2 g of oil with H2SO4, fuming HNO3 and ﬁnally HClO4. The digest was used for determination of trace Se content by atomic absorption. Safety is a key concern in all open vessel digestions. Chemical fume hoods must have adequate ﬂow to sweep away the copious fumes from digestions. The use of HClO4 requires particular care. HClO4 should only be added after a sample has been thoroughly digested with HNO3 and H2SO4, never at the start of a digestion! Whenever HClO4 is used for digestion, the laboratory fume hood used must be of the perchloric acid type that can be washed down with water on a regular basis to eliminate potentially explosive perchlorate deposits from forming in the top of the hood or in the ductwork [22]. Alternately, a perchloric acid collection system must be employed. Our laboratory abandoned the use of HClO4 for digestions many years ago because of the potential hazards. A safer substitute for ﬁnishing a digestion is 30 –35% H2O2. Even without HClO4, all acid mineralizations should be considered as having potential for violent reactions. Directions for the order and amount of sample and reagents added must be followed precisely. The analyst must wear proper personal protective equipment (ﬁreproof lab wear, acid proof gloves, safety glasses with side shields or goggles, and/or face shield) and be shielded from the sample by a hood sash or lab shield. Acid mineralizations can be very time consuming and can require constant operator attention. Automatic addition modules for reagents are available for digestion apparatus that make these procedures easier and safer. Microwave assisted open vessel digestions Acid mineralization of petroleum can be conducted in a microwave assisted digestion apparatus using cavity or focused microwave energy. The digestion vessels are not completely open but are maintained at atmospheric pressure. The focused apparatus, available from several vendors, somewhat resembles a Kjeldahl digestion rack. Two–six digestion vessels are seated in a focused microwave unit. Reagent addition is performed manually or automatically at speciﬁed intervals from the tops of the vessels located outside the microwave unit. Another version uses a microwave oven containing an assembly of two digestion vessels attached to an apparatus that allows cooling of the upper part of the vessels for reﬂux action and automatic reagent addition. Maximum sample sizes up to 10 g are claimed but oil digestions may have to be scaled down. A recent review of focused microwave assisted strategies for sample preparation is included as Ref. [23]. More information on microwave digestion techniques can be found in Chapter 8.

699

R.I. Botto

Digestion of 0.6 g samples of lubricating oil was performed in a focused microwave digestion apparatus using an initial addition of HNO3/H2SO4 followed by multiple additions of HNO3 and a ﬁnal treatment with H2O2 [24]. A total of 25 min was required for the entire digestion. The aim of the study was to evaluate the efﬁciency of the decomposition of the oil using a system with six digestion vessels, one magnetron, one waveguide and six slots to control the transfer of energy to each of the six vessels. The digestion efﬁciency was measured by residual carbon content in the digests which varied from 2.4 to 9.7%, depending on whether all of the cavities were used simultaneously or each one individually. Digestions were not totally complete in 25 min even with these small samples, as indicated by the residual carbon content of the digests. Residual carbon containing species could interfere with analytical measurements using ICP-MS, ICP-OES and other techniques. In another study, 2% residual carbon content was achieved with an 8 g sample of lube base oil in a single open focused unit [25]. More work needs to be done to determine what reagents and conditions are required to yield complete mineralization of oil using focused microwaves. Ash solubilization by atmospheric pressure microwave digestion. The microwave oven can be used to solubilize ash from high or low temperature ashing using disposable 50 ml polypropylene centrifuge tubes as the digestion vessel. A special carousel for holding 36 or 52 of these tubes is supplied by CEM Corporation. The plastic caps for the tubes as supplied by CEM are perforated for insertion of a temperature probe. The hole also serves as an outlet for gas pressure. The digestion of the ash is accomplished at sub-boiling conditions while the temperature of a reference tube containing the acid mixture is monitored. Below is the procedure used in our laboratory for siliceous ash materials, such as catalyst ﬁnes: (1) Weigh up to 0.2 g of ash (Warning! Sample must be completely ashed before using this procedure!) into a 50 ml centrifuge tube and add 3 ml concentrated HCl, 1 ml concentrated HNO3 and 2 ml concentrated HF. (2) Cap the tube and place it on the turntable. Continue to prepare and load other samples. Label each tube with a marker on the outside body. (3) Prepare a temperature reference tube containing 6 ml of concentrated HNO3. Fit this tube with a glass thermowell and place it in the inner ring of the turntable. (4) Install the turntable in the oven, placing the ﬁber optic temperature probe in the reference tube thermowell. (5) Program the oven to ramp the temperature to 1108C over a period of 45 min and hold this temperature for 15 min. Start the digestion. (6) Allow the temperature to cool below 408C, then, open each tube (including the temperature reference) and add 40 ml of 5% (wt.) boric acid solution. Replace the caps and reposition the tubes in the turntable. Reload the turntable in the oven and replace the temperature probe.

700

Sample preparation for crude oil, petroleum products and polymers

(7) Program the oven to ramp to 908C in 45 min and hold for 15 min. Start the program and allow the tubes to cool below 408C after it ﬁnishes. (8) Pour the contents of each tube into a 125 ml disposable polyethylene bottle and make up to 100 g with deionized water. Calculate individual sample dilution factors on ash or original sample basis. Solutions are now ready for elemental analysis. 23.2.2.2 Elevated pressure digestions Elevated pressure acid digestions can achieve higher temperatures and more complete mineralization of petroleum materials. Closed vessel digestions also have the advantage of retaining potentially volatile elements such as Hg, P and Se. Vessels for elevated pressure digestion are designed to operate up to some maximum temperature and pressure at which point there is a controlled release via a release mechanism or a potentially catastrophic failure. The problem with digestion of oils is that large quantities of CO2 and other gases are generated by the sample decomposition adding to the pressure in the vessel due to the acid vapor and elevated temperature. Small amounts of oils can create high vessel pressures when undergoing decomposition with oxidizing acids such as HNO3. With some organic materials and HNO3/H2SO4 it is also possible to synthesize explosive compounds in the vessel. Pressure and safety considerations severely limit sample size for most elevated pressure digestions of oil. Because of its unpredictability, HClO4 should never be used in elevated pressure digestions [26]. Oven/autoclave type digestions Teﬂon “bomb” digestions. Steel-jacketed PTFE lined acid digestion bombs have been widely used for digesting inorganic materials [26]. Our laboratory has used them for coal, oil shale and ashes from petroleum and polymers [2]. A range of capacities and maximum operating temperatures and pressures is represented in the current line from Parr Instrument Company, a major supplier. The bombs will accommodate up to 5 g of inorganic solid and 15 ml of acids in the largest model. A high strength model can withstand 2758C and pressures up to 5000 psig. Ashes from high temperature ashing of ﬁnished polymer products may contain inorganic “ﬁllers”, colorizing agents, ﬁre retardants and other additives that can be difﬁcult to dissolve on a hot plate or in a microwave assisted open vessel digestion. The following is a typical procedure for solubilizing a polymer ash sample: (1) Weigh 0.1 –0.2 g of ash sample into the Teﬂon liner of a 23 ml general purpose acid digestion bomb. Add 1 ml each of concentrated HCl, HNO3 and HF. Cap and assemble the bomb. (2) Place in preheated oven at 130 –1508C for 2 h. Remove and cool the bomb. (3) Open and add 1 g of boric acid and 5 ml deionized water. Reseal and place in oven at 130 –1508C for 1 h. Remove and cool the bomb.

701

R.I. Botto

(4) Wash contents into a polyethylene beaker and dilute to 100 g ﬁnal weight. Calculate dilution factor on ash basis (weight of solution/weight of ash) or original sample basis (weight of solution/(weight of ash aliquot) £ (weight of polymer/total ash weight)). Elevated pressure digestions for petroleum must be approached with caution as noted above. Sample size (0.1 g for 23 ml bomb) and acid loading should never exceed the manufacturer’s recommendations. Sample size severely limits utility of method for trace element analysis. Metal-jacketed PTFE bombs tend to corrode and the corrosion products may be a source of contamination. Pressure decomposition vessels without metal jackets are available but ability to retain a complete seal limits use temperature. All ﬂuorocarbon type containers are somewhat porous and may retain contamination from manufacture or from use. Extensive acid leaching and/or vessel annealing may be required to reduce contamination. A detailed discussion of the characteristics and use of elevated pressure digestion vessels is included in Ref. [16]. A recent study comparing closed elevated pressure digestions for residual fuel oil with open focused microwave found signiﬁcant amounts of residual carbon species in the former and essentially complete digestion in the latter. Unfortunately, the open focused microwave digestions exhibited poorer recovery for many of the elements studied, possibly through volatilization losses [27]. High pressure autoclave and ampoule digestions. High pressure autoclaves using quartz, or ﬂuorocarbon vessels have been used extensively for decomposition of refractory inorganic solids [16]. They could potentially be used for petroleum acid mineralization although they would probably not be able to accommodate more than a few tenths of a gram of material. Sealed quartz ampoules have been used to decompose extremely resistant materials due to the very high pressures achieved. Again, sample size is limited and so is the utility of such systems for petroleum trace element analysis. Microwave assisted pressure vessel digestions During the past 15 years, the laboratory microwave oven has evolved into a precision scientiﬁc tool for accomplishing a wide variety of elevated pressure sample decompositions for inorganic and organic materials [28]. The modern laboratory microwave oven with pressure and temperature control provides the ﬂexibility to optimize methods for petroleum and polymer samples of various types and sizes. Knowing temperature and pressure at all times helps to ensure complete digestion—vessels may also be vented at one or more stages in the digestion to release CO2 and other gases. See also Chapter 8 for additional information on microwave digestions. Moderate pressure microwave digestions. Moderate pressure microwave digestions are conducted at pressures below 500 psig and at temperatures up to

702

Sample preparation for crude oil, petroleum products and polymers

2008C. Most commercial laboratory microwave oven equipment works at moderate pressures, most often 150 –300 psig and temperatures of 120 –2008C. At moderate pressures, oils may or may not be completely digested in 30–60 min using HNO3 alone or with H2O2. Sample sizes used are typically 0.1 –0.5 g. Often, the limiting factor for sample size is the amount of gas evolved during digestion. Venting after an initial stage of digestion to remove accumulated gases may permit larger sample sizes. Undigested organic carbon present at the end of the procedure may or may not be evident to the eye. The material could appear as a yellowish waxy residue, particularly in the case of heavy oils. The undigested material may also be present as nearly invisible water and acid soluble species (organic acids for example). Undigested organics could cause a matrix problem in certain methods of elemental analysis such as inductively coupled plasma-mass spectrometry (ICP-MS) where molecular interferences might arise from carbon species. US Environmental Protection Agency (US EPA) Method 3052 [29] uses moderate pressure microwave digestion for “siliceous and organically based matrices” including oil contaminated soil and waste oil. The method digests up to 0.5 g of soil or up to 0.25 g of oil using 9 ml of HNO3 and 0.5 –5 ml of HF (scaled according to amount of siliceous material present) and, if needed for complete digestion, 0.1 –2 ml of H2O2. Vessel temperature is programmed to rise to 1808C in 5.5 min and hold at 180 ^ 58C for 9.5 min. The method claims near complete digestion of oil and oily soils and complete solubilization of most inorganic materials in soils. Element recoveries for ﬁve “wear metals” in oil were near 100%. A few have reported some success with moderate pressure microwave sample preparation for oils [30,31]. Our own laboratory trials were less than satisfactory. Attempted digestions of heavy fuel oil yielded incomplete digestions and were impacted by contamination from the materials of construction of the digestion vessels [32]. Cutting back on sample size to improve the digestion made it more difﬁcult to determine the trace elements due to dilution and contamination. High pressure microwave digestions. High pressure microwave digestions provide for more complete mineralization of oil and can potentially accommodate larger sample sizes. Commercial microwave digestion vessels are available to work at pressures up to about 1500 psig and temperatures up to 3008C. One manufacturer has published method application notes for digestion of wax, grease, oily ﬁltrate, butadiene rubber and reprocessed fuel oil [33]. Sample sizes range from 0.1 to 0.5 g and digestions are performed using HNO3 and H2O2. During the digestion of 0.3 g of wax vessel temperatures reach 2108C and pressures peak close to 50 bar (730 psig). Another manufacturer recommends organic sample sizes of 0.1 – 0.2 g for their high pressure microwave digestion vessels [34]. If vessels are not vented during digestion either intentionally or unintentionally, potentially volatile elements are retained. The determination

703

R.I. Botto

of trace As, Sb, Se and Hg in oily waste was accomplished using high pressure microwave digestion [35]. Precision for element determinations from multiple digestions ranged from 5.4 to 6.6% relative standard deviation and recoveries for the elements ranged from 89 to 105%. Another laboratory developed a nine step digestion program for residual fuel oil (NIST SRM 1634b) using HNO3 and H2O2 that resulted in complete digestion of 0.25 g and produced good agreement for certiﬁed values for eight trace elements determined by ICPMS [36]. New developments in microwave digestions. Present day problems and limitations with microwave digestions of petroleum materials may be soon laid aside by innovative new designs for digestion vessels and systems. Vessels that can vent and reseal at prescribed pressures will allow for release of the early digestion gases and allow for much larger sample sizes [37]. Microwave digestions that use UV light to enhance the decomposition rate of organic materials are just coming onto the market [38]. These have the potential of providing more complete digestions and more gentle conditions. “I had another thought Joe” “If we could just ash this goop inside some kind of closed system we could retain all of the volatile elements.” “Not a bad idea.” “We have the Teﬂon digestion bombs and our 500 psig microwave digestion vessels available to us” Joe replied. “The problem is that the amount of organic material we can digest safely is no more than a couple tenths of a gram.” “The sample dilution will be at least 200 fold making some trace determinations kind of iffy…” “Yeah and in my experience with heavy oils you will still have a yellow waxy residue after the digestion Joe” “Well keep thinking Rob Botto—as much as I hate to multiply work we might have to use more than one sample prep for all of the elements”

23.2.3 Oxygen combustion 23.2.3.1 Oxygen combustion bomb The oxygen combustion bomb is a useful tool for determining potentially volatile elements in petroleum products and polymers. Commonly used for calorimetric measurements for coal, the device is capable of combusting approximately 1 g of an organic material in a completely sealed environment under approximately 30 psig of pure oxygen [16]. The equipment used in many laboratories is manufactured by the Parr Instrument Company [39]. They make oxygen bombs especially for trace element analysis with quartz liners and Pt plated interior parts. Bombs completely lined with Pt/Ir alloy are also available.

704

Sample preparation for crude oil, petroleum products and polymers

A few milliliters of a solution of dilute HNO3 or HNO3/H2O2 are placed in the bottom of the combustion bomb to absorb the elements released during the combustion. After combustion, the bomb is disassembled and the solution removed with rinsing of the interior of the bomb. A dilution factor of approximately 50 is usually achieved. Elements commonly determined are As [2,40], Se [40,41], S [42,43], P, B, halogens, Pb and Hg [44 – 46]. Determination of nitrogen is also possible if air is carefully removed from the bomb before the pure oxygen is introduced [44]. An ASTM procedure uses sodium carbonate solution to absorb Cl from combustion of petroleum products [47]. Our laboratory has used the oxygen combustion bomb for the determination of As and Hg in butyl rubber. Analyses of spiked rubber revealed recoveries of 90 þ %. 23.2.3.2 Oxygen combustion vessel Oxygen combustion of petroleum samples may be done at atmospheric pressure in closed vessels or accomplished using a burner attached to an apparatus to condense and trap the analyte elements. The oxygen ﬂask method is the classic Schoeniger method (since 1955) [48]. A sample is placed in a ﬂask containing oxygen and an absorbing solution and ignited by electrical means or with infrared radiation. The gases are taken up into the solution by agitation of the ﬂask contents and the solution is used for determination of the elements. Sulfur and halogens were commonly determined using this method but it has been superseded by the oxygen bomb method that is safer and easier to operate. The burner (or lamp) combustion methods are still in use in some reﬁnery laboratories for sulfur determination in petroleum liquids. The sample is burned in a closed system using an oxygen/hydrogen burner (Wickbold type) or oxygen/CO2 atmosphere. Combustion products are absorbed and sulfur oxides are oxidized to sulfate for determination by gravimetry, titration or turbimetry [49,50]. Samples of LPG and even gases can be analyzed using this apparatus. Halogens and other elements can potentially be determined at very low concentrations because sample sizes can be reasonably large (20–40 g or more). Unfortunately, the apparatus is complex and hazardous to operate so it is disappearing from petroleum laboratories. “Joe do you have an oxygen combustion bomb like our Coal Laboratory in Baytown uses for heating value of coal and shale?” “I’m thinking that we could combust these samples in the bomb and recover all of the volatile elements from the washings…” “Great idea Rob but unfortunately we don’t mine a lot of coal around Jerusalem and we have been concerned with bombs of a rather different sort lately.” “Oh”

705

R.I. Botto

23.2.4 Sample component separations/extractive sample preparation Breaking down a petroleum sample into components may have advantages for analysis. Physical and chemical separations can be used to isolate and concentrate analytes of interest, avoid interferences and eliminate the unwanted organic matrix. Here are a few useful types of separations that could be performed. 23.2.4.1 Physical separations Distillation [51] is most common procedure for fractionating petroleum samples. Metals tend to concentrate in the “bottoms” fraction yielding a concentration technique for Fe, Ni, V, etc. Volatile forms of some elements (such as Hg) may concentrate in lower boiling fractions. Evaporation or weathering is another method of concentrating heavier components in a volatile sample. Asphaltenes may be precipitated from crudes or lube stocks by pentane dilution [52]. Asphaltenes contain the polar components of the oil and a large fraction of the metals, particularly the Ni and V porphyrins. Thus, we have a way of concentrating the metals rich fraction of the sample without distillation. Separation of broad compound types in oils having an initial boiling point can be accomplished using a clay –gel absorption column chromatographic method [53]. The method yields separate fractions containing the polar compounds (including the asphaltenes), aromatic compounds and saturate compounds. Sample size of the original oil is typically 5–10 g. The clay –gel preparative separation scheme can be used as a starting point for element speciation studies. 23.2.4.2 Aqueous extraction Aqueous extraction of oil may be used to separate inorganic ions (halides) or salts (sodium, calcium, etc.) Aqueous extraction may be used as part of a scheme to determine what portion of the total concentration of the element (such as Cl) is inorganic. Dilute aqueous acids or bases may be used to extract components such as sulfur and nitrogen species and organic acids from an oil sample. Oxidative extractions have been used to remove Hg compounds from oil [54]. Another application of oxidative extraction is the “de-metallisation” of used industrial oils for analysis as aqueous solutions [55]. 23.2.4.3 Organic solvent extraction Solvent extraction can be used to separate hydrocarbons of diverse type. For example, methyl ethyl ketone is used to extract non-parafﬁnic components from commercial wax production [56]. Samples that consist of mixtures of inorganic and organic components may be extracted in a Soxhlet type device to separate them. This is preferable to washing and ﬁltration if there is a tendency to form emulsions. Once a sample of sludge or scale has been “deoiled” in this manner it

706

Sample preparation for crude oil, petroleum products and polymers

can be characterized using X-ray diffraction and compositional techniques. The speed and efﬁciency of solvent extraction techniques have been increased with the use of microwave methods [57]. Additional information on solvent extraction techniques may be found in Chapters 11 and 15. 23.2.4.4 Solid phase extraction Solid phase extraction is being used to isolate and separate organic compounds from aqueous systems [58]. Solid phase extraction cartridges or disks that attract polar compounds from oils may also be used to concentrate the metals they carry to provide for more sensitive analysis by x-ray ﬂuorescence and other techniques. Samples may also be ashed and the metals concentrated on ion exchange disks for-ray ﬂuorescence. Non-aqueous ion exchange solid phase extraction can be used to isolate acids or bases from petroleum such as in the quantitative analysis of naphthenic acids in crude oils [59]. Chapters 13 and 14 contain much additional information on solid phase extraction techniques. 23.2.5 Organic sample dilutions and dissolutions 23.2.5.1 Sample/solvent compatibility “Dilute and shoot” seems the easiest way to analyze a petroleum sample. Dilution is the simplest form of sample preparation. Dilute in what? The solvent must be compatible with the analytical method (having the appropriate volatility, for example), free from contaminants or substances that could interfere and it must also solubilize the entire sample. Incomplete solubilization results in fractionation and potential analyte loss or analytical bias. It is also desirable that the solvent has low toxicity and that it does not attack the materials of construction of the sample introduction equipment used for analysis. One must also carefully consider the impact of solvent choice on the instrumental analysis. Matrix related effects caused by analyte/matrix volatility and/or differences in physical properties between samples, or from mismatch between samples and calibration standards may have a big impact on the accuracy of elemental analysis by direct dilution ICP-OES. Solvent choice may be a key to minimizing these effects [60]. Internal standards are routinely used to compensate for sample transport and matrix effects in organic ICP [61,62]. Direct dilution ICP-MS methods for determination of trace elements and elemental speciation in a variety of petroleum materials from light condensates and naphthas to tars have been published [63 –69]. Sample/solvent compatibility is the key issue in diluting petroleum samples and solvent polarity is the key variable that determines compatibility and solution stability. “Like dissolves like” is a good general rule. The solvent polarity index (Table 23.5) puts this concept on a quantitative basis. Keep in mind that ﬁnished oils may contain additive packages that include compounds that are

707

R.I. Botto TABLE 23.5 Solvent polarity index: a scale for sample/solvent compatibility Solvent Heptane Octane, decane Isoparafﬁns Kerosene Cyclohexane Decalin Carbon tetrachloride Toluene O-xylene Benzene Tetrahydronaphthalene Chlorobenzene Nitrobenzene Diethyl ether Methylene chloride 2-Propanol Tetrahydrofuran Chloroform Methyl isobutyl ketone Ethyl acetate Dioxane Methyl ethyl ketone Acetone Methanol Pyridine Dimethyl sulfoxide Water

Index value 0.1

0.2 1.6 2.4 2.5 2.7 2.7 2.8 3.1 4.0 4.0 4.1 4.2 4.4 4.8 4.8 5.1 5.1 5.3 7.2 10.2

Typical sample types Parafﬁn wax White spirit, mineral oil Diesel, jet fuel Petroleum distillates Petroleum ether, naphtha Gas oils Aromatic naphtha Lube base stocks Crude oils Fuel oils Heavy petroleum fractions Asphaltenes, tar Coal liquids, creosote Shale oils Jet engine oils (synthetic) Hydraulic ﬂuids Transmission ﬂuids Lube additives Coal, shale “bottoms”

much more polar than the base oils. High performance synthetic oils (such as jet engine lubes) may be ether based and have high polarity compared to petroleum lubes. Hydraulic and transmission ﬂuids also fall in this category. Metal-organic standards used for preparing solutions for petroleum analysis are available from many commercial laboratories. Many of these organometallic compounds are chelates having signiﬁcant polarity. Stabilizers are sold by most vendors of these standards to help keep them in solution after they are diluted to prepare working standards. Still, many of these compounds

708

Sample preparation for crude oil, petroleum products and polymers

will not be stable over long periods when diluted in solvents such as n-hexane or isooctane, even when using the stabilizer. Solvents like xylenes or tetralin will provide better long-term stability for metal-organic standards. Stability of standards should be veriﬁed by testing. In some cases, depending on the element of interest, samples may need to be stabilized. 23.2.5.2 Polymer dissolutions Polymer dissolutions can require considerable patience. High molecular weight polymers will dissolve very slowly and may not dissolve completely. Cross-linked polymer structures may not dissolve at all. Solvent selection follows “like dissolves like” but it should be kept in mind that although high-density polyethylene has a long chain parafﬁn type structure it is not likely to go easily into heptane! Polymer dissolutions are usually done in a reﬂux type apparatus allowing plenty of time for completion. Even if the polymer dissolves, the inorganic contaminants, catalyst residues, additives and ﬁllers may not. Extraction of soluble polymer away from insoluble components may be accomplished in a Soxhlet type apparatus. Microwave assisted polymer dissolutions claim increased speed and effectiveness [70]. “Joe if we could ﬁnd a solvent system that could dissolve the heavy oil would the solids be a problem?” “Could we aspirate this material right into your ICP?” “Don’t you have one of those high solids nebulizers?” “Whoa Rob!” “One question at a time.” “I suppose if we found a solvent to dissolve it our V-groove nebulizer could handle the solids…slurry nebulization and all that…” “THF” “It dissolves everything!” “Including my pump tubing and ﬁttings!” “Tetralin might be a better choice Rob.” “What about the water phase, Joe?” “We might end up with a slimey emulsion.” “Emulsion?” “Hummm…”

23.2.6 Stable emulsions 23.2.6.1 Emulsiﬁcation of crude oils and naphthas Use of stable emulsions for analysis of petrochemical materials is gaining popularity. By preparing an emulsion we convert an organic matrix to a more or less aqueous matrix, one that is compatible with mineral acids, bases and dissolved ionic salts. Emulsions then can be analyzed like aqueous solutions, spiked with aqueous standards for standard additions and aqueous internal standards.

709

R.I. Botto

Stable microemulsions can be prepared from crude oil [71–73]. An example preparation follows from Ref. [71]: (1) Weigh 0.5 g crude oil into a tared 60 ml glass bottle. Record weight to nearest 0.1 mg). (2) Add the equivalent weight of 1,2,3,4-tetrahydronaphthalene (tetalin) and mix until a homogeneous solution is obtained. (3) Add 1.0 g of Triton X-100 surfactant and mix thoroughly again until a homogeneous solution is obtained. Use a vortex mixer or ultrasonic bath for mechanical agitation. (4) Add deionized water stepwise in 1 ml increments with continual agitation in a vortex mixer until at least 5 ml have been added and a homogeneous gel has been prepared. Add internal standard spike, if used. This gel microemulsion may now be diluted in one step to a ﬁnal weight of 50.0 g. Tetralin serves to reduce the viscosity of the crude oil and help it to mix with the surfactant. Microemulsions prepared this way are reported to be transparent isotropic solutions stable for weeks or even months. Naphthas and gasoline can also be analyzed as an emulsion. Emulsiﬁcation with water helps to reduce the effects of the volatile hydrocarbons on an analytical method such as ICP-OES or ICP-MS. A method for determination of Pb in gasoline by ICP-OES was developed by Brenner et al. [74]. Emulsions containing 10% (wt.) gasoline in water and Y as an internal standard were treated with iodine and tricapryl methyl ammonium chloride to ﬁx Pb as a non-volatile iodide complex and analyzed using emulsiﬁed calibration standards prepared with decalin. Good accuracy was obtained with a stable ICP operation. Kumar and Gangadharan using a similar emulsion technique [75] accomplished the analysis of naphtha for V, Ni and As by ICP-MS. Recoveries in the 86 –101% range were obtained for several elements when spiked in organometallic form. 23.2.6.2 Acid emulsiﬁcation for analysis of wear metals in used lube oils Metals from engine wear are present as particles suspended in used lube oils. Engine oils are monitored for wear metals to assess engine wear. This is a particularly important activity in the commercial trucking, heavy vehicle, railroad, aircraft and ﬂeet bus/automobile industries and in the military where vehicle reliability means money and lives. Most wear particles are sub-micron size to a few microns but particles indicating catastrophic engine wear just prior to failure may be up to millimeter size [76]. Metal analysis techniques using solution nebulization do not have the capability to transport such large particles into the atomization device nor does the ﬂame or plasma have the ability to atomize them effectively. The wear particles must be solubilized and homogenized with the oil for accurate determination. This is accomplished by acid emulsiﬁcation,

710

Sample preparation for crude oil, petroleum products and polymers

otherwise know as the “particle size independent” method for wear metals analysis [77–81]. The following procedure for automotive lubes is from Ref. [81]: (1) Mix 1.0 g of used oil with 0.1 ml of concentrated HCl in a clean, dry tube. (2) Place in an ultrasonic bath for 10 min at 60–808C. (3) Add 0.1 ml of concentrated HNO3 and place in the ultrasonic bath for an additional 10 min. (4) Dilute to 10 ml with a 2:1 mixture of xylenes/butylcellosolve.

“Joe I can’t believe it!” “These measurements are actually precise!” “The microemulsion runs almost like an aqueous sample on the ICP-OES.” “We are seeing some carbon background from the organic material but not nearly as much as I would expect with pure organic ICP.” “Dilution factor not too bad…detection limits look good Rob!” “We can conﬁrm accuracy with aqueous standard addition.” “I can’t believe we prepped this three phase mess with one simple procedure—Joe that was brilliant!” “We should celebrate.” “How about a dip in the Dead Sea?” “I’ll take you to the Plasma Spectroscopist’s Ritual Bath in Ein Gedi.” “There we will cover our bodies with therapeutic black sulfur mud and bask in the 110 degree desert heat!” “Sounds like another black goopy sample problem to me Joe!” “I’m game!” 23.2.7 Scrubber sampling for C1 – C4 hydrocarbons and gases The analysis of fuel gas or C1 –C4 hydrocarbons for non-particulate trace metals requires special apparatus for trapping the elements of interest. Sampling is often done directly from the unit where a sampling point provides a controlled stream of gas. Alternatively, a liqueﬁed gas sample is collected in a steel sample cylinder. The sample point or cylinder sample is attached via a regulator and needle valve assembly to the ﬁrst of a series of gas scrubber or impinger vessels (scrubbers) in a train (Figure 23.1). The gas is fed through the scrubbers at a steady rate in which intimate gas/liquid contact is maintained without “slugging”. The more gas that is collected, the lower the concentration levels that can be determined. Reagents in the scrubbers trap the elements in the gas. A wet or dry test meter follows the scrubber train to measure the amount of gas sampled. The outlet side of the test meter is vented. Our laboratory uses the following scrubber train (250 ml plastic scrubbers) for general fuel gas sampling: Scrubber #1: 100 ml of deionized water

711

R.I. Botto

Fig. 23.1. Gas scrubber train for fuel gas sampling for trace element analysis.

Scrubber #2: Scrubber #3: Scrubber #4: Scrubber #5:

100 ml 100 ml 100 ml 100 ml

of 4% (vol.) HNO3 of 4% (vol.) HNO3 of 30 –35% H2O2 of 4% (vol.) HNO3

The ﬁrst scrubber would trap salts and particle contamination; the following scrubbers are for trapping metal carbonyls, metal-organic compounds, metal hydrides, etc. The 4% (vol.) HNO3 concentration matches our ICP-OES calibration solutions. For analysis, 4 ml of HNO3 is added to the ﬁrst scrubber solution. The H2O2 scrubber solution is digested with HNO3 to remove the H2O2 and it is made up to 4% (vol.) HNO3. The scrubber solutions are usually analyzed separately and the element concentrations summed to yield the total. Scrubbers #2–#5 can be combined and analyzed as one sample (after H2O2 removal), however. The contents of Scrubber #1 should not be combined with the other four in case there has been contamination of this solution from the sampling equipment (ﬁne dust from corrosion scale for example). After analysis, if it has been determined that there has been no gross contamination of Scrubber #1; the results can be added to those from the other scrubbers. Special reagents may be used to trap particular analytes, silver diethyldithiocarbamate solution for arsine, for example [82]. A solution of 3% KMnO4 in 10% H2SO4 has been found effective for trapping Hg [83]. Special techniques for the analysis of volatile hydrocarbons and petrochemicals by ICP-OES are found in three publications [32,84,85].

712

Sample preparation for crude oil, petroleum products and polymers

23.3

CLEANLINESS AND QUALITY ASSURANCE

23.3.1 Equipment cleaning Cleanliness is next to Godliness in the analytical laboratory. Many topics could be discussed relative to equipment cleaning, clean room facilities, ﬁlters and hoods. However, the intention here is to provide just a few pointers related to petroleum sample preparation. 23.3.1.1 Glassware How do you get that greasy goop off the beakers? The days of throwing oily glassware into a big vat full of hazardous (to you and the environment) chromic acid/sulfuric acid solution are gone. If you have a beaker containing oily mess, start by pouring the oil that will pour out into an appropriate safety container designated for slop oil. You might be surprised to learn that reﬁneries and petrochemical facilities do not want oil in their sewers. The next step would be to rinse the beaker with an appropriate solvent of relatively low toxicity such as isooctane, kerosene, denatured alcohol or varsol, as appropriate, to remove the oil. This should be done in a chemical fume hood with solvent proof gloves. The washings should go into the slop oil can. After the oil is removed, the glassware can be dried over a stream of air or nitrogen. The next step is a thorough scrubbing with non-ionic laboratory detergent (containing no metals). This should remove the residual oil ﬁlm. Rinse with water. The glassware can now be placed in an acid leaching bath or simply heated on a hot plate ﬁlled two-thirds full with 1:1 HCl or HNO3. Heat to boiling and allow to simmer 30 min. The acid may be saved and reused. Finish the process with a triple rinse in 18 Mohm cm quality deionized water. Glassware can be quickly dried by placing it in a laboratory oven or furnace, being careful that only clean surfaces are contacted. Carefully inspect clean glassware for chips and ﬂaws. Discard chipped or broken glassware. Quartz beakers used in high temperature ashing should be discarded when their bottoms appear etched and no longer transparent. 23.3.1.2 Fluoropolymers Fluorocarbon polymers like Teﬂon PFA are by nature semi-permeable membranes. They permit certain gases, small quantities of water and acids to penetrate the structure of the polymer, as well as some volatile element species such as elemental Hg, osmium tetroxide, SeH2 and probably other similar compounds [28]. Acid gases will gradually penetrate the Teﬂon liners of the steel jacketed digestion bombs and corrode the steel. Corrosion products then begin to penetrate the outside surface of the liner. When this happens the steel jackets should be cleaned and the liners replaced. Corroded steel is a signiﬁcant contamination risk for trace metal analysis.

713

R.I. Botto

Teﬂon PFA digestion vessels should be leached for several hours in hot 1:1 HNO3 for cleaning. It may be possible to clean them by running HNO3 “blanks” in the microwave oven but this may also tend to drive some contaminants deeper into the Teﬂon surface. Be careful not to use abrasives when cleaning Teﬂon vessels. Scratching could increase surface contamination and affect the seal. Some manufacturers recommend annealing new Teﬂon PFA vessels in a conventional oven before use. New vessels also will require acid leaching to remove contamination from the manufacturing process. Carefully inspect Teﬂon vessels and liners for damage or distortion. Discard any damaged vessels. 23.3.1.3 Ovens, furnaces and hoods Ovens and furnaces should be blown free of dust with a stream of compressed air on a regular basis to avoid contaminating samples. Spills in ovens should be cleaned up promptly. Quartz watch glasses may be placed over beakers in mufﬂe furnaces to help protect against dust or ﬁne ash from other samples or the refractory walls. To maintain a ﬂow of oxygen into the beaker use quartz rod spacers to provide a small gap. Chemical fume hoods are rarely designed for acid digestions. The typical hood has metal ﬁxtures and materials of construction as well as the screws that hold it together. Metal ductwork may also have been installed. All of this tends to corrode with use and generate scale particles that can contaminate samples. Our laboratory installed specially designed hoods constructed of serpentine rock and including a wash down feature. All screws were nylon and the ﬁxtures plastic. Ductwork above was constructed of ﬁberglass composition with nonmetallic ﬁttings. Even the cabinets below were constructed of wood. Although these hoods were expensive to construct they have served us well for 17 years with only minor repair to the stonework. Painting the interior with acid resistant epoxy is another technique used to extend useful lifetime of acid digestion hoods. 23.3.2 Clean techniques and disposable equipment “It’s a jungle out there!” The laboratory is full of lurking sources of contamination for trace element analysis. Flaking painted walls, ceiling tiles, corroding steel or painted bench tops, dusty cabinets, hoods, drawers, ﬂoors and equipment all pose potential contamination “hazards”. Short of constructing a clean room facility, keeping the lab as clean and dust free as possible is a step in the right direction. Paper towels are a source of lint but even worse, if you are not careful as to their source, may be a source of metals contamination. Paper whiteners Ti and Zn are present in the wrong types of paper towels. Check your supply by ashing one and analyzing the ash. Corroded metal cabinets should not be used to store clean glassware for analysis. Use clean, dust-free desiccators to store beakers for ashing or vessels for digestion. Polish

714

Sample preparation for crude oil, petroleum products and polymers

stainless steel bench tops and coat them with furniture spray oil or similar protective coating. Never use metal soaps or phosphate detergents for cleaning lab equipment. Wherever it is possible to use disposable equipment for sample preparation you have the advantage of avoiding contamination of a sample from the one run just before it. The procedure in Section “Ash solubilization by atmospheric pressure microwave digestion” that uses all disposable equipment is an example of a procedure that completely avoids the possibility of sample cross contamination and leaves no equipment to clean up afterward. Disposable equipment made from polyethylene, polysulfone, polycarbonate or similar “clean” polymers may not require pre-cleaning before use. Test this by running blanks in the equipment to check for contamination. Avoid using plastic equipment having ﬁlled (often white or colored) polymers for digestion or sample storage. If cleaning of plastic equipment is required, an acid leach followed by rinsing with 18 Mohm cm deionized water should sufﬁce. More information on techniques for avoiding analyte contamination has been provided by Schramel and Knapp in Chapter 2. 23.3.3 Quality assurance How many laboratories ignore the concept of quality assurance for the sample preparation procedure because by itself it does not produce numerical results for a QC chart? Are we good about running QC checks on our analytical instruments, verifying calibrations and correcting for interferences while we ignore the possibilities that our sample preparations are plagued with losses and contamination? To verify the accuracy of the entire analysis, recovery studies must be performed through the entire sample preparation procedure. This usually involves the use of standards containing known concentrations of the elements in the sample matrix or standard additions to the sample matrix. These recoveries should be repeated or a reference material analyzed at regular intervals to make sure nothing has changed. Standards for elemental analysis for the petroleum industry are now available from many commercial sources. Materials available from the National Institute of Standards and Technology include mixtures of organometallic elements in lube oil, trace elements in reference fuel oils, lead in hydrocarbon, and oils containing certiﬁed concentrations of S, N and Cl. A searchable on-line catalog is available. Can you obtain Certiﬁed Elements in Butyl Rubber? Not now. For many petrochemical materials there are no reference materials available to represent them. Naphtha, gasoline, most polymers and tars would all fall into that category. For these materials and many others you will have to certify your own “standard”. Our laboratory developed a standard for butyl rubber by obtaining a substantial quantity of a carefully chosen plant lot of

715

R.I. Botto TABLE 23.6 Analysis of butyl rubber by ICP-AES, 40 replicate determinations Element

Mean (ug/g)a

Sigma (ug/g)a

% RSDa

Preparation technique

Al B Ca Cr Cu Fe Na Ti V Zn

148 3.45 14.5 1.15 0.13 7.73 32.6 1.37 0.027 13.9

8.9 0.49 1.1 0.18 0.04 0.90 3.51 2.68 0.06 1.31

6.0 14 7.3 15 31 12 11 200 200 9.4

Dry/wet ashingb Dry ashing Dry/wet ashingb Dry ashing Dry/wet ashingb Dry/wet ashingb Dry/wet ashingb Dry ashing Dry/wet ashingb Dry/wet ashingb

a

Data collected over 8 year period—all points included. Results from a dry and wet ashing preparation were averaged to produce each result.

b

Fig. 23.2. Calcium in butyl rubber QC chart.

716

Sample preparation for crude oil, petroleum products and polymers

Fig. 23.3. Ti in butyl rubber QC chart.

material and analyzing it repeatedly. The method was veriﬁed by spike recovery studies as described earlier and shown in Table 23.4. Table 23.6 shows the precision obtained for 40 analyses of the butyl rubber “standard” performed over a period of 8 years. The quality control (QC) chart for Ca is typical (Figure 23.2). One “out of three sigma” value for Ca probably represented an incidental contamination. As would be expected, low concentrations are associated with larger %RSD values, as they are being determined closed to the detection limit. However, the Ti data (Figure 23.3) reveal a period of months in which high and extremely variable results were being produced. During this period we searched for the cause. A source of contamination, probably a particular brand of paper products being used in the laboratory was assigned the blame. A more recent event for V (Figure 23.4) was traced to coke samples containing high V content that were being ashed in the same mufﬂe furnaces at roughly the same time. If possible, self-certiﬁed standards should have adequate concentrations of the elements of interest for valid precision data. A standard having

717

R.I. Botto

Fig. 23.4. V in butyl rubber QC chart.

non-detectable amounts of the element would only indicate when contamination events had occurred. To obtain adequate concentrations it may be necessary to spike a batch of material with the analyte elements. This can be a difﬁcult procedure for polymer samples and homogeneity may be difﬁcult to achieve. For liquid samples, spiking is straightforward but stability is the issue and liquid “standards” may have to be spiked just prior to each use. Further discussions on quality assurance for elemental analysis in the petroleum industry may be found in Refs. [2,86].

ACKNOWLEDGEMENTS My special thanks go to I.B. (Joe) Brenner, R.A. (Kishore) Nadkarni and Frank C. McElroy for many interesting and productive discussions on sample preparation.

718

Sample preparation for crude oil, petroleum products and polymers

REFERENCES 1 2 3 4

5

6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

21 22

23 24 25 26 27 28

J.H. Gary and G.E. Handwerk, Petroleum Reﬁning: Technology and Economics, 3rd edn. Marcel Dekker, Inc., New York, 1994. R.I. Botto, Spectrochim. Acta Rev., 14 (1991) 141. W.F. Baade, G.D. Snyde and J.M. Abrardo, Hydrocarbon Process., 72 (1993) 117. V. Sychra, I. Lang and G. Sebor, Analysis of petroleum and petroleum products by atomic absorption spectroscopy and related techniques, Prog. Anal. At. Spectrosc., 4 (1981) 341. R.A. Nadkarni, A review of modern instrumental methods of elemental analysis of petroleum related material: Part I—occurrence and signiﬁcance of trace metals in petroleum and lubricants. In: R.A. Nadkarni (Ed.) Modern Instrumental Methods of Elemental Analysis of Petroleum Products and Lubricants. ASTM publication STP 1109, 1991 ASTM method D 4057-95, 1997. ASTM method D 4177-95, 1995. R.A. Nadkarni, Guide to ASTM Test Methods for the Analysis of Petroleum Products and Lubricants, ASTM stock number MNL44, 2000. ASTM method D 482-95, 1996. ASTM method D 128-98, 1998. ASTM method D 4422-94 (Re-approved 1998). R.A. Nadkarni, Anal. Chem., 52 (1980) 929. ASTM method D 874-96, 1996. R.A. Nadkarni and R.R. Ledesma, SAE publication 952548, 1995. ASTM method D 5184-91 (Re-approved 1995). Z. Sulcek and P. Povondra, Methods of Decomposition in Inorganic Analysis. CRC Press, Boca Raton, FL, 1989. R.A. Nadkarni, personal communication, 2003. ASTM method D 3228-96, 1996. ASTM method D 1091-91 (Re-approved 1995). R.A. Hofstader, O.I. Milner and J.H. Runnels (Eds.), Analysis of Petroleum for Trace Metals, American Chemical Society Advances in Chemistry Series 156. Washington, DC, 1976. H.H. Walker, J.H. Runnels and R. Merryﬁeld, Anal. Chem., 48 (1976) 2056. K. Everett and F.A. Graf Jr., Handling perchloric acid and perchlorates. In: N.V. Steere (Ed.), Handbook of Laboratory Safety, Second Edition. CRC Press, West Palm Beach, FL, 1971. J.A. Nobrega, L.C. Trevizan, G.C.L. Araujo and A.A. Nogueira, Spectrochim. Acta, 57B (2002) 1855. L.M. Costa, F.V. Silva, S.T. Gouveia, A.R.A. Nogueira and J.A. Nobrega, Spectrochim. Acta, 56B (2001) 1981. J.H. Liu, R.E. Sturgeon and S.N. Willie, Analyst, 120 (1995) 1905. Operating Instructions for Parr Digestion Bombs, Parr Instrument Company publication No. 249M. T. Wondimu and W. Goessler, Bull. Chem. Soc. Ethiopia, 14 (2000) 99. H.M. Kingston and L.B. Jassie (Eds.), Introduction to Microwave Sample Preparation: Theory and Practice. American Chemical Society, Washington, DC, 1988.

719

R.I. Botto 29

30 31 32 33 34 35 36 37

38 39 40 41 42 43 44

45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65

720

Microwave assisted acid digestion of siliceous and organically based matrices including ash, biological tissue, oil, oil contaminated soil, sediment, sludge, and soil, USEPA Method 3052, 1995. M.B. Martin-Garcia, D. Bellido-Milla, A. Jimenez-Jimenez and M.P. HernandezArtiga, Fresenius J. Anal. Chem., 364 (1999) 527. C. Sanz-Segundo, M.P. Hernandez-Artiga, J.L. Hidalgo-Hidalgo de Cisneros, D. Bellido-Milla and I. Naranjo-Rodriguez, Mikrochim. Acta, 132 (1999) 89. R.I. Botto, J. Anal. At. Spectrom., 8 (1993) 51. Milestone S.r.I Digestion Application Reports #06-002, #06-003, #06-004, #06-005 and #10-008. Operating Instructions for Parr Microwave Acid Digestion Bombs, Parr Instrument Company publication No. 243M. T. Wondimu, W. Goessler and K.J. Irgolic, Fresenius J. Anal. Chem., 367 (2000) 35. M. Horton, D. Leong and J.D. Hwang, ICP Inform. Newsl., 28 (2002) 331. H.M. Kingston, Short Course on Microwave Sample Preparation for Inorganic Analysis ST-07 (Notes), 2002 Winter Conference on Plasma Spectrochemistry, Scottsdale, Arizona, January 2002. A.G. Howard, L. Labonne and E. Rousay, Analyst, 126 (2001) 141. Parr Oxygen Combustion Bomb, Parr Instrument Company Bulletin 4700. H. Narasaki, Anal. Chem., 57 (1985) 2481. Y. Nakamoto and T. Tomiyama, Bunseki Kagaku, 46 (1997) 665. M. Murillo, N. Carrion and J. Chirinos, J. Anal. At. Spectrom., 8 (1993) 493. ASTM method D 129-95, 1995. R.A. Nadkarni, Comprehensive elemental analysis of coal and ﬂy ash. In: E.L. Fuller (Ed.), Am. Chem. Soc. Symp. Ser. 205. American Chemical Society, Washington, DC, 1982, p. 147. R.A. Nadkarni, Am. Lab., 13 (1981) 22. R.A. Nadkarni, Int. Lab., 26 (1981). ASTM method D 808-95, 1995. W. Schoeniger, Mikrochim. Acta, (1955) 123 and (1956) 869. ASTM method D 1266-98, 1998. ASTM method D 2784-98, 1998. ASTM method D 86-87 (Re-approved 1999). ASTM method D 4055-97, 1997. ASTM method D 2007-98, 1998. L. Liang, M. Horvat and P. Danilchik, Sci. Total Environ., 187 (1996) 57. V. Fernandez-Perez, M.M. Jimenez-Carmona and M.D. Luque de Castro, Anal. Chim. Acta, 433 (2001) 47. ASTM method D 721-97, 1997. B.W. Renoe, Am. Lab.,(August) (1994) 34. M.J. Jager, D.P. McClintic and D.C. Tilotta, Appl. Spectrosc., 54 (2000) 1617. D.M. Jones, J.S. Watson, W. Meredith, M. Chen and B. Bennett, Anal. Chem., 73 (2001) 703. R.I. Botto, Spectrochim. Acta, 42B (1987) 181. I.B. Brenner and A.T. Zander, ICP Inform. Newsl., 20 (1995) 738. J.G. Bansal and F.C. McElroy, SAE Technical Paper Series #932694. SAE International, Warrendale, PA, 1993. S.D. Olsen, S. Westerlund and R.G. Visser, Analyst, 122 (1997) 1229. F. McElroy, A. Mennito, D. Ebenezer and R. Thomas, Spectroscopy, 13 (1998) 42. R.I. Botto, Can. J. Anal. Sci. Spectrosc., 47 (2002) 1.

Sample preparation for crude oil, petroleum products and polymers 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86

D. Hausler, Spectrochim. Acta, 42B (1987) 63. ASTM method D 4951-96, 1996. ASTM method D 5185-97, 1997. ASTM method D 5708-95a, Test method A, 1995 Polymer Solubilities, Solvents and Dissolution Temperatures, CEM Corporation Application Note. C.J. Lord III, Anal. Chem., 63 (1991) 1594. M. Murillo and J. Chirinos, J. Anal. At. Spectrom., 11 (1996) 253. H.M. Al-Swaidan, Talanta, 43 (1996) 1313. I.B. Brenner, A. Zander, S. Kim and J. Shkolnik, J. Anal. At. Spectrom., 11 (1996) 91. S.J. Kumar and S. Gangadharan, J. Anal. At. Spectrom., 14 (1999) 967. K.J. Eisentraut, R.W. Newman, C.S. Saba, R.E. Kauffman and W.E. Rhine, Anal. Chem., 56 (1984) 1086A. C.S. Saba and K.J. Eisentraut, Anal. Chem., 51 (1979) 1927. J.R. Brown, C.S. Saba, W.E. Rhine and K.J. Eisentraut, Anal. Chem., 52 (1980) 2365. R.E. Kauffman, C.S. Saba, W.E. Rhine and K.J. Eisentraut, Anal. Chem., 54 (1982) 975. M.P. Hernandez-Artiga, J.A. Munoz-Leyva and R. Cozar-Sievert, Analyst, 117 (1992) 963. Nakamura, et al., Bunseki Kagaku, 34 (1985) T85. D. Liederman, J.E. Bowen and O.I. Milner, Anal. Chem., 31 (1959) 2052. U. Telgheder and V.A. Khvostikov, J. Anal. At. Spectrom., 12 (1997) 1. R.I. Botto and J.J. Zhu, J. Anal. At. Spectrom., 9 (1994) 905. R.I. Botto and J.J. Zhu, J. Anal. At. Spectrom., 11 (1996) 675. R.I. Botto, Spectrochim. Acta, 39B (1984) 95.

721

This page intentionally left blank

Chapter 24

Sample preparation of geological samples, soils and sediments Philip J. Potts and Philip Robinson

24.1

INTRODUCTION

Sample dissolution for the determination of trace elements in geological samples including soils and sediments is an enormously varied, demanding, but at the same time fascinating topic of study. These attributes are due to the nature of this type of sample. For example, silicate rocks comprise a wide range of different mineral species, which gives rise to fundamental issues of how to select and prepare a representative sample prior to dissolution. Then there are questions related to the enormous range of different elemental and isotopic measurements required by the various branches of geological sciences. Indeed, in the most demanding research applications, analytical parameters such as the precision, accuracy and the detection limit capability must often be state-ofthe-art. The realization of these expectations is directly related to the quality of the sample preparation procedure. And, ﬁnally, all the above attributes are reﬂected in the large variety of sample preparation procedures that have been developed to accommodate the wide range of elemental and mineralogical compositions that fall within this ﬁeld taking account of the users’ requirements for data quality. This chapter reviews the sample preparation and dissolution procedures that currently support geological and environmental ﬁelds of study in the analysis of rocks, soils and sediments. 24.2

SAMPLE PREPARATION

Sample preparation is often considered to cover procedures used to prepare a selected test portion prior to an analytical measurement. However, in the analysis of geological and environmental samples (rocks, soils and sediments), sample preparation has much wider implications that must be taken into account before a laboratory measurement may be made and interpreted with conﬁdence. There are two principal considerations. First, the selection and collection of a representative sample from the ﬁeld. Second, the procedures Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

723

P.J. Potts and P. Robinson

used for crushing and grinding the collected sample to ensure that a statistically representative test portion can be taken for analysis. 24.2.1 Sample collection A very wide range of samples are collected by scientists undertaking geochemical and environmental research. However, to illustrate a number of issues, this discussion will be restricted to two applications: (i) research involving the ﬁeld collection of silicate rock samples and (ii) sample collection associated with the characterization of contaminated land. (i) In undertaking geochemical research that involves the collection of rock samples from the ﬁeld, the usual aim is to select and bring back to the laboratory samples that have a chemical composition that is representative of a rock outcrop, lava ﬂow or even an entire unit of rock marked on a geological map. There are often signiﬁcant logistical constraints that must be applied to this process in terms of ease of access, if the ﬁeld area is remote, and serious limitations in the amount of material that may be transported out of the ﬁeld area. These limitations often mean that the mass of material returned to the laboratory is restricted to a single sample, for each sampling locality, of 1–2 kg [1]. In order to ensure that this sample is as representative as possible, great care is usually taken to hammer the material directly from the outcrop (i.e., to avoid collecting loose material that is not in situ) and that every care is taken to ensure that the sample is free of weathered surfaces, veining or secondary alteration effects. Whereas this sampling process is the foundation on which the majority of “hard-rock” geochemistry research is based, there are a number of limitations to this approach that could affect the quality of analytical results. The most important is that collection of a single sample provides no information about either the local or extended homogeneity of the sampling target. Uncertainties associated with such effects cannot then be accounted for in the interpretation of analytical results. If the sampling target is a large or extended geological unit (in the extreme a whole mountain, the majority of which is, of course, inaccessible beneath the surface), a sampling design based on some sort of grid covering the entire surface of the sampling target would provide information on its homogeneity, and hence provide additional information of value in the conﬁdent interpretation of analytical results. Accepting that this approach may not always be practical because of logistical issues or considerations of time and expense, another approach is to consider the collection of duplicate samples from a locality. Differences between duplicate samples that have each been prepared and analyzed independently provides some additional information on sampling effects for a more conﬁdent interpretation of results. In fact, duplicate samples collected from a grid of locations covering a target may provide a means of evaluating sampling uncertainty, as summarized in (ii) below.

724

Sample preparation of geological samples, soils and sediments

A further consideration in the collection of ﬁeld samples is the mass of material required so that a representative test portion can be selected for analysis in the laboratory. Representative mass depends on a number of factors, including grain size of the rock and the way the analyte is distributed in host materials. In one investigation of the coarse grained Murchison granite, it was found that a minimum of 3 kg of material was required to form a representative sample (Paul Davidson, Personal Communication). These issues in relation to analytical test portions are discussed further in Section 24.2.3. (ii) The collection of samples in environmental applications, for example, in the assessment of contaminated soil, has reached a much higher level of sophistication. This is probably related to the problem that if the objective is to make a judgment as to whether or not the level of a contaminant exceeds a trigger or action level, decisions are much less related to the uncertainty of measurements in the laboratory, but rather uncertainty associated with the distribution of the contaminant at the site itself. At many contaminated sites, hazardous elements are likely to be distributed in soil in an inhomogeneous manner. Very different analytical results may then be obtained from soil samples collected sometimes even a few centimeters apart from a particular locality. Interpretation of results then critically depends on an assessment of sampling uncertainty (i.e., uncertainty associated with the composition of nominally identical samples all collected from the same locality). One effective way of characterizing this problem in the assessment of soil at contaminated sites has been developed over a number of years by Ramsey and co-workers [2, 3]. The principle is that having set out a grid of sampling localities marked out at a ﬁeld site under investigation, duplicate samples are collected at typically 10% of these localities (selected at random). One convention is that these duplicate samples should not be collected at exactly the same position, but separated by a distance representative of the accuracy with which that locality could be relocated if a follow-up investigation were justiﬁed. An analysis of variance of results from all the samples collected at the site will provide the following information: – differences between the composition of duplicate test portions from individual samples will provide a measure of analytical uncertainty (associated with the laboratory/analytical procedure); – differences between the average composition of the duplicate samples collected from each locality will provide a measure of sampling uncertainty (associated with how representative the composition of a sample is of a particular locality); Variations in the average composition of samples from each locality at the site will provide a measure of the heterogeneity of the site, or the way in which a contaminant is distributed across the site, taking into account both analytical and sampling uncertainties. The criterion proposed by Ramsey and co-workers [2,3] is that if real differences in the composition of samples taken from

725

P.J. Potts and P. Robinson

localities are to be identiﬁed and interpreted with conﬁdence, the variance associated with analytical and sampling uncertainty should, together, not exceed 20% of the total variance of the site data set. The rigor of this approach sets a standard to which all geochemical sample collection studies should aspire, taking into account the time, expense and other logistical restrictions. 24.2.2 Crushing and grinding Having collected a representative sample from the ﬁeld, there are further considerations in the crushing and grinding of a sample to provide a laboratory sample from which representative test portions can be taken for analysis. These considerations can be illustrated by considering the procedures relevant to the analysis of silicate rocks. The fundamental issue is that most silicate rocks are crystalline in nature and are intrinsically inhomogeneous if the mass of test portion is reduced beyond a certain limit. To some extent, this limitation can be overcome by applying effective milling procedures to reduce the mean particle size in the material presented to the laboratory for analysis. However, although standard milling equipment is capable of achieving a particle size distribution that will pass a 200 mesh sieve (individual grain size ,74mm), further grinding to achieve an even ﬁner grain size distribution becomes progressively less effective. Furthermore, all grinding equipment will cause some degree of contamination, so extended contact with such equipment may cause a signiﬁcant unwanted enhancement of certain trace elements that may be of interest. For this reason, agate-lined milling equipment is the choice for the most demanding trace element applications, to avoid the varying degrees of contamination that will result from the use of tungsten carbide (W, Co and Ta) or steel (Fe, Mn and Cr) equipment. A typical procedure for a rock is, therefore, as follows: (i)

remove, by hammering or sawing, any surface contamination, weathering or veining; (ii) break up large pieces of material with a hammer or rock splitter; (iii) further break up these pieces to provide material of a coarse sand-like grain size (2– 5 mm). Note that some laboratories use a jaw crusher for this process. However, unless the blades of the jaw crusher are rigorously cleaned before use, cross-contamination of samples can occur. After cleaning, a preliminary portion of sample can be crushed and discarded to minimize this cross-contamination effect; a better alternative is to crush rock samples down to ,5 mm fragments with a hydraulic press ﬁtted with tungsten carbide plates. The crushed rock, including the ﬁne dust, can then be coned and quartered down to about 80–100 g for the next milling stage. (iv) if several batches must be combined and homogenized, this is best undertaken at this stage;

726

Sample preparation of geological samples, soils and sediments

(v)

mill the resultant material in an agate-lined swing mill or orbital mill for a time appropriate to crush the sample to a grain size that will pass a 200– 250 mesh sieve. Although it may be appropriate to sieve a test portion to evaluate the effectiveness of the milling process, dry sieving the entire sample to 200/250 mesh is impractical because of rapid clogging of the sieve mesh and the risk of introducing further contamination from the sieve and indeed, segregation of different mineral types. Rock powder may be washed through a sieve with water, but after drying, the sieved powder will cake and need re-milling and there is always the risk that water may leach out any labile analytes. Furthermore, sieving may cause the segregation of some mineral species from the bulk matrix; (vi) the resultant powder may then be stored in a bottle or packet for analysis. 24.2.3 Selecting an appropriate test portion In many non-geological applications, instructions to analysts require that the sample presented to the laboratory for analysis is homogenized by shaking or blending, and then a suitable test portion is removed for analysis. In the case of silicate rocks, there are additional considerations arising from the fact that most samples do not contain a uniform distribution of particles, particularly with respect to shape and composition, and that conventional blending processes may be ineffective. If the sample contains ﬂaky minerals (e.g., micas) or grains having a high density contrast (e.g., native gold grains in a silicate matrix), simple shaking will not guarantee homogenizing a powder, and use of a sample splitter may be required to give conﬁdence that a statistically representative test portion has been selected for analysis. In addition, the mass of the test portion required to obtain a representative analytical sample is not a trivial consideration for some trace element analytes. The “nugget effect” is a well-known phenomenon in the determination of gold and the platinum-group elements (PGEs). This phenomenon arises from the fact that gold and the PGE are generally present as discrete grains that cannot be ground by milling processes. The mass of test portion must then be sufﬁciently large to contain a representatively large number of individual grains to minimize differences in composition between test portions of identical mass caused by statistical variations in the number of gold grains they contain. In addressing this phenomenon, Clifton et al. [4] developed a nonogram that indicates the mass of test portion in relation to the bulk concentration of gold and the average size of individual grains of native gold. This nonogram is based on the assumption that each test portion should contain on average 25 individual grains. Results show that if samples contained low concentrations of gold in relatively large grain sizes, the mass required could be in the tens or even hundreds of kilograms range. Although the nugget effect is not normally as serious a problem in the determination of the more common trace elements, it still requires careful

727

P.J. Potts and P. Robinson

consideration for elements that are normally present at high concentrations in minor or rare phases. Examples include Zr, Hf (e.g., zircon), Cr (e.g., chromite), Sn (cassiterite), REE, U, Th (various minor phases). This problem is exacerbated by the widespread adoption of ICP-MS techniques, in which it is possible to take advantage of the high sensitivity (coupled to an intolerance of solutions of high dissolved salt content) by selecting 0.1 g or even smaller test portions. As the analyst may have no prior knowledge of the mineralogy of samples presented for analysis, the only effective approach is to analyze duplicate test portions of a proportion of samples in a batch and ensure that the agreement between results matches the repeatability speciﬁcation of the technique. This action should in any case be part of a laboratory’s routine quality assurance programme.

24.3

CHOICE OF APPROACH

A number of criteria must be assessed before deciding the most appropriate method for the preparation of geological and environmental samples for trace element determination. These criteria include the properties of individual, or groups of elements and the choice of a suitable technique. However, whatever the detail, the overall assessment should be based on ﬁtness-for-purpose criteria. 24.3.1 Fitness-for-purpose In the context of trace element determinations, ﬁtness-for-purpose means that there should be a match between the quality of analytical results (in terms of precision, accuracy, detection limits, cost and other factors) and the use/interpretation to which the results will be subjected. Simple examples of this principle are that if a technique does not have a detection limit ten times less than the lowest elemental concentration that must be measured, it may not be ﬁt-for-purpose, and an alternative technique should be chosen. Conversely, if the precision of analytical measurements is an order of magnitude better than that required to satisfy an application, it may well be that the laboratory is wasting analytical resource. That said, there are only a limited number of techniques that are widely used in the analysis of geological and environmental samples, their popularity depending on high sensitivity in the determination of trace elements and efﬁciencies gained by their multi-element capabilities. 24.3.2 Choice of sample preparation procedure based on choice of technique The most popular techniques used for routine trace element determination of geological and environmental samples are X-ray ﬂuorescence and ICP-MS

728

Sample preparation of geological samples, soils and sediments

(see for example, Potts [5]). In addition, instrumental neutron activation analysis (INAA), ICP-atomic emission spectrometry and atomic absorption spectrometry remain popular. Choice of technique depends in part on the characteristics of the instrumentation and may be summarized as follows: XRF: Trace elements are normally determined in the laboratory on samples prepared as compressed powder pellets. The alternative form of sample preparation, glass disks, is routinely used for the determination of the major elements, although schemes of trace element analysis have also been developed [169], particularly for applications where insufﬁcient sample is available for a conventional powder pellet (see Section 24.5). INAA: This technique is effective for the determination of Au, Sb, As, selected REE, Ta, Hf, Th, Ba, Cs, Sc and several other elements, although the requirement to irradiate samples in a nuclear reactor has reduced the popularity of the technique. Conventional sample preparation comprises simply encapsulating 0.3–0.5 g of powder in a suitable polyethylene capsule. Note that for some exploration applications (e.g., Au), alternative sample presentation schemes have been developed involving the irradiation of 10–50 g samples, to allow measurements to be made on representative test portions. The radioactivity of the irradiated sample is subsequently counted by gamma ray spectrometers without further sample processing. In a more sophisticated version of this technique, the irradiated sample is subjected to sample dissolution and ion exchange separation to remove the matrix to allow the REE and other element fractions to be counted with enhanced sensitivity. This procedure is now almost obsolete, largely replaced by ICP techniques capable of providing equivalent or superior data. AAS, ICP-OES, ICP-MS: The conventional form of sample preparation for all these techniques is dissolution to form a liquid. ICP-MS is most sensitive in terms of detection limit capability, but also the most susceptible to interferences from the presence of high levels of dissolved salts and acid. AAS is least sensitive on both counts. The choice of sample preparation procedure—typically either acid dissolution, molten salt fusion or partial extraction—is then inﬂuenced by the nature of both sample matrix and analyte. In some cases, a further enhancement in analytical performance can be achieved by isolating the analyte from the matrix. In other cases, a sequence of partial extractions may give rise to speciation information of direct relevance to the interpretation in a particular application. The characteristics of all these sample preparation procedures will be considered in more detail in Sections 24.6–24.9. 24.3.3 Choice of sample preparation based on the characteristics of elements Although standard sample preparation procedures have been developed to permit the determination of a range of elements by the most popular

729

P.J. Potts and P. Robinson

instrumental forms of analysis, some groups of elements demand special consideration. (i)

Gold and the PGEs: Conventional sample dissolution techniques can be used in some applications. However, ﬁre assay has been developed speciﬁcally to permit the analysis of large test portions (20–50 g) and so minimize sampling uncertainty (caused by the nugget effect, referred to above). This specialized sample preparation technique offers effective chemical separation of the elements of interest from potentially interfering elements. Further details of these procedures are included in Section 24.7.4). (ii) Hg, As, Sb, Se, Sn, Bi and Te: Elements in this group have chemical properties that make them particularly suited to forming volatile species, either by cold vapor generation (Hg) or hydride generation (As, Sb, etc.). This property provides a selective and quantitative extraction of the elements of interest and was ﬁrst exploited in atomic absorption spectrometry and has been used in conjunction with ICP-OES. Further details of these procedures are included in Section 24.8.3. 24.4

METHODS THAT DO NOT REQUIRE ANY SAMPLE DIGESTION— IN SITU METHODS OF ANALYSIS

Although there is a presumption that sample dissolution is a prior requirement to any form of trace element analysis, this is not the case in some important geochemical applications. The sample preparation requirements for the XRF powder pellet technique and INAA have already been referred to in the previous section. However, there are three other categories of measurement that do not require sample dissolution for trace element determinations. To undertake a trace element determination for the majority of analytical techniques, a sample must be removed from the sampling target and taken to a laboratory for sample preparation and analysis. However, there are a small number of techniques that can be transported to the sampling target and placed in contact with the material to be analyzed without any form of sample preparation. As the options available to the analyst/investigator comprise sample selection rather than sample preparation, there are some important limitations in the interpretation of analytical results. Two of the more important techniques in this category are portable X-ray ﬂuorescence analysis and gamma ray spectrometry. 24.4.1 Portable X-ray ﬂuorescence Modern instruments use either a radioactive excitation source or a miniature X-ray tube, coupled to some form of energy dispersive detector (e.g., Si(PIN) diode or mercury(II) iodide crystal). In the in situ mode of operation,

730

Sample preparation of geological samples, soils and sediments

the portable analyzer is placed in contact with the sample surface and an X-ray ﬂuorescence spectrum acquired for 1–5 min. Spectrum analysis and quantiﬁcation is then undertaken in real time so that the analytical results are immediately available to the operator. To illustrate capabilities, an assessment of the trace element detection limit capabilities of one commercial P-XRF instrument that uses 109Cd, 55Fe and 241Am excitation sources, coupled to a HgI2 detector is given in Table 24.1 [6]. In the analysis of geological and environmental samples, there are several applications where this in situ ﬁeld method of analysis shows clear advantages. If the instrument is being used for the assessment of contaminated soil at a ﬁeld site (e.g., Argyraki et al. [7]), the immediate availability of results provides the operator with valuable information to base decisions on what to sample and analyze next. If the aim is to understand the distribution of a contaminant at the ﬁeld site or to locate hot spots of contamination, this interactive sampling and analysis approach has clear advantages. The range of elements that can be effectively assessed by P-XRF in contaminated soil at environmentally signiﬁcant levels depends on land use. In the latest UK soil guideline values [8], P-XRF has an adequate detection limit capability to determine lead (Pb), whatever the land use, and the elements As, Cd, Cr, Hg and Se for land used for commercial and industrial purposes, but not domestic use (see Table 24.1). A second important area of application of P-XRF for trace element determination that takes advantage of its non-destructive capabilities is in TABLE 24.1 P-XRF detection limits [6] (mg g21) compared with UK soil guideline values for various toxic elements of environmental interest Element

P-XRF detection limits

UK soil guideline values [8] Allotment Commercial Residential Residential and industrial with plant without plant uptake uptake

As 60p Cd 250p Cr(VI) 1080 Hg(total) 80p Ni 116 Pb 39 Se 45p

20 1,2,8a 130 8 50 450 35

20 30 200 8 75 450 260

20 1,2,8a 130 15 50 450 35

500 1400 5000 480 5000 750 8000

P-XRF detection limits: 200 s count time (p ¼ estimated from the response of adjacent elements in the XRF spectrum).

Cr: Assumes all Cr is Cr(VI). a

Cadmium values represent data for soil pH values of 6, 7 and 8, respectively.

731

P.J. Potts and P. Robinson

the analysis of a range of rock artifacts, including museum specimens, e.g., Neolithic stone axes and other lithic archaeological artifacts [9] and works of art Caesareo et al. [10]. In general, these types of material cannot be sampled (because they are too valuable and/or would be seriously disﬁgured by sampling), and cannot be removed to the laboratory for analysis (too large, ﬁxed in situ, or cannot be moved because of museum restrictions). In each case, portable XRF can provide in situ, non-destructive trace element analysis for applications that are often related to provenancing the artefact to a potential source to, conservation or to the detection of forgery. In each of these applications, care is needed in the interpretation of results when essentially a surface analysis by P-XRF is being used to estimate the bulk composition of the sample. The area on the surface of a sample from which the analytical signal is derived is typically 25 mm in diameter and the depth is usually between 0.1 and 1 mm, depending on the penetration depth of the ﬂuorescence X-rays of interest. When interpreting results, care is required to take into account surface contamination or alteration effects.

24.4.2 Gamma spectrometry Gamma spectrometry is also an in situ ﬁeld technique that is capable of determining U, Th and K from the gamma ray emissions of naturally-occurring isotopes derived from these elements found in rocks [11]. Instrumentation comprises a NaI(Tl) scintillation gamma ray detector that is used to acquire the gamma spectrum in the range 1–3 MeV. Gamma rays from selected isotopes are then measured and quantiﬁed. One consideration is the mass of sample from which the analytical signal originates, which for ﬁeld measurements is of the order of 80 kg, taking into account the range of the relevant gamma rays in rock. Gamma spectrometry has generally been used for ﬁeld mapping the distributions of the trace elements U and Th (and K).

24.4.3 Laser ablation techniques and other microprobe/ microanalytical techniques Laser ablation techniques can be regarded as microprobe in situ methods of analysis and are widely used for the determination of trace elements in geological samples. Samples are normally prepared as small polished slabs which are mounted in a suitable cell and excited with a laser, typically 50 mm in diameter. The laser beam is designed to have a sufﬁciently large power density to ablate material from the surface of the sample as particulates that should ideally have as small a particle size distribution as possible. This material is swept by argon carrier gas into an inductively couple plasma where evaporation, atomization and ionization of this material occurs, prior to

732

Sample preparation of geological samples, soils and sediments

analysis by ICP-MS. The frequency of the laser has a signiﬁcant inﬂuence on ablation characteristics (including the degree of fractionation associated with the ablation process observed in measurements). Almost all current applications use UV lasers with frequencies of 266, 213, 193 and 157 nm that represent the spectrum extending from the routine through to the current frontier of research developments. Laser ablation ICP-MS is a rapidly developing ﬁeld of considerable contemporary research interest and is discussed in detail in Chapter 20. The same technique can be used to determine the bulk (as opposed to microprobe) trace element composition of silicate materials, and there has been recent interest in combining the capabilities of XRF (for the determination of the major elements and LA-ICPMS (for the determination of the trace elements) on samples prepared as glass disks by fusion with lithium borate ﬂuxes (see, for example, Yu et al. [12]). Other microprobe techniques are available for the determination of trace elements and isotope ratios, normally on geological samples prepared as either polished thin sections or small slabs of material. These techniques include the ion probe, PIXE, synchrotron radiation XRF and to some extent the electron microprobe (although the latter technique is generally used for the determination of the major elements in minerals at the micrometer resolution scale). Further details of these specialized techniques can be found in Ref. [13]. 24.5

METHODS BASED ON SOLID SAMPLES

One of the unusual aspects of the determination of trace elements in rocks, soils and sediments is that, for some techniques, sample dissolution is not required. In order to give a complete overview of techniques and procedures in this ﬁeld, the preparation of samples that can be analyzed in the laboratory directly for trace elements is reviewed here. 24.5.1 Direct determinations on powders The preferred form of sample preparation for a technique such as INAA is to seal typically about 0.3 g of powder into a polyethylene capsule and to irradiate it in a nuclear reactor for a suitable time, together with other unknown and calibration samples and neutron ﬂux monitors. After irradiation, the gamma ray spectrum from encapsulated sample can be measured directly with no further pre-treatment. As with other techniques, the principal issue is to ensure that the test portion selected for irradiation is representative of the bulk sample. Loose powders can also be analyzed for trace elements by XRF. The sample powder is simply poured into an XRF sample cup, which normally has a bottom comprising a thin X-ray transparent polymer ﬁlm through which measurements on the sample are made.

733

P.J. Potts and P. Robinson

24.5.2 Powder pellet for XRF A more formal sample presentation scheme is widely used for the determination of trace elements by XRF in which sample powders are conﬁgured as compressed pellets. A typical scheme has been described by Watson [14]. (i) (ii) (iii) (iv) (v) (vi)

Weigh out approximately 10 g of rock powder and mix with 0.7 –0.9 ml of polyvinylpyrrolidone –methyl cellulose (PVP –MC) binder in a plastic bag. Rub the bag and contents to ensure intimate mixing of the sample and binder to form a moist paste. Place the paste into a mould and ﬁt the piston. Place the mould and piston in position in a hydraulic press and compress at 2– 6 ton in.2 depending on the nature of the sample. After releasing the hydraulic pressure, remove the powder pellet and dry it in an oven for at least 2–3 h at 105–1108C. After removing from the oven and cooling, the pellet can be analyzed by XRF.

24.5.3 Glass disks for XRF trace determinations The standard method for the determination of the major elements by XRF involves the fusion of the sample powder with a lithium metaborate/tetraborate ﬂux. After dissolution of the sample in the ﬂux, the melt is poured into either a mould and quenched with a plunger, or a casting dish and quenched by cooling to form a ﬂat transparent glass disk that is suitable for XRF analysis. For the determination of major elements followed by LA-ICP-MS for the determination of the trace elements, a sample to ﬂux ratio of between 1– 3 and 1–5 would normally be used. However, “low dilution” methods for the more sensitive determination of trace elements by XRF on glass disks use a sample to ﬂux ratio of about 1–2. These latter procedures are particularly useful for trace element determination by XRF of samples where insufﬁcient material is available for the preparation of conventional powder pellets. A typical scheme of analysis for the preparation of XRF glass disks is as follows: (i) Dry the rock powder in an oven at 1108C overnight. (ii) Weigh the appropriate amount of dried ﬂux into a platinum –5% Au crucible and then weigh in the appropriate amount of dried rock powder (Pt –5% Au crucibles have non-wetting properties). (iii) After mixing the powders with a polythene rod (care to avoid crosscontamination), place the crucible in a mufﬂe furnace that has been preheated to 11008C for 15– 20 min. Swirl the mixture every 5 min to ensure thorough mixing. (iv) When a clear molten glass has been obtained, pour the contents of the crucible into a shaped brass mould (preheated to about 2008C to reduce

734

Sample preparation of geological samples, soils and sediments

thermal shock) and ﬁrmly lower a plunger to quench the glass into a ﬂat transparent disk. As an alternative, the molten glass can be poured to ﬁll a Pt/Au casting dish (preheated brieﬂy in the furnace) and quenched by cooling with a stream of air to take up the shape of the casting dish. As an alternative to manual manipulations, automatic equipment is available to undertake this entire process, based on either RF induction or gas burner heating. 24.6

DISSOLUTION METHODS BASED ON ACID ATTACK

Strong mineral acids such as HF, HClO4, HNO3, HCl and H2SO4 have long been favored for the decomposition of rocks and minerals. Either in open vessels or in closed ones under pressure, most samples will dissolve, given sufﬁcient time and an appropriate temperature. Aspects of these procedures have been detailed in Chapters 6 and 8. There are good reasons for using acids in the trace element analysis of geological materials: 1. Modern instrumental techniques such as ICP-MS require dilute solutions. The alternative of using alkali fusion adds considerably to the salt content of the solution. 2. Use of HF results in removal of the major element Si as volatile SiF4. This reduces the risk of precipitates, as Si is difﬁcult to keep in solution, and also results in a more dilute solution. 3. Very low reagent blanks can be achieved with acids, which are essential for samples such as ultramaﬁc and carbonate rocks that can have extremely low trace element contents. Although it is possible to obtain some pure alkali ﬂuxes nowadays, acids can be readily vapor distilled in silica or PTFE stills. Many trace element laboratories produce their own double distilled acids or alternatively they can be purchased commercially. 4. Attack on the digestion vessel is less with acids where PTFE is the standard material used. Fusions require expensive Pt or less satisfactory carbon, Zr, Ni or Fe crucibles. Fusion ﬂuxes, such as Na2O2 and NaOH, are excellent for decomposing refractory phases but attack Pt. These aspects have been discussed in detail by Claisse in Chapter 9. Acid digestion has disadvantages too. The reaction is slow compared with a fusion. However, even fusions can be time consuming. For example, for accurate analysis using isotope dilution ICP-MS, fusion salts are best removed by ion exchange before analysis [15]. Although most resistant minerals can be decomposed by acids (even zircons and chromite) there are some (e.g., cassiterite), which will not dissolve. Another disadvantage, which is often cited, is precipitation of insoluble ﬂuorides. However, when HF is used in conjunction with HClO4 or H2SO4, this is rarely a problem. Acids also have

735

P.J. Potts and P. Robinson

highly hazardous properties and appropriate laboratory safety protocols must be followed. 24.6.1 Properties of acids used in the decomposition of geological materials The common mineral acids used in the decomposition of geological materials are discussed in Refs. [11,16– 19]. The physical properties of commonly used acids have been summarized by Matusiewicz in Chapter 6. (a) Hydroﬂuoric acid: This is the main constituent of any acid decomposition procedure involving silicate rocks, readily dissolving silica to form SiF6 22 in acid solution. HF by itself is more effective than when mixed with another mineral acid. However, even though it is sometimes used alone [20], it is almost always mixed with another oxidizing, higher boiling point acid, such as HClO4, to drive off the last traces of ﬂuoride from the residue. Silicon is removed as volatile SiF4 and insoluble ﬂuorides converted to a soluble product. Even dilute solutions of HF will etch glass, so PTFE labware is normally used. It is also one of the most hazardous mineral acids in use in the laboratory causing agonizing pains some time after an accident and full safety precautions (gloves, mask, PVC apron, fume cupboard, etc.) are essential. Continuous drenching with water should be followed by calcium gluconate jelly and medical attention started immediately if an accident occurs. (b) Perchloric acid: HClO4 is one of the strongest mineral acids known and is a powerful oxidizing agent when hot and concentrated. It reacts explosively with organic compounds and thus geological samples containing organic material should ﬁrst be treated with HNO3 or an HNO3 – HClO4 mixture. Inorganic perchlorates are stable in solution but some become spontaneously inﬂammable in the anhydrous form. Perchlorate salts are very soluble except for (K, Rb and Cs). Due to the high boiling point (2038C), there is more efﬁcient attack on refractory minerals than, e.g., HNO3 (b.p. 1228C) and better removal of HF during the evaporation stage. One drawback in the use of HClO4 is that an expensive, dedicated fume cupboard is required, free from organics, equipped with fume scrubbing and wash-down facilities. Without such a facility, HClO4 cannot be used safely. (c) Nitric acid: When hot and concentrated, HNO3 is a strong oxidizing agent, which will liberate trace elements as highly soluble salts. The relatively low boiling point (1228C), however, results in long digestion times. It is useful where other acids, such as HClO4 or HCl, cause interference, e.g., polyatomic ion species in ICP-MS. HNO3 is the best acid medium for ICP-MS. It’s constituents (H2, N2 and O2) are already present in air entrained in the plasma and the range of polyatomic ions are not increased signiﬁcantly by an HNO3 matrix. Aqua regia is a fresh mixture of concentrated HNO3 and HCl in a volume ratio of 1:3. It has a much stronger oxidizing and dissolving power than

736

Sample preparation of geological samples, soils and sediments

HNO3 alone due to the presence of free Cl2 and NOCl. It must be freshly prepared. Major uses are the decomposition of silicates (mixed with HF) and sulﬁdes and for the dissolution of Au, Pt and Pd. Being a more moderate oxidizing agent than HClO4, HNO3 is also used as a preliminary treatment for samples containing easily oxidized organic material. (d) Hydrochloric acid: Concentrated hydrochloric acid is a weakly reducing acid, which dissolves carbonates, phosphates and many metal oxides. For silicate analysis, it is usually used in combination with other acids, such as HF and HNO3. Under elevated temperatures and pressures, many silicates, refractory oxides, sulfates and ﬂuorides are attacked. It dissolves acid volatile sulﬁdes and some sulfates but not pyrite or barite. In combination with HNO3 (aqua regia) or KClO3 oxidants, many sulﬁde minerals are decomposed and brought into solution. For AAS analysis, interferences are less with chlorides but for ICP-MS analysis chloride-bearing polyatomic ions (e.g., ArClþ, ClOþ and ClOHþ) cause major interferences on As and V and, to a lesser extent, on many other trace elements (Cr, Fe, Ga, Ge, Se, Ti and Zn) below m/z 80 [18]. However, since the boiling point of HCl is low (1108C), HCl can be removed by evaporation with HNO3. Remnant Cl2 ions in a ﬁnal dilute HNO3 solution do not cause signiﬁcant polyatomic interferences in ICP-MS. When HCl is used in acid digestion, there is potential loss of the volatile chlorides of Ge, As, Se, Sn, Sb and Hg. (e) Sulfuric acid: Concentrated sulfuric acid is dehydrating, mildly oxidizing and has a high boiling point (3388C), making for long evaporation times in PTFE. It forms some insoluble sulfates (particularly Ba, Ca, Pb and Sr), is viscous, thus depressing the signal in AAS, causes polyatomic interferences in ICP-MS analysis and attacks the ICP’s nickel cones. For these reasons, it has not found wide popularity in the decomposition of geological materials. Despite these disadvantages, however, it is a highly effective reagent with HF, decomposing most resistant minerals such as zircon, chromite, monazite and many naturally occurring halides (e.g., ﬂuorite). After lengthy evaporation, where the high boiling point is very effective at removing ﬂuorine, the sample is treated with HClO4, dried again and taken up in 2% HNO3 for ICP-MS analysis [21,22]. H2SO4 also decomposes As, Sb, Te, Se minerals and, when mixed with (NH4)2SO4, Nb and Ta minerals. (f) Phosphoric acid: H3PO4 has a high boiling point with low volatility and is infrequently used in whole rock analysis because phosphate ions can cause interferences by precipitating some of the elements to be determined. In ICPMS analysis, difﬁculties encountered are polyatomic species of P and erosion of nickel sampler cones. However, the acid is very useful for dissolving oxides and even chromite (FeCr2O4). Its effectiveness depends on the formation of condensed polyphosphoric acids (CPA) on heating. In this form, it can be used for the decomposition of minerals, ores and oxides that would otherwise have to be fused to be dissolved, e.g., bauxites, iron, chrome, Nb, Ta and W ores. Difﬁcult carbonates, such as siderite, are dissolved although sulﬁdes are not

737

P.J. Potts and P. Robinson

attacked unless mixed with H2SO4 or HClO4 [17]. The resulting solution is analyzed by colorimetry, AAS or ICP-AES, although there is a depressive effect on the latter two techniques. Hannaker and Hou [23] found that CPA at 2908C, either alone or, even better, mixed with HClO4, decomposed 70 natural minerals. Reaction times were less than 45 min, except for corundum, staurolite, tourmaline and zircon, which took up to 180 min. Pure acid is expensive to produce, as the low vapor pressure does not allow distillation. Either triple sublimation of P2O5 or ion exchange is needed.

24.6.2 Open vessel and low-pressure acid digestion This method refers to acid attack in open containers or screw top PTFE vials (e.g., Savillexw) placed on a hotplate. It has long been a popular and simple method for the decomposition of geological materials. HF in combination with a variety of higher boiling point acids (e.g., HNO3, HClO4 and H2SO4) attacks silicates with the formation of ﬂuorosilicic acid, which is decomposed and driven off as the volatile SiF4. SiO2 þ 6HF ! H2 SiF6 þ 2H2 O

ð24:1Þ

H2 SiF6 ! SiF4 " þ 2HF

ð24:2Þ

HF is also driven off at the higher temperature and the residue can be dissolved in dilute HNO3 or HCl acid. There are, however three main problems with this technique: 1. refractory minerals, such as zircon, chromite, rutile, monazite, garnet and magnetite, will only be partially attacked. With felsic samples, a high pressure closed “bomb” system or an alkali fusion is needed; 2. with samples larger than 100 mg, insoluble ﬂuorides can be produced which can have severe effects on trace element determinations. Croudace [24] identiﬁed ralstonite by XRD and suggested this problem could be overcome by initially evaporating the solution to dampness, not to dryness. Boer et al. [25] studied ﬂuoride precipitation in relation to rare earth elements (REE) and Yokoyama et al. [26] found 100% recovery of incompatible elements Rb, Sr, Y, Cs, Ba, REE, Pb, Th and U by using larger amounts of HClO4 and evaporating to dryness in a stepwise fashion. However, the high ﬁeld strength elements (HFSE) were lost as insoluble oxides; 3. volatile ﬂuorides, such as As, B, Ge, Sb and Se, could be lost. However, many form stable ﬂuoride complexes which help keep them in solution [27]. Jenner et al. [28], Eggins et al. [29] and Robinson et al. [30] used an HF/ HNO3 digestion in closed (low pressure), screw top PTFE vials kept at 1308C on

738

Sample preparation of geological samples, soils and sediments

a hotplate for 48 h. This procedure (described in detail below) works well for samples without refractory minerals such as basalts, carbonates (after pretreatment with dilute acid) and silicate samples containing sulﬁdes (after pretreatment with fresh aqua regia). It has the advantage of avoiding the use of HClO4, which cannot be employed in many laboratories for safety reasons unless specialized fume cupboard facilities are available. Open vessel acid digestions using HF/HClO4, HF/HClO4/HNO3 and HF/HNO3/HCl have been used extensively [26,31–39]. The high boiling point of HClO4 (2038C) compared with HF (b.p. 1128C) ensures complete removal of ﬂuorides and, with the high solubility of perchlorates, there is usually better solution of resistant phases than with HF/HNO3. However, many refractory minerals are still only partially attacked. For example, Hall and Plant [40] found the “common mixed acid attack” (HF/HClO4/HNO3/HCl) inefﬁcient and resorted to an alkali fusion for HFSE and REE. Tang et al. [41], analyzing eleven Chinese geological reference materials, found no signiﬁcant difference between HF/HNO3/HClO4, HF/aqua-regia/HClO4 and HF/HClO4 in an open acid digestion. However, results for Zr, Hf, Y, Sn and some HREE were low compared with a fusion attack. HF/H2SO4 was used in classical methods for major element separations and Fe2þ determinations but because of the formation of insoluble sulfates has gained little favor in open beaker digestions for trace element analysis. Hoops [42] digested samples in an open system with HF/H2SO4/HNO3 overnight on a steam bath using the analytical scheme outlined by Shapiro and Brannock [43], evaporating the acid mixture until SO3 fumes were evolved. XRD analysis of the insoluble residues revealed that many refractory minerals were still present. Multiple acid attacks, drying down in between acid addition, are often made with varying degrees of success. Sometimes, any residue left is ﬁltered and decomposed by alkali fusion or by acid in a closed pressure vessel. The two solutions are combined before analysis. Examples of this approach include the procedures used by Roelandts and Michel [44], Jarvis and Jarvis [45], Park and Hall [46], Sen Gupta [47], Pin and Joannan [48] and Lara et al. [49]. However, recent observations with ICP-MS analysis (e.g., Zr in granites) have shown that a clear solution does not guarantee all trace elements are in solution. In conclusion, open vessel or low-pressure screw top vial digestion is successful for samples without refractory minerals. Elements such as Zr, Hf, Cr, Sn, Mo, Sc, Ba, HREE and HFSE are often present in those minerals and recovery will be incomplete. Volatile elements, such as Se, Hg, As, Ge, Te, Re, Os and Ru, may be lost using this method [17]. Clearly, knowledge of the petrology and mineralogy of samples to be analyzed is essential before deciding whether an alternative method of attack, such as a closed system acid digestion or fusion with an alkali ﬂux, must be used.

739

P.J. Potts and P. Robinson

24.6.3 HF – HNO3 decomposition method in Savillexw screw top vials [30] This method is suitable for silicate samples such as basalts which have no refractory mineral phases. Powdered sample (100 mg) is weighed into a 7 ml Savillexw vial and moistened with ultrapure water. If sulﬁdes are present, add HCl (3 ml) and HNO3 (1 ml) and evaporate to dryness. HF (2 ml) and HNO3 (0.5 ml) are added to the sample, the vial sealed and placed on a hotplate for 48 h at 1308C. At least twice during the ﬁrst 24 h, the container is removed from the hotplate, cooled and placed in an ultrasonic bath for 2 min. Alternatively, it can be shaken by hand every few hours. After 48 h, the vials are allowed to cool and then opened and the contents evaporated to incipient dryness. The evaporation is repeated twice more after additions of 1 ml HNO3. The residue is dissolved in 2 ml HNO3 and 3–5 ml ultrapure water, heated at 60–808C overnight, transferred to a polycarbonate container and diluted to 100 ml (1000 £ dilution of sample) prior to analysis. 24.6.4 Closed vessel high pressure acid digestion Often referred to as “bombs” (cf. Chapters 6 and 8), these PTFE containers sealed inside a stainless steel jacket produce very high pressures (7 –12 MPa) when sample and acids are subjected to high temperatures (110–2508C). Under these conditions, there is considerably more efﬁcient decomposition of resistant minerals. Other advantages are that volatile elements, such as As, B, Cr, Hg, Sb, Se and Sn, remain within the vessel in solution and contamination is reduced because the sealed system excludes introduction of airborne particles during decomposition [18]. Care must be taken to avoid explosive rupturing of the vessel and samples containing organic material should not be mixed with oxidizing acids. Carbonates must be pre-treated with dilute acid to remove CO2. Digestion bombs must never be ﬁlled with liquid to more than 10–20% of their total volume. They are often used non-routinely because of their cost and potential danger. If only a small number of vessels are available, preparation of a large number of samples is also time consuming. One alternative for greater throughput is an assembly of six vessels, sometimes referred to as the “six pack” [50]. Another is the PicoTracew apparatus [51], which heats one or two PTFE coated metal blocks each holding 16 tightly sealed PTFE vessels, on a PTFE coated hotplate. Time and temperature are electronically controlled and a solid PTFE evaporation plate under vacuum allows rapid removal of acid vapors, which are absorbed in sodium hydroxide solution. Early work on geological materials [32,52] used HF and a small volume of HNO3 or aqua regia at 110 –1508C for 1–2 h. On cooling, boric acid was added to complex the HF so that major and trace elements could be determined. However, the resulting solution still attacks glassware and silica so is not

740

Sample preparation of geological samples, soils and sediments

favorable for ICP-MS nebulizers. Furthermore, the total dissolved solids content is too high for good ICP-MS analysis. For trace elements, it is better to expel the SiF4 on a hotplate after opening the vessel. Subsequent studies have shown that refractory materials as accessory minerals associated with silicate rocks take anywhere from a few hours to several days for complete decomposition and the choice of accompanying acid (e.g., HNO3, HCl, HClO4, H2SO4) is critical. An HF/HNO3 attack is often used in isotope geochemistry prior to ion exchange separations. Krogh [53] digested , 20(–50) mg samples of zircon in a bomb at 2208C for 7 days for U and Pb isotopic analysis. David et al. [54], digested samples with HF/HNO3 in screw top Savillex vials for 2 days at 1508C followed by further attack on the residue with extra acid in bombs for several days at 2208C for Zr/Hf and 176Hf/177Hf isotope studies. Mu¨nker et al. [55], in a study on the separation of Nb, Ta, Zr, Hf and Lu in rock samples, used HF/ HNO3 in Parrw bombs at 1808C. However, zircon-bearing samples were fused with Li2B4O7 prior to isotope dilution (ID) and ion exchange. There is a paucity of published data on multi-trace element analysis using HF/HNO3 in digestion bombs. Hollocher et al. [56] used digestion bombs with HF/HNO3 (7 days at 1508C), to analyze reference materials of basalt, a rhyolite obsidian and a phosphate rock. Somewhat low results were reported for Zr, Hf, Ba and HREE in some samples, although the authors considered that dissolution of Zr is not usually a problem with their technique. Diegor et al. [57] compared the use of HF/HNO3 in high pressure bombs (2008C for 3 days) and Savillex screw top containers (708C) with determinations by XRF in the analysis of soils and sediments. As expected, the bomb results were higher in Zr, Hf, Y, HREE, Nb and U than the Savillex container method, but only Nb, Pb and Ba were reported for comparison with XRF. Barium was signiﬁcantly lower compared to the XRF results due to poor dissolution of BaSO4. Liang and Gre´goire [58] reported a multi-stage HF/HNO3 dissolution wherein 26 Chinese geological reference materials were analyzed for 37 elements by ICP-MS. Samples (100 mg) were treated for 12 h at 2008C, followed by concentrated HNO3 twice and ﬁnally 40% HNO3 heated to 1408C for 3 h in the bomb. Good results were obtained for most elements, although Zr, Hf and Ba were 2–8( – 15)% lower than recommended values. As in an open vessel digestion, the use of HF/HClO4 under pressure in a closed system is more successful than HF/HNO3. It has been used by Xie and Kerrich [59], Mu¨nker et al. [55], Robinson et al. [30], Yu et al. [60] and Dulski [61]. Van Eenbergen and Bruninx [62] reported no loss of elements during decomposition. However, some refractory minerals, e.g., zircons, are not fully attacked. Dulski [61] used a multi-stage method with follow-up attacks of HCl and reported good results for Rb, Sr, Y, Cs, Ba, REE, Pb, Th and U. Maﬁc samples had good Zr and Hf values but occasional low values were obtained for felsic rocks. Yu et al. [60] found Zr and Hf recoveries in three granites of 80 –92% using a similar method. The use of HF/HClO4 is

741

P.J. Potts and P. Robinson

preferred to HF/HNO3 for trace elements in ultramaﬁc rocks where chromerich spinels are effectively decomposed [63,64], although Cr itself is often lost as chromyl chloride [65]. HF/H2SO4 attacks nearly all accessory phases (except cassiterite and barite). Its use is reported by Heinrichs and Herrmann [50], Schnetger [66], Mu¨nker [21] and Yu et al. [22]. Treatment of the ﬁnal residue with HClO4 gives a solution in which almost all trace elements can be determined. Exceptions are Sr and Pb, which can be difﬁcult at high concentrations and the precipitation of BaSO4 does not allow Ba to be determined. Evaporation times can be long (2 –5 days) as the acid boils at 3388C. However, no operator time is taken up during the evaporation and the end result is not much different in the overall time taken compared to multi-stage attacks when using other acids. Details of an HF/H2SO4 single stage and a HF/HClO4 –HCl multi-stage closed vessel digestion are described below. Even though PicoTracew apparatus is used in each case, the methods could be used with any other approved pressure digestion system.

24.6.5 HF/H2SO4 decomposition method in closed, high pressure vessels [60] Aliquots (100 mg) of powdered sample are weighed into 30 ml PTFE PicoTracew digestion containers. After wetting the sample with a few drops of ultrapure water, 3 ml HF and 3 ml H2SO4 are slowly added. After thorough mixing by shaking the PTFE containers a few times, the PTFE containers are left in the digestion block at 180 – 1908C for 16 h. The digestion mixture is then evaporated to dryness at 200 – 2108C for approximately 4 days in the evaporation block. HClO4 (1 ml) is added to the residue and dried before adding 2 ml HNO3, 1 ml HCl and 10 ml ultrapure water. The role of HCl is to stabilize Nb and Ta [21]. The residue is dissolved by warming the solution in the digestion block at 60–708C for an hour or so. After the solution becomes clear, it is transferred into a polypropylene bottle, indium added as the internal standard for HR-ICP-MS analysis and the solution is diluted to 100 ml.

24.6.6 HF/HClO4 – HCl decomposition method in closed, high pressure vessels [61] Accurately weighed 100 mg aliquots of sample powder are transferred into PTFE digestion vessels and wetted with a few drops of Milli-Q water. HF (3 ml of 23 mol l21) and HClO4 (3 ml of 12 mol l21) are added carefully, in order to avoid a vigorous reaction for samples containing carbonates. After closing the digestion vessels with PTFE lids and ﬁtting the pressure plate (gas tight sealing),

742

Sample preparation of geological samples, soils and sediments

samples are digested for 16 h at 1808C under pressure. After cooling, samples are evaporated at 1808C for approximately 4 h to near dryness, redissolved with 5 ml HCl (10 mol l21) and heated again at 1808C to incipient dryness. The moist hot residues are taken up again in 5 ml HCl (10 mol l21) and, after closing the digestion system, treated at 1308C for 12 h. After cooling, the solutions are evaporated to incipient dryness again and the hot sample cakes carefully redissolved in 2 ml HCl (10 mol l21) and 10 ml Milli-Q water (the ﬁnal 2 ml HCl could be replaced by HNO3 (16 mol l21) in ICP-MS analysis if the trace elements required suffer an interference from polyatomic Cl ions). The resulting, generally clear solutions, are poured into 50 ml volumetric ﬂasks, made up to volume with Milli-Q water and then transferred to 50 ml polyethylene bottles. Prior to ICP-MS analysis, internal standards (Ru and Re) are added and the mixtures diluted 10-fold.

24.6.7 Microwave oven digestion Decomposition with acids in a closed vessel by microwave heating allows rapid heating and cooling compared with a steel-jacketed digestion bomb. Electromagnetic radiation with a frequency of 2450 MHz is used to heat the solution and the microwaves are also absorbed by sample molecules. This increases the kinetic energy of the matrix and causes internal heating and differential polarization, which expand, agitate and rupture surﬁcial layers of the solid material, exposing fresh surfaces to acid attack [18,67,68]. A detailed description of this process and its applications has been presented in Chapter 8. The incomplete digestion of refractory minerals soon tempered early expectations of fast total digestions of geological materials. Lamothe et al. [69] tested 56 reference material samples with HF/HNO3/HCl but found minerals such as chromite, rutile, corundum, cassiterite and zircon to be resistant. Totland et al. [34] used HF/HNO3/HClO4 and noted low results for Cr, Hf and Zr in some samples. Sen Gupta and Bertrand [70,71] determined 30 elements in 25 geological reference materials using HF/HNO3/HCl with the addition of boric acid and EDTA in a second stage heating. The results compared well with recommended values except for some low HREE. However, Hf was low and Zr not reported. More recently, Yu et al. [22] used a newer microwave system with higher power (650 W) and considerably higher pressure (maximum 11 MPa). Acid mixtures of HF/HNO3 and HF/HNO3/HClO4/H2SO4 were used in tetraﬂuormethaxil vessels. Recoveries for Sc, Rb, Y, Zr, Cs, Ba, REE, Hf, Ta, Pb, Th and U were only 80– 100% for two basalts and 69– 84% for two granites. It appears that the relatively short digestion times (30 min) in a microwave system are insufﬁcient to decompose all minerals compared with the long (up to several days) digestion times used in a conventional PTFE oven-heated pressure bomb.

743

P.J. Potts and P. Robinson

Microwave ovens have found widespread use in the digestion of soils, sediments and environmental samples where easily dissolved elements are required. Sen Gupta and Bouvier [72] determined Ag, Cd, Pb, Bi, Cr, Mn, Co, Ni, Li, Be, Cu and Sb in waters, rocks, lake and stream sediments. Only Cr was low in a dunite due to incomplete dissolution of chromite. McGrath [73] used HF/HNO3/HCl in a microwave digestion for the estimation of the heavy metal (Cd, Cr, Cu, Ni, Pb and Zn) content of soils in a geochemical survey and Bettinelli et al. [74] analyzed Cd, Co, Cr, Cu, Mn, Ni, Pb and Zn in soils and sediments, also with HF/HNO3/HCl for “complete solubilization”. There are also ofﬁcial testing methods for soils, sediments and sludge (e.g., ASTM 5258-92 [75], SW-846 EPA 3050B [76] and ISO 11466 [77]) that employ acid mixtures and microwave heating. 24.6.8 Partial acid attack In mineral exploration and environmental programs involving rock, soil and stream sediment analysis, complete digestion is often unnecessary [78]. Partial acid attack using aqua regia or occasionally HNO3, without HF, is often used. Such methods are simple, fast and economical. Other advantages of aqua regia, compared with HF and fusion techniques, are that the volatile components (As, Hg, S, Sb, Se, etc.) are invariably retained and the total concentration of dissolved solids in the resulting ﬁltered solution is minimized. In the search for ore deposits where base metals and sulﬁdes are of interest, HClO4 and HClO4/ HCl/HNO3 are also used. Cyanide leach procedures are useful for gold exploration where extraction from very large samples can sometimes detect small gold anomalies. Acetic acid or dilute HCl dissolve carbonate rich samples without attacking the silicate matrix, which is of interest in sedimentological studies of carbonate mineralogy and chemical characteristics of depositional and diagenetic solutions [79]. Church et al. [80] investigated the effect of oxalic acid on over 200 minerals and compounds. Oxalic acid primarily attacks compounds formed during secondary geochemical processes, i.e., where weathering is high. Church et al. also investigated aqua regia, which will digest the primary sulﬁde phases as well as the secondary phases. Balaram et al. [81] and Bettinelli et al. [74] reported the use of aqua regia for rapid dissolution of soils and sediments, the latter listing several ofﬁcial European and US standard methods. Florian et al. [82] and Sastre et al. [83] compared the use of aqua regia with HNO3 leaches on sediments, soils, sludges and environmental samples. Elements analyzed in these studies were Ag, Cd, Co, Cr, Cu, Fe, Hg, Mn, Ni, Pb, Sb and Zn. Whilst aqua regia gave higher results for most samples, HNO3 was preferred for those environmental samples with very high (.70%) organic matter. Aqua regia extraction has been adopted as a standard method for extraction of trace elements in soil (ISO 11466 [77]).

744

Sample preparation of geological samples, soils and sediments

One problem encountered with determinations based on partial leaching of the material is that there are few recognized reference materials available that can be used for validation [84]. 24.6.9 Difﬁcult minerals Acids can decompose most refractory minerals in a steel-jacketed highpressure PTFE “bomb” system, although for some, an alkali fusion must be used. Table 24.2 gives digestion information for many common resistant minerals, which are not easily attacked in an open or low-pressure (Savillexw type screw-top container) acid digestion. Fusion techniques have been discussed in detail by Claisse in Chapter 9. Zircons have traditionally been attacked by a high ratio of HF to HNO3 in a “bomb”, often with further treatment of HClO4 to expel ﬂuorides (e.g., Ref. [53] for U and Pb isotopic studies, Refs. [85,86] for Lu–Hf and Zr–Hf geochemistry, Refs. [87,88] for REE analysis and Ref. [89] for the analysis of zircon reference samples). Temperatures of 170 –2508C and time frames of 2–8 days have been used. Grain size, the amount of sample and the nature of the zircon’s crystal structure affect the ease of decomposition. There is some doubt about HF/HNO3 (and HF/HClO4) digestions for whole rock trace element analysis in e.g., granites, unless prolonged treatment (4 –7 days) is applied. Mu¨nker [21] and Yu et al. [22] preferred an HF/H2SO4 digestion taking 16 h. An alternative is to resort to an alkali fusion [90 –92]. Pure chromite is dissolved by boiling in a mixture of H2SO4 and HClO4, H3PO4, or, for faster decomposition, a MnO2 –Li2SO4 –H2SO4 mixture [93]. However, these solutions are inadequate for determining total trace element contents in a silicate rock containing minor chromite. Typically, ultramaﬁc samples have 1–3% spinel, which is not attacked by HF/HNO3, e.g. Eggins et al.’s [29] results for Cr, Ni and Ti were low in peridotite PCC-1 and Sharma et al. [63] found the REE too low in ultramaﬁc samples. Longerich et al. [94], Ionov et al. [35] and Robinson et al. [30] used HF/HClO4 in the analysis of ultramaﬁc reference materials although Cr can be lost [65]. Sahuquillo et al. [95], however, successfully determined Cr in four river sediment reference materials using HF/HNO3/HClO4 followed by HF/HClO4, HClO4 and ﬁnally HNO3. Use of HF/H2SO4 [22] gives good results in ultramaﬁcs for all trace elements, provided care is taken to obtain good ultrapure H2SO4. HF/HClO4 or HF/H2SO4 decompose many resistant minerals in a high pressure –temperature closed system. The latter acid mixture is more effective and breaks down beryl, garnet, ilmenite, rutile, spinel, staurolite, tourmaline and the aluminum rich minerals andalusite, corundum, kyanite and sillimanite. Iron oxides and hydroxides are digested with HCl, hematite more slowly than limonite, goethite or magnetite. Tungsten minerals, scheelite

745

746

TABLE 24.2 Decomposition of common resistant minerals from Refs. [16,17,96] General chemical formula

Acid digestion (open vessel)

Acid (high pressure–temperature closed vessel)

Fusion decomposition

Andalusite Barite Bauxite, e.g., gibbsite Beryl

Al2SiO5 BaSO4 Al(OH)3

H3PO4/HCl very slowly HClO4 Polyphosphoric acid

HF/H2SO4 HF/HClO4 HCl (250 –3008C) in quartz tube HF/H2SO4

Cassiterite

SnO2

Chromite

FeCr2O4

Corundum

Al2O3

Fluorite

CaF2

K2S2O7, B2O3/Li2CO3 LiBO2, Na2CO3, NaOH LiBO2, Na2O2, Na2CO3/Na2B4O7 LiBO2, Na2O2, NH4F, NaF/H3BO3 LiBO2, Na2O2/NaOH, KOH, KHF2 Na2CO3/KClO3, Na2O2, Na2CO3/Na2B4O7 LiBO2, Na2O2, Na2CO3/Na2B4O7 LiBO2, NaOH

Garnet

e.g., almandine Fe3Al2(SiO4)3 Ca2Fe2þAl2(SiO4)3 resistant to acid Fe2O3

Hematite

Be3Al2Si6O18

H2SO4/HClO4, H3PO4/HClO4 (2908C) H3PO4/HClO4 (2908C) very slowly 7 M HNO3 slowly, 8%AlCl3/HCl, H3PO4/HClO4 Depends on garnet. HCl, boiling H2SO4, H3PO4/HClO4 HCl, slower than other Fe oxides

HCl (250 –3008C) in quartz tube HF/H2SO4, HF/HClO4 (Cr lost) HF/H2SO4 (240–2508C) HF/HNO3 þ HCl

HF/H2SO4, HF/HClO4

LiBO2, Na2CO3 þdilute HClO4

LiBO2

P.J. Potts and P. Robinson

Mineral

TABLE 24.2 (continuation) General chemical formula

Ilmenite

FeTiO3

Kyanite

Al2SiO5

H3PO4/HCl very slowly

HF/H2SO4, HF/HClO4

Magnetite Monazite

Fe3O4 (Ce,La,Th)PO4

HCl H2SO4, HClO4/HNO3, HF/HClO4

HF/H2SO4 22% HCl (4 days, 2208C)

Rutile

TiO2

Scheelite

CaWO4

Sillimanite

Al2SiO5

Spinel

MgAl2O4

Staurolite

(Fe,Mg)2(Al,Fe)9O6 (SiO4)4(O,OH)2

Sulﬁdes

Acid digestion (open vessel)

Acid (high pressure–temperature closed vessel)

Fusion decomposition

HF/H2SO4

LiBO2, Na2CO3/Na2O2, Na2O2 K2S2O7, B2O3/Li2CO3, Na2CO3/Na2B4O7 LiBO2, K2S2O7 LiBO2 (þ HCl, not HNO3), KOH, Na2CO3, Na2O2 LiBO2, K2S2O7, Na2CO3, NaOH LiBO2, alkali carbonate

HF/H2SO4 HCl, aqua regia, HNO3/H2SO4, HF/HNO3, HF/HClO4

H3PO4 (2908C)

HF/H2SO4, HCl (3008C) in quartz tube HF/H2SO4 HF/H2SO4, HF/HClO4 HF/HNO3/HClO4 (bomb retains volatiles)

LiBO2, K2S2O7, B2O3/Li2CO3 LiBO2, Na2O2, K2S2O7, Na2B4O7 LiBO2 LiBO2 after pre-oxidation with LiNO3, Na2CO3/Na2O2 continued

Sample preparation of geological samples, soils and sediments

Mineral

747

748

TABLE 24.2 (continuation) Acid digestion (open vessel)

Acid (high pressure– temperature closed vessel)

Fusion decomposition

Chalcopyrite

CuFeS2

Aqua regia

HF/HNO3/HClO4 (bomb retains volatiles)

Galena

PbS

HCl, HClO4, aqua regia slow

HF/HNO3/HClO4 (bomb retains volatiles)

Molybdenite

MoS2

Aqua regia /Br2, KClO3/HCl (þ HNO3)

HF/HNO3/HClO4 (bomb retains volatiles)

Pyrite

FeS2

Aqua regia

HF/HNO3/HClO4 (bomb retains volatiles)

Pyrrhotite

Fe12xS where x ¼ 0 2 0.125

HCl, aqua regia

HF/HNO3/HClO4 (bomb retains volatiles)

Sphalerite

ZnS

HCl, aqua regia

HF/HNO3/HClO4 (bomb retains volatiles)

LiBO2 after pre-oxidation with LiNO3, Na2CO3/Na2O2 LiBO2 after pre-oxidation with LiNO3, Na2CO3/Na2O2 LiBO2 after pre-oxidation with LiNO3, Na2CO3/Na2O2 LiBO2 after pre-oxidation with LiNO3, Na2CO3/Na2O2 LiBO2 after pre-oxidation with LiNO3, Na2CO3/Na2O2 LiBO2 after pre-oxidation with LiNO3, Na2CO3/Na2O2

P.J. Potts and P. Robinson

General chemical formula

Mineral

Mineral

General chemical formula

Acid digestion (open vessel)

Topaz

Al2(SiO4)(OH,F)2

Tourmaline

e.g., dravite NaMg3 Al6B3Si6O27(OH,F)4

Boiling H2SO4/H3PO4

Wolframite

(Fe,Mn)WO4

Xenotime

YPO4

HCl, aqua regia, HF with either HNO3, HClO4 or H2SO4 H2SO4

Zircon

ZrSiO4

Acid (high pressure– temperature closed vessel)

Fusion decomposition

Resists HF and HF/HClO4

K2S2O7, NaF/H3BO3, (Na2CO3, NaOH very slow) LiBO2, alkali carbonates (add SiO2), NH4F LiBO2, alkali carbonate

HF/HClO4, HF/H2SO4, HCl (3008C)

HF/H2SO4 (16 h), HF/HNO3 (2–7 days)

(LiBO2), KOH, Na2CO3/Na2B4O7 LiBO2/Li2B4O7, Na2O2, KHF2

Sample preparation of geological samples, soils and sediments

TABLE 24.2 (continuation)

749

P.J. Potts and P. Robinson

and wolframite, are dissolved by aqua regia or when present as minor constituents in silicate rocks, by HF/HNO3 (or HClO4). The rare earth phosphate monazite requires HClO4/HNO3 or H2SO4 and xenotime, H2SO4. Apatite dissolves in HCl. Fluorite will dissolve slowly in 7 mol l21 HNO3, 8% AlCl3 in dilute HCl or with a high pressure HF/HNO3 digestion followed by HCl. Sulﬁdes (chalcopyrite, galena, molybdenite, pyrite, pyrrhotite and sphalerite) are digested with aqua regia. Addition of an oxidizing agent, such as Br2 or KClO3, enhances the reaction. A closed vessel prevents loss of volatile elements such as As, Se and Sb. HClO4 or HF/HClO4 is recommended for barite or a LiBO2 fusion. Cassiterite is particularly insoluble and the best option is fusion with LiBO2 or Na2O2.

24.7

DECOMPOSITION BY MOLTEN SALT FUSION

24.7.1 Total fusion Fusion with alkali salts is used to decompose samples containing refractory minerals that are resistant to acids. It ensures complete quantitative attack and subsequent dissolution of all elements present in a geological sample. There are virtually no known silicate minerals that cannot be brought into solution if the appropriate ﬂux is used. Fusions do not require hazardous reagents such as HF or HClO4 or specialized apparatus such as PTFE digestion bombs. The main disadvantage with fusion for the decomposition of rocks and minerals is the high level of total dissolved solids in solution and, of course, the elements that comprise the ﬂux (Li, Na, K, B and S) cannot be determined. High salt content in the ﬁnal solution affects modern instrumental methods of analysis. For example, in ICP-MS analysis there is a decrease in instrumental sensitivity resulting in poor detection limits and the need for frequent cleaning of cones, nebulizer and spray chamber. When LiBO2 has been used, it can be difﬁcult to determine Li or B in subsequent acid digestion runs due to high background levels left in the ICP-MS. Large quantities of an element (e.g., Na, K) may cause polyatomic interferences in ICP-MS analysis [18]. High dilutions are required with fusions prior to analysis, which also results in deterioration in detection limits. Although reaction time with fusion is quick compared with acid digestion, it is labor intensive and more difﬁcult to streamline for large numbers of geological samples. However, one automated system is described by Govindaraju and Mevelle [97]. Purity of the ﬂux with regard to trace element contamination is a big issue and reagent blanks should be constantly monitored. Loss of volatile elements (e.g., Sn, Sb, Tl, Pb and Zn) has been reported in lithium borate fusions by Totland et al. [34] and Yu et al. [22].

750

Sample preparation of geological samples, soils and sediments

LiBO2, and to a lesser extent Li2B4O7, are the most common ﬂuxes replacing former widely used ﬂuxes such as Na2CO3 and NaOH. Na2O2 is a powerful oxidizing agent decomposing many refractory minerals. The general effectiveness in attacking silicate rocks increases in the order Na2CO3 , NaOH , Na2O2. K2S2O7 is useful for the decomposition of refractory oxides (e.g., rutile, ilmenite) and oxides of Nb and Ta, although not for silicates. Alkali ﬂuorides (KHF2, NH4F) are used chieﬂy for opening up refractory oxides and silicates (e.g., tourmaline, zircon, beryl, cassiterite and some niobotantalates). Table 24.2 has information on the fusion decomposition of common resistant minerals. LiBO2 decomposes all major rock forming minerals and most accessory minerals [98,99]. Cremer and Schlocker [100] and Feldman [90] studied the effect of a LiBO2 fusion followed by dilute nitric acid dissolution on many common minerals. Under the conditions used (20–100 mg samples, 15 –20 min at 9508C and with various sample:ﬂux ratios) some metal oxides and many sulﬁdes (e.g., chromite, zircon, cassiterite, galena and chalcopyrite) were less well attacked than the silicates. The addition of SiO2 facilitated decomposition of some minerals. Longer fusion times with smaller sample sizes may be required. For example, Jain et al. [88] were successful with zircon, and chromitites have been analyzed after lithium borate fusion [101]. LiBO2 is readily available as a high purity reagent and fusions are carried out in nonwetting Pt –5% Au or carbon crucibles. Examples of its use are found in Refs. [33,34,40,102,103]. Fusions were performed at 950 –10508C with sample:ﬂux ratios of 1:3– 1:5. Li2B4O7 has a higher melting point (9308C) than LiBO2 (8458C), is less active and slower to dissolve. It is better for basic (low SiO2) rocks whereas LiBO2 is preferred for acidic (high SiO2) rocks. Yu et al. [22] used Li2B4O7, digesting the melt in HF/HNO3, to remove some of the Si and B as volatile ﬂuorides so reducing the total dissolved solids in solution. Na2CO3 (m.p. 8518C) decomposes many silicate materials but does not attack some refractory minerals. Fusions are carried out for at least 1 h in platinum crucibles at 1000–12008C with a sample to ﬂux ratio of 1:3 or 1:5 for acidic rocks (,70% SiO2) and up to 1:15 for basic rocks with less than ,40% SiO2 [17]. Volatile elements, such as As, Se, Tl and Hg, are lost. Addition of an oxidizer, such as KNO3, Na2O2 or KClO3, enhances the decomposition. NaOH and Na2O2 are extremely efﬁcient at dissolving silicate rocks, and melt at much lower temperatures than do Na2CO3 ﬂuxes [11]. Commonly used together, NaOH –Na2O2 decomposes accessory phases such as zircon, cassiterite, ilmenite and monazite. A ﬂux to sample ratio of 1:4 and fusion for 15–30 min at 8008C can be used with glassy carbon crucibles, but Pt is attacked, as are Au, Ag, Ni, Fe and Zr crucibles, to a lesser extent. Na2O2 is also used for digestion prior to PGE separation and analysis.

751

P.J. Potts and P. Robinson

24.7.2 A LiBO2 fusion procedure [33,34] Samples 0.250 g are mixed carefully with 1.250 g pure LiBO2 and the powders transferred to a clean 30 ml carbon or platinum crucible. Fuse at 10508C in a mufﬂe furnace for 20 min (45 min for refractory samples, for example, those rich in zircon or chromite) and pour the hot melt into 150 ml aliquots of 0.8 mol l21 HNO3. Solutions are stirred continuously until all solids are dissolved (30 –60 min) and then ﬁltered, to remove carbon particles (if carbon crucibles are used), before diluting to 250 ml in a volumetric ﬂask. Store solutions in polypropylene bottles (1:1000 dilution) and dilute to 1:5000 for ICPMS analysis.

24.7.3 Sintering Sintering reduces the salt content in solution, the principal disadvantage of an alkali salt fusion, and the lower temperature leads to less corrosion of the decomposition vessel. Powders must be ﬁne and well mixed before heating to generally just below the melting point of the reagent. Na2O2 is the most suitable, dissolving many resistant phases (e.g., zircon, tourmaline and chromite), given sufﬁcient time. A sample-to-Na2O2 ratio of 1:4 is used with the mixture heated in a platinum crucible for 1 h at 4808C [104]. On cooling, water is carefully added, the mixture centrifuged and the cake washed twice with water, centrifuging again. Sodium salts and silica are removed and the cake dissolved in HCl acid. The technique has been used by Robinson et al. [105], Longerich et al. [94] and Yu et al. [22], who determined the REE, Sc, Y, Th and Sr. Yu et al. [22] also measured Rb, Zr, Nb, Mo, Sn, Sb, Cs, Ba, Hf, Ta, Tl, Pb and Bi in nine reference rocks, ﬁnding poor recoveries for these elements. However, Meisel et al. [106] obtained good agreement with published values in three reference rocks for Zr, Hf, Ta and Nb reporting a reproducibility of 5–10%.

24.7.4 Fire assay Fire assay is used for the fusion and separation of the precious metals, Au, Ag and the PGEs, Ru, Rh, Pd, Os, Ir and Pt [16, 107, 11, 108, 17 and 109]. Typically, a large sample (5 –100 g) is taken to avoid the “nugget effect”, and is mixed with a ﬂux of PbO, SiO2, Na2CO3, CaO, Na2B4O7, KNO3 and ﬂour and fused at 10008C in a ﬁreclay crucible. The precious metals are partitioned into a Pb button that settles to the bottom of the crucible. After cooling, this button is separated and cupelled (i.e., oxidized) in a furnace to give a noble metal prill, which can be dissolved for quantitative analysis of Au, Pt and Pd. Rhodium, Ir, Ru, and Os are partially lost [110,111]. In order to collect all the PGEs, a NiS

752

Sample preparation of geological samples, soils and sediments

button is prepared instead of a Pb one, using, for example, Li2B4O7, Na2CO3, Ni, SiO2 and S, although some loss of Au and Os can occur [112–114]. Coprecipitation with Te during dissolution of the NiS button is often used to increase yields [115–117]. Gros et al. [118] even report that such a method gave a good Os recovery. One disadvantage for samples containing low levels of PGEs and Au is the contamination caused by reagent blanks owing to the large amount of chemicals used as ﬂuxes. Nickel, produced by the carbonyl process must be used. Some workers [119,120] reduced the amount of NiS and still obtained good recoveries of PGEs in two ore reference materials, although Frimpong et al. [121] noted different yields when an ore sample was compared with a komatiite. Low blanks have been obtained by some workers and ID with NiS ﬁre assay has been used for the determination of the PGEs (Pt, Pd and Ir) and Os isotope ratios [122]. Unfortunately, Re/Os ratios cannot be obtained because NiS does not allow high precision Re analysis. This is due to variable partitioning of Re into the sulﬁde melt, as well as large and variable Re reagent blanks. NiS beads have also been analyzed by laser ablation ICP-MS [123,124] to save time and avoid losses of PGEs in the acid dissolution step. Finally, it must be borne in mind that ﬁre assaying is a skilled technique where conditions have to be varied depending on the nature of the sample and considerable experience is required to obtain good results.

24.8

PRE-CONCENTRATION AND SEPARATION PROCEDURES

Pre-concentration and separation techniques are used for two main reasons: (i)

analysis of trace elements below or near the detection limit, e.g., REE analysis by ICP-OES; (ii) removal of matrix interferences that would preclude accurate analysis, e.g., high salt content in solution and polyatomic interferences in ICP-MS analysis. Ion exchange, solvent extraction, co-precipitation, vapor generation and ﬁre assay are the main methods employed in the analysis of geological materials. Very high precision in the trace element analysis of geochemical samples can be achieved by ID-ICP-MS combined with analyte separation, e.g., S, B, Zr, Hf, Mo, Sb, W, REE, Cd, PGEs, Rb, U, Th, Ge and Sn (see review by Pin and Le Fe`vre [15]). 24.8.1 Ion exchange Ion exchange is the most widely used pre-concentration technique in geological applications, particularly for the analysis of REE, trace elements by RNAA, isotope ratios and PGEs.

753

P.J. Potts and P. Robinson

Cation exchange resin (e.g., Dowex 50W-X8) is used for the group separation of the REE, Sc and Y after a LiBO2 fusion or Na2O2 sinter. Major elements are ﬁrst removed with HCl acid followed by the Al and Ca tail, Sr and Ba using HNO3. The separation has been used prior to XRF analysis on ion exchange papers by Robinson et al. [105] and ICP-OES analysis by Crock et al. [125], Roelandts and Michel [44], Jarvis and Jarvis [45], Croudace and Marshall [126] and Rucandio [127]. Watkins and Nolan [128] and Lihareva and Delaloye [129] also separated Hf along with REE, Sc and Y. Lara et al. [49] used ICP-OES with ultrasonic nebulization, ﬂow injection (FI) and an on-line anionexchange iron separation. Boaventura et al. [130] developed both off-line and on-line ICP-OES methods for REE determination using mini-columns for separation in a continuous ﬂow system. The more sensitive ICP-MS instrumentation does not usually require prior separation of the REE although ion exchange has been used to concentrate low levels in silicate rocks [48,94]. RNAA detection limits are improved signiﬁcantly in germanium gammaray spectrometers after removal of gamma active isotopes such as 24Na, 59Fe, 60 Co, 45Sc and 50Cr due to a dramatic increase in signal to background ratio. Chemical separation schemes use ion exchange, solvent extraction and precipitation. REE and PGEs have been determined in this way (see Ref. [11]). Isotope ratio analysis by TIMS or multi-collector ICP-MS invariably requires ion exchange separation, either in applications involving the determination of the established isotopes, such as Sr, Nd and Pb [131] and U–Th [132] or in newer studies on the isotope geochemistry of Lu –Hf [133], Zr–Hf and Nb– Ta [39] and the stable isotopes of B [134], Li [135], Mg, Ca, Fe, Cu, Zn, Ge and Tl [136]. Pre-concentration is essential in the analysis of PGEs and Au, which are usually present in very low concentrations. Cation exchange procedures followed by ICP-MS analysis have been developed by Jarvis et al. [137] and Ely et al. [138] who determined Ru, Rh, Ir, Pt, Pd and Au in the 1– 10 ng g21 range typically found in unmineralized samples. Pt and Au can be low and volatile OsO4 lost in the preparation. Anion exchange resins are also used to separate PGEs but not generally as a whole group. Low recovery (75–95%) requires ID to compensate for the loss [139]. Pearson and Woodland [140] used a Carius tube digestion, ID, solvent extraction and anion-exchange separation to determine Os, Ir, Pt, Pd and Ru, and Re –Os isotopes. 24.8.2 Solvent extraction and co-precipitation Solvent extraction with 4-methylpentan-2-one (MIBK) has been used for many years in AAS analysis, particularly for Au [141], Ag [142], Mo [143], In and Te [144] were analyzed by GF-AAS after organic solvent extraction. Separation of the group Nb, Mo, Ta and W from a silicate rock matrix was achieved using Nbenzoyl N-phenylhydroxylamine (BPHA) in CHCl3 followed by ICP-MS

754

Sample preparation of geological samples, soils and sediments

analysis of pentanol solutions [145]. MIBK has been employed for Tl extraction [46] and a TOPO–MIBK extraction for Sn separation [146]. The most common co-precipitation method is that of Te for the separation of PGEs (e.g., Ref. [118]). Zirconium, Nb, Hf and Ta have been analyzed by ICPMS after cupferron precipitation [91], W and Mo were pre-concentrated from geological materials by adsorption of their 8-hydroxyquinolates on activated charcoal [147] and Bi, Se and Te were co-precipitated with La(OH)3 prior to hydride generation (HG), AAS and ICP-MS analysis [148]. For the ICP-MS determination of Cd in sediments, the Zr and Sn interferences were removed by co-precipitation with the hydrolysis compounds of Al and Fe [149]. Finally, analysis of REE, Cd, In, Tl, Th, Nb, Ta, Zr and Hf in soils and sediments was achieved with a Ti(OH)4 – Fe(OH)3 co-precipitation before ICP-MS analysis [150].

24.8.3 Vapor generation Vapor generation approaches are discussed by Cai in Chapter 19. HG by quartz tube AAS or ICP-MS is used with geological materials for the analysis of hydride-forming elements As, Bi, Sb, Se, Te, Ge and Sn. The gaseous hydride is formed at room temperature by addition of a strong reducing agent, usually NaBH4, to an acid leachate of the sample and the vapor swept into the instrument [148,151]. Interferences from transition metals, such as Ni, Cu, Co and Fe can be severe, so masking agents, ion exchange or co-precipitation are used to reduce or remove them. Selenium was determined in geochemical samples by HG-ICP-AES in combination with FI and on-line anion-exchange separation of Fe [152], by HG-AAS with standard addition [153] and in sediments by FI-HG-ICP-MS, which focuses on short reaction times and rapid separation of the gaseous products [154]. Tin was analyzed by HG-ICP-MS after removal of transition metals by anion exchange [155] and in soils after KOH fusion [156]. Mercury is determined by cold vapor generation after thermal release or acid digestion followed by reduction in solution to form elemental Hg [157,158]. A review of vapor generation as compared with electrothermal AAS is given by Tsalev [159]. Another vapor phase technique (sparging) is being used for the introduction of volatile OsO4 into MC –ICP-MS instruments for study of the Re –Os isotopic system [160].

24.9

SEQUENTIAL EXTRACTIONS AND DISSOLUTIONS

Although the main aim of sample preparation for trace element determination is to achieve total dissolution of the sample, in recent years, interest has been shown in sequential dissolutions, because of the potential of this technique to provide additional speciation information, particularly in some environmental

755

P.J. Potts and P. Robinson

and exploration applications. The principle of this type of procedure, detailed by Sahuquillo and Rauret in Chapter 39, is that a sample is progressively dissolved, ﬁrst with a relatively mild reagent, through a number of increasingly reactive stages (typically between 3 and 5 in total) until the ﬁnal stage results in complete dissolution of the sample. The solution resulting from each dissolution stage is analyzed separately. The advantage of this approach is that results provide information on the concentration of an element associated with a particular mineral phase (or group of phases). This information may facilitate an understanding of the processes involved in an environmental application (based on elemental mobility/bioavailability) that is not available from a single dissolution of a sample. 24.9.1 Procedure of Tessier et al. [161] One of the ﬁrst proponents of this approach was Tessier et al. [161] and the beneﬁts of this approach can be illustrated by examining a summary of Tessier et al.’s method for the determination of trace elements of environmental interest in soil and sediment samples. The method involves a ﬁve-stage sequential extraction in which the elements of interest (e.g., Cd, Co, Cu, Ni, Pb, Zn and Fe) are extracted using progressively more aggressive reagents from progressively more resistant minerals. Fraction 1: exchangeable: 1 g of sediment—add 8 ml of 1 mol l21 magnesium chloride solution or sodium acetate solution at pH 8.2 for 1 h at room temperature. Fraction 2: bound to carbonates: Residue from (1) is leached with 8 ml of 1 mol l21 sodium acetate solution adjusted to pH 10 with acetic acid at room temperature. Fraction 3: bound to Fe – Mn oxides: The residue from (2) is extracted with 20 ml of either 0.3 mol l21 Na2S2O4 þ 0.175 mol l21 sodium citrate þ 0.025 mol l21 citric acid or 0.04 mol l21 NH2OH·HCl in 25% v/v acetic acid (the latter extraction is performed at 968C). Fraction 4: bound to organic matter: Residue from (3) is reacted with 3 ml of 0.02 mol l21 nitric acid and 5 ml of 30% hydrogen peroxide adjusted to pH 1 with nitric acid and heated to 858C for 2 h. A second 3 ml aliquot of 30% v/v hydrogen peroxide is added and the mixture heated to 858C for 3 h. To prevent the adsorption of extracted metals onto oxidized sediment, 5 ml of 3.2 mol l21 ammonium acetate in 20% (v/v) nitric acid are added and the sample diluted to 20 ml. Fraction 5: residual (silicate phases): The residue from (4) is digested with HF/HClO4 to achieve total dissolution. Although this procedure is capable of providing a wealth of elemental speciation information, difﬁculties have been encountered in obtaining

756

Sample preparation of geological samples, soils and sediments

reproducible results when the method is implemented in different laboratories. These differences may arise from the sensitivity of the method to small changes in the concentration of reagents, temperature, contact time and other procedural effects. A detailed evaluation of this method was undertaken by Li et al. [170], speciﬁcally to enable large batches of soil samples to be processed rapidly using an ICP-OES determination. In addition to an assessment of precision and accuracy, this work demonstrated the applicability of the method to contaminated soils from an old mining region, indicating the high solubility and bioavailability of Pb in smelting regions.

24.9.2 The “BCR” method In order to develop a more robust methodology for sequential extractions, Ure et al. [162] published the results of an investigation sponsored by the BCR (the former EU Community Bureau of Reference). In this report, the simpliﬁed scheme for the sequential extraction of sediments was as follows. Step 1: 1 g sediment þ 40 ml of 0.1 mol l21 acetic acid. Shake for 16 h at room temperature. Centrifuge and decant the supernatant liquid. Step 2: Break the cake from Step 1 with a vibrating rod. Add 40 ml of 0.1 mol l21 hydroxylamine hydrochloride solution. Shake for 16 h at room temperature. Centrifuge and decant the supernatant liquid. Step 3: Break the cake from Step 1 with a vibrating rod. Add 10 ml of 8.8 mol l21 (300 mg g21) hydrogen peroxide solution. Digest at room temperature for 1 h. Continue digestion for 1 h at 858C. Evaporate to a few ml. Add a further 10 ml aliquot of hydrogen peroxide solution and heat again to 858C for 1 h. Again, evaporate to a few ml. Add 50 ml of 1 mol l21 ammonium acetate solution. Shake for 16 h at room temperature. Centrifuge and decant the supernatant liquid. Further investigations of this scheme have taken place in which the type and speed of shaking designed to keep a sediment sample in suspension has been shown to have a signiﬁcant effect on results. An evaluation of the results from two interlaboratory trials was reported by Quevauviller et al. [163] with the aim of certifying the trace element extractable contents of a sediment reference material (BCR CRM 601). There has also been interest in characterizing lake and stream sediment reference materials for element concentrations derived from partial extractions [164].

24.9.3 Selective extractions for geochemical exploration Selective extractions are also of potential interest in the analysis of trace elements in soil in geochemical exploration applications, particularly in

757

P.J. Potts and P. Robinson

investigating the presence of deeply buried mineral deposits. In a scheme developed by Hall et al. [165,166], sequential selective extractions are undertaken to identify trace elements associated with (i) adsorbed/exchangeable/carbonate phases, (ii) humic and fulvic components, (iii) crystalline iron oxides, (iv) refractory organics and sulﬁdes, and (v) residual silicate and more resistant phases. Further perspectives on the performance and application of this procedure were subsequently reported by Hall [167] and Hall and Pelchat [168]. 24.10 SUMMARY AND CONCLUSIONS As can be seen from the scope and extent of this chapter, sample dissolution for trace element determination in rocks, soils and sediments covers almost the entire gamut of analytical chemistry. Because of the important role trace elements play in advancing knowledge in the geochemical and environmental sciences, geoanalytical chemists are among the vanguard in exploiting each advance in analytical technology. Indeed, there is no evidence that the pace of advance will diminish in the future. Although new capabilities are often associated with advances in instrumentation, reliable and reproducible methods of sample dissolution are fundamental to the acquisition of high quality data. Details in this chapter are designed to give an overview of the options available to support present and future developments in this ﬁeld.

REFERENCES 1 2 3 4 5 6 7 8

9 10

11 12

758

R.S. Thorpe and G.C. Brown, The Field Description of Igneous Rocks. Wiley, Chichester, 1991. M.H. Ramsey, M. Thompson and M. Hale, J. Geochem. Explor., 44 (1992) 23. M.H. Ramsey and A. Argyraki, Sci. Total Environ., 198 (1997) 243. H. Clifton, R.F. Hunter, F.J. Swanson and R.L. Phillips, USGS Prof Paper 625-C, 1969. P.J. Potts, Analyst, 122 (1997) 1179. P.J. Potts, P.C. Webb, O. Williams-Thorpe and R. Kilworth, Analyst, 120 (1995) 1273. A. Argyraki, M.H. Ramsey and P.J. Potts, Analyst, 122 (1997) 743. UK Department of the Environment, Food and Rural Affairs and the Environment Agency. An Overview of the Development of Soil Guideline Values and Related Research, CLR7, 2002. O. Williams-Thorpe, P.J. Potts and P.C. Webb, J. Archaeol. Sci., 26 (1999) 215. R. Caesareo, C. Cappio Borlino, G. Stara, A. Brunetti, A. Castellano, G. Buccoliero, M. Marabelli, A.M. Giovagnoli, A. Gorghinian and G.E. Gigante, J. Trace Microprobe Tech., 18 (2000) 23. P.J. Potts, A Handbook of Silicate Rock Analysis. Blackie, Glasgow, 1987, 622 pp. Z. Yu, M.D. Norman and P. Robinson, Geostand. Newslett. J. Geostand. Geoanal., 27 (2003) 67.

Sample preparation of geological samples, soils and sediments 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49

In: P.J. Potts, J.F.W. Bowles, S.J.B. Reed and M.R. Cave (Eds.), Microprobe Techniques in the Earth Sciences. Chapman & Hall, London, (1995), 419pp. J.S. Watson, X-ray Spectrom., 25 (1996) 173. C. Pin and B. Le Fe`vre, Geostand. Newslett. J. Geostand. Geoanal., 26 (2002) 135. W.M. Johnson and J.A. Maxwell, Rock and Mineral Analysis, 2nd ed., Wiley, New York, 1981, 489 pp. Z. Sulcek and P. Povondra, Methods of Decomposition in Inorganic Analysis. CRC Press, Boca Raton, FL, 1989, 325 pp. I. Jarvis, in: K.E. Jarvis, A.L. Gray and R.S. Houk (Eds.), Handbook of Inductively Coupled Plasma Mass Spectrometry. Blackie, Glasgow, 1992, pp. 172– 224. T.T. Chao and R.F. Sanzolone, J. Geochem. Explor., 44 (1992) 65– 106. A. Makishima, E. Nakamura and T. Nakano, Geostand. Newslett. J. Geostand. Geoanal., 23 (1999) 7. C. Mu¨nker, Chem. Geol., 144 (1998) 23. Z. Yu, P. Robinson and P. McGoldrick, Geostand. Newslett. J. Geostand. Geoanal., 25 (2001) 199. P. Hannaker and Q.L. Hou, Talanta, 31 (1984) 1153. I.W. Croudace, Chem. Geol., 31 (1980) 153. R.H. Boer, G.J. Beukes, F.M. Meyer and C.B. Smith, Chem. Geol., 104 (1993) 93. T. Yokoyama, A. Makishima and E. Nakamura, Chem. Geol., 157 (1999) 175. R. Bock, A Handbook of Decomposition Methods in Analytical Chemistry. Blackie, London, 1979, 444 pp. G.A. Jenner, H.P. Longerich, S.E. Jackson and B.J. Fryer, Chem. Geol., 83 (1990) 133. S.M. Eggins, J.D. Woodhead, L.P.J. Kinsley, G.E. Mortimer, P. Sylvester, M.T. McCulloch, J.M. Hergt and M.R. Handler, Chem. Geol., 134 (1997) 311. P. Robinson, A.T. Townsend, Z. Yu and C. Munker, Geostand. Newslett. J. Geostand. Geoanal., 23 (1999) 31. J.P. Riley, Anal. Chim. Acta, 19 (1958) 413. F.J. Lamgmyhr and P.E. Paus, Anal. Chim. Acta, 43 (1968) 397. K.E. Jarvis, Chem. Geol., 83 (1990) 89. M. Totland, I. Jarvis and K.E. Jarvis, Chem. Geol., 95 (1992) 35. D.A. Ionov, L. Savoyant and C. Dupuy, Geostand. Newslett., 16 (1992) 311. M.M. Cheatham, W.F. Sangrey and W.M. White, Spectrochim. Acta, 48B(3) (1993) E487–E506. C.-D. Garbe-Scho¨nberg, Geostand. Newslett., 17 (1993) 81. V. Balaram, S.L. Ramesh and K.V. Anjaiah, Geostand. Newslett., 20 (1996) 71. S. Weyer, C. Mu¨nker, M. Rehka¨mper and K. Mezger, Chem. Geol., 187 (2002) 295. G.E.M. Hall and J.A. Plant, Chem. Geol., 95 (1992) 141. Y.Q. Tang, K.E. Jarvis and J.G. Williams, Geostand. Newslett., 16 (1992) 61. G.K. Hoops, Geochim. Cosmochim. Acta, 28 (1964) 405. L. Shapiro and W.W. Brannock, US Geol. Surv. Bull., 1144-A (1962) 56. I. Roelandts and G. Michel, Geostand. Newslett., 10 (1986) 135. K.E. Jarvis and I. Jarvis, Geostand. Newslett., 12 (1988) 1. C.J. Park and G.E.M. Hall, J. Anal. At. Spectrom., 3 (1988) 355. J.G. Sen Gupta, Geostand. Newslett., 18 (1994) 111. C. Pin and S. Joannan, Geostand. Newslett. J. Geostand. Geoanal., 21 (1997) 43. R. Lara, R.A. Olsina, E. Marchevsky, J.A. Ga´squez and L.D. Martinez, At. Spectrosc., 21 (2000) 172.

759

P.J. Potts and P. Robinson 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87

760

H. Heinrichs and A.G. Herrmann, Praktium der analytischen Geochemie. Springer, Berlin, 1990, 669 pp. A. Schneider, PicoTracew, Bovenden, Germany, Personal Communication, 2002. B. Bernas, Anal. Chem., 42 (1968) 1682. T.E. Krogh, Geochim. Cosmochim. Acta, 37 (1973) 485. K. David, J.L. Birck, P. Telouk and C.J. Alle`gre, Chem. Geol., 157 (1999) 1. C. Mu¨nker, S. Weyer, E. Scherer and K. Mezger, Geochem. Geophys. Geosyst. (G3), 2 (2002) 9. K. Hollocher, A. Fakhry and J. Ruiz, Geostand. Newslett., 19 (1995) 35. W. Diegor, H. Longerich, T. Abrajano and I. Horn, Anal. Chim. Acta, 431 (2001) 195. Q. Liang and D.C. Gre´goire, Geostand. Newslett. J. Geostand. Geoanal., 24 (2000) 51. Q. Xie and R. Kerrich, Chem. Geol., 123 (1995) 17. Z. Yu, P. Robinson, A.T. Townsend, C. Mu¨nker and A.J. Crawford, Geostand. Newslett. J. Geostand. Geoanal., 24 (2000) 39. P. Dulski, Geostand. Newslett. J. Geostand. Geoanal., 25 (2001) 87. A. Van Eenbergen and E. Bruninx, Anal. Chim. Acta, 98 (1978) 405. A.L. Sharma, O. Alard, S. Elhou and N.J. Pearson, J. Conf. Abstr. Goldschmidt Conf., 5(2) (2000) 914. O. Alard, unpublished PhD thesis, GEMOC, Macquarie University, Sydney, 2000. K.A. Rodgers, Mineral. Mag., 38 (1972) 882. B. Schnetger, Fresenius J. Anal. Chem., 359 (1997) 468. In: H.M. Kingston and L.B. Jassie (Eds.), Introduction to Microwave Sample Preparation, ACS Prof. Ref. Book, Am. Chem. Soc., Washington, DC, 1988, p. 263. H. Matusiewicz and R.E. Sturgeon, Prog. Anal. Spectrosc., 12 (1989) 21. P.J. Lamothe, T.L. Fries and J.J. Consul, Anal. Chem., 58 (1986) 1881. J.G. Sen Gupta and N.B. Bertrand, Talanta, 42 (1995a) 1947. J.G. Sen Gupta and N.B. Bertrand, Talanta, 42 (1995b) 1595. J.G. Sen Gupta and J.L. Bouvier, Talanta, 42 (1995) 269. D. McGrath, Talanta, 46 (1998) 439. M. Bettinelli, C. Bafﬁ, G.M. Beone and S. Spezia, At. Spectrosc., 21 (2000) 50. ASTM 5258-92, Am. Soc. for Testing and Materials, Philadelphia, PA, USA, 1992. SW-846 EPA 3050B, U.S. Environ. Protection Agency, Washington, DC, 1995. ISO 11466, Soil Quality—Extraction of Trace Elements Soluble in Aqua Regia, Berlin, Germany, 1995. T.T. Chao, J. Geochem. Explor., 20 (1984) 101. P. Robinson, Chem. Geol., 28 (1980) 135. S.E. Church, E.L. Mosier and J.M. Motooka, J. Geochem. Explor., 29 (1987) 207. V. Balaram, P.V. Sunder Raju, S.L. Ramesh, K.V. Anjaiah, B. Dasaram, C. Manikyamba, M. Ram Mohan and D.S. Sarma, At. Spectrosc., 20(4) (1999) 155. D. Florian, R.M. Barnes and G. Knapp, Fresenius J. Anal. Chem., 362 (1998) 558. J. Sastre, A. Sahuquillo, M. Vidal and G. Rauret, Anal. Chim. Acta, 462 (2002) 59. J.N. Walsh, R. Gill and M.F. Thirlwall, in: R. Gill (Ed.), Modern Analytical Geochemistry. Longman, Harlow, Essex, 1997, p. 29. P.J. Patchett and M. Tatsumoto, Contrib. Mineral. Petrol., 75 (1980) 263. K. David, J.L. Birck, P. Telouk and C.J. Alle`gre, Chem. Geol., 157 (1997) 1. D.C. Gre´goire, K.M. Ansdell, D.M. Goltz and C.L. Chakrabarti, Chem. Geol., 124 (1995) 91.

Sample preparation of geological samples, soils and sediments 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107

108 109 110 111 112 113 114 115 116 117 118 119 120 121 122 123

J.C. Jain, C.R. Neal and J.M. Hanchar, Geostand. Newslett. J. Geostand. Geoanal., 25 (2001) 229. M. Wiedenbeck, P. Alle´, F. Corfu, W.L. Grifﬁn, M. Meier, F. Oberli, A. Von Quadt, J.C. Roddick and W. Spiegel, Geostand. Newslett., 19 (1995) 1. C. Feldman, Anal. Chem., 55 (1983) 2451. G.E.M. Hall, J.C. Pelchat and J. Loop, J. Anal. At. Spectrom., 5 (1990) 339. W.T. Perkins, N.J.G. Pearce and R. Fuge, J. Anal. At. Spectrom., 7 (1992) 611. R. Chattopadhyay and M. Mistry, Anal. Chim. Acta, 295 (1994) 325. H.P. Longerich, G.A. Jenner, B.J. Fryer and S.E. Jackson, Chem. Geol., 83 (1990) 105. A. Sahuquillo, R. Rubio, G. Rauret and B. Griepink, Fresenius J. Anal. Chem., 352 (1995) 572. P. Dulski, Personal Communication, 2002. K. Govindaraju and G. Mevelle, J. Anal. At. Spectrom., 2 (1987) 615. C.O. Ingamells, Talanta, 11 (1964) 665. C.O. Ingamells, Anal. Chim. Acta, 52 (1970) 323. M. Cremer and J. Schlocker, Am. Mineral., 61 (1976) 318. P.J. Potts, C.J.B. Gowing and K. Govindaraju, Geostand. Newslett., 16 (1992) 81. M. Thompson and J.N. Walsh, Handbook of Inductively Coupled Plasma Spectrometry. Blackie, Glasgow, 1989, 316 pp. J. Carignan, P. Hild, G. Mevelle, J. Morel and D. Yeghicheyan, Geostand. Newslett. J. Geostand. Geoanal., 25 (2001) 187. T.A. Rafter, Analyst, 75 (1950) 485. P. Robinson, N.C. Higgins and G.A. Jenner, Chem. Geol., 55 (1986) 121. T. Meisel, N. Scho¨ner, V. Paliulionyte and E. Kahr, Geostand. Newslett. J. Geostand. Geoanal., 26 (2002) 53. P.E. Moloughney, Assay Methods Used in Canmet for the Determination of Precious Metal. Canada Centre for Mineral and Energy Technology, 1986, SP86-1E. G.E.M. Hall and G.F. Bonham-Carter, J. Geochem. Explor., 30 (1988) 255. R.R. Barefoot, J. Anal. At. Spectrom., 13 (1998) 1077. J.C. Van Loon and R.R. Barefoot, Determination of the Precious Metals. Wiley, New York, 1991. G.E.M. Hall and J.C. Pelchat, Chem. Geol., 115 (1994) 61. D.C. Gre´goire, J. Anal. At. Spectrom., 3 (1988) 309. R. Juvonen, E. Kallio and T. Lakomaa, Analyst, 119 (1994) 617. H.-G. Plessen and J. Erzinger, Geostand. Newslett. J. Geostand. Geoanal., 22 (1998) 187. S.E. Jackson, B.J. Fryer, W. Gosse, D.C. Healey, H.P. Longerich and D.F. Strong, Chem. Geol., 83 (1990) 119. M. Sun, J. Jain, M. Zhou and R. Kerrich, Can. J. Appl. Spectrosc., 38 (1993) 103. K. Oguri, G. Shimoda and Y. Tatsumi, Chem. Geol., 157 (1999) 189. M. Gros, J.-P. Lorand and A. Luguet, Chem. Geol., 185 (2002) 179. M. Asif and S.J. Parry, Analyst, 114 (1989) 1057. G.S. Reddi, C.R.M. Rao, T.A.S. Rao, S. Vijaya Lakshmi, R.K. Prabhu and T.R. Mahalingham, Fresenius J. Anal. Chem., 348 (1994) 350. A. Frimpong, B.J. Fryer, H.P. Longerich, Z. Chen and S.E. Jackson, Analyst, 120 (1995) 1675. G. Ravizza and D. Pyle, Chem. Geol., 141 (1997) 251. K.E. Jarvis, J.G. Williams, S.J. Parry and E. Bertalan, Chem. Geol., 124 (1995) 37.

761

P.J. Potts and P. Robinson 124 125 126 127 128 129 130 131 132 133 134 135 136 137 138 139 140 141 142 143 144 145 146 147 148 149 150 151 152 153 154 155 156 157 158 159 160 161 162

762

A.P. de S. Jorge, J. Enzweiler, E.K. Shibuya, J.E.S. Sarkis and A.M.G. Figueiredo, Geostand. Newslett. J. Geostand. Geoanal., 22 (1998) 47. J.G. Crock, F.E. Lichte and T.R. Wildeman, Chem. Geol., 45 (1984) 149. I.W. Croudace and S. Marshall, Geostand. Newslett., 15 (1991) 139. M.I. Rucandio, Fresenius J. Anal. Chem., 357 (1997) 661. P.J. Watkins and J. Nolan, Chem. Geol., 95 (1992) 131. N. Lihareva and M. Delaloye, Fresenius J. Anal. Chem., 357 (1997) 314. G.R. Boaventura, R.C. De Oliveira and R.E. Santelli, Geostand. Newslett. J. Geostand. Geoanal., 26 (2002) 63. J.D. Woodhead and J.M. Hergt, Geostand. Newslett. J. Geostand. Geoanal., 25 (2001) 261. C.-C. Shen, R. Lawrence Edwards, H. Cheng, J.A. Dorale, R.B. Thomas, S. Bradley Moran, S.E. Weinstein and H.N. Edmonds, Chem. Geol., 185 (2002) 165. J. Blichert-Toft, Geostand. Newslett. J. Geostand. Geoanal., 25 (2001) 23. S. Kasemann, A. Meixner, A. Rocholl, T. Vennemann, M. Rosner, A.K. Schmitt and M. Wiedenbeck, Geostand. Newslett. J. Geostand. Geoanal., 25 (2001) 405. Y. Nishio and S. Nakai, Anal. Chim. Acta, 456 (2002) 271. M. Rehka¨mper, M. Scho¨nba¨chler and C.H. Stirling, Geostand. Newslett. J. Geostand. Geoanal., 25 (2001) 23. I. Jarvis, M.M. Totland and K.E. Jarvis, Chem. Geol., 143 (1997) 27. J.C. Ely, C.R. Neal, J.A. O’Neill Jr. and J.C. Jain, Chem. Geol., 157 (1999) 219. M. Rehka¨mper and A.N. Halliday, Talanta, 44 (1997) 663. D.G. Pearson and S.J. Woodland, Chem. Geol., 165 (2000) 87. S. Terashima, S. Itoh and A. Ando, Geostand. Newslett., 16 (1992) 9. S. Terashima, Geostand. Newslett., 15 (1991) 195. S. Terashima, Geostand. Newslett. J. Geostand. Geoanal., 21 (1997) 93. S. Terashima, Geostand. Newslett. J. Geostand. Geoanal., 25 (2001) 127. R. Goguel, Fresenius J. Anal. Chem., 344 (1992) 326. J. Yoshinaga, A. Nakama and K. Takata, Analyst, 124 (1999) 257. G.E.M. Hall, J.-C. Pelchat and K. Nimalasiri de Silva, Analyst, 112 (1987) 631. G.E.M. Hall, A.I. MacLaurin, J.C. Pelchat and G. Gauthier, Chem. Geol., 137 (1997) 79. K. Inagaki, A. Takatsu, A. Uchiumi, A. Nakama and K. Okamoto, J. Anal. At. Spectrom., 16 (2001) 1370. D. Taicheng, C. Hangting and Z. Xianjin, J. Anal. At. Spectrom., 17 (2002) 410. G.E.M. Hall and J.-C. Pelchat, Geostand. Newslett. J. Geostand. Geoanal., 21 (1997) 85. L.D. Martinez, E. Saidman, E. Marchevsky and R. Olsina, J. Anal. At. Spectrom., 12 (1997) 487. S. Terashima and N. Imai, Geostand. Newslett. J. Geostand. Geoanal., 24 (2000) 83. C. Moor and J. Kobler, J. Anal. At. Spectrom., 16 (2001) 285. Y.-L. Feng and H. Narasaki, Talanta, 46 (1998) 1155. T.J. Hosick, R.L. Ingamells and S.D. Machemer, Anal. Chim. Acta, 456 (2002) 263. J.G. Viets and R.M. O’Leary, J. Geochem. Explor., 44 (1992) 107. S. Terashima, Geostand. Newslett., 18 (1994) 199. D.L. Tsalev, Spectrochim. Acta B, 55 (2000) 917. M. Norman, V. Bennett, M. McCulloch and L. Kinsley, J. Anal. At. Spectrom., 17 (2002) 1394. A. Tessier, P.G.C. Campbell and M. Bisson, Anal. Chem., 51 (1979) 844. A. Ure, P. Quevauviller, H. Muntau and B. Griepink, Anal. Chem., 51 (1993) 135.

Sample preparation of geological samples, soils and sediments 163 164 165 166 167 168 169 170

P. Quevauviller, G. Rauret, J.-F. Lopez-Sanchez, R. Rubio, A. Ure and H. Muntau, Sci. Total Environ., 205 (1997) 223. J. Lynch, Geostand. Newslett. J. Geostand. Geoanal., 23 (1999) 251. G.E.M. Hall, J.E. Vaive, R. Beer and M. Hoashi, J. Geochem. Explor., 56 (1996) 59. G.E.M. Hall, J.E. Vaive and MacLaurin, J. Geochem. Explor., 56 (1996) 23. G.E.M. Hall, J. Geochem. Explor., 56 (1998) 59. G.E.M. Hall and P. Pelchat, Water Air Soil Pollut., 112 (1999) 41. I. Croudace and O. Williams-Thorpe, Archaeometry, 30 (1988) 227. X. Li, B.J. Coles, M.H. Ramsey and M. Thompson, Chem. Geol., 124 (1995) 109.

763

This page intentionally left blank

Chapter 25

Sample preparation for food analysis Milan Ihnat

25.1

INTRODUCTION

Materials of interest to this chapter are, to a ﬁrst approximation, foods/foodstuffs, which include animal and plant-based materials. There is a compositional similarity, in some ways, of foods to other organic materials such as feedstuffs of animal and plant origin, animal tissues, clinical tissues and, occasionally, ﬂuids and organic waste products. Thus, we can logically extend the material coverage to include “biological materials” in general. Therefore, any analytical sample treatment methodologies developed for and applicable to a wide range of biological materials are of interest to the food analyst and are worth considering and evaluating. This range of biological materials hence encompasses foods, feeds, plants, animal tissues and ﬂuids, agricultural and clinical materials, and related materials such as water and organic wastes and is characterized by a wide range of chemical composition that inﬂuence the performance of chemical analytical measurements; thus the target generally is foods, feedstuffs and related biological materials. Publications dealing with sample treatment/preparation of these commodities for determination of trace element content were consulted for the preparation of this chapter. This list can be peripherally extended to a very brief consideration of geological materials (which relate to siliceous residues found in some plants and thus plant-based foods, and which at times need to be decomposed to release trapped, soughtafter elements). A wide range of elements occur in foods as naturally occurring and purposely added elements as well as those arising by adventitious contamination. These mineral elements can generally be classiﬁed as nutritionally essential major elements, such as Na, K, Mg, Ca, N, P; nutritionally essential minor and trace elements: B, Si, F, V, Cr, Mn, Fe, Co, Cu, Zn, As, Se, Br, Mo, Sn, I; and those regarded as toxic or with an essential/toxic Contribution 2210 from Paciﬁc Agri-Food Research Centre. Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

765

M. Ihnat

duality: F, V, Cr, Mn, Co, Ni, Zn, As, Se, Mo, Pd, Cd, Sn, Hg, Tl, Pb. The survey of sample treatment approaches presented here thus covers some of these elements. The goal of sample treatment is the quantitative release of the element of interest from the organic/inorganic matrix. This is usually associated with the concept of total quantitative decomposition, destruction and dissolution of the organic and inorganic constituents of the sample leading to the element in an aqueous inorganic solution for presentation to the determinative device for quantitative measurement of total elemental content. Although the ostensible goal of most sample treatment procedures is just that, the complete decomposition of the material for quantitative determination of total elemental content is generally achieved, reported recovery and method performance studies suggest that this is not always the case. Naturally, extraction techniques may, or most likely may not, provide quantitative extractions; these partial decomposition procedures can be considered as alternatives to total, complete decompositions for various reasons (quick, amenable to automation) when extractive recovery is judged to be adequate. Other chapters in this volume present in-depth, detailed discussion of principles and background of sample treatment related also to preparation of food materials. Those of special relevance to the total decomposition of food and related matrices are: 6 Wet Digestion Methods, 7 Dry Ashing, 8 MicrowaveBased Digestion; those of relevance to partial extraction of elements for estimation of total elemental contents or for selective extraction of speciﬁc compounds (speciation) are: 10 Supercritical Fluid Extraction, 11 Accelerated Solvent Extraction, 12 Sonication as Sample Preparation Method, 13/14 Solid Phase Extractions, 15 Chelation Solvent Extraction; as well as those dealing with trace element speciation in Section 25.4. The various sections in this chapter deal with a review of the literature; pretreatment; classiﬁcation of sample treatment methods; compilation of sample decomposition procedures for various matrices and elements; a discussion of speciﬁc method/element/matrix cases; performance of sample treatment procedures; examples of speciﬁc, recommended, sample treatment procedures; and closing remarks. For the central, main portion of this chapter (Section 25.5) containing summary tables, selection of information is from a large literature base of relevant recent literature, which also includes some older literature to cover procedures not currently in vogue but which are classical ones still possessing redeeming value. 25.2

LITERATURE

Countless thousands of analytical publications have directly dealt with or have included sample treatment as part of a greater scheme of activities.

766

Sample preparation for food analysis

There has been no let-up, and currently a constant stream of research studies into older and newly developing approaches to sample treatment applied to foods and related materials continues to appear. Together with these original works are many excellent reviews, including: entire books on sample decomposition; books including chapters or sections on or discussing sample treatment; papers and publications containing major writing; reviews devoted to sample treatment; other reviews including coverage of sample treatment. 25.2.1 Books on sample treatment, decomposition A monograph on the destruction of organic matter was published in 1970 by Gorsuch based on his pioneering investigations [1]. Comprehensive treatises by Bock (A Handbook of Decomposition Methods in Analytical Chemistry) [2] and Dolezal et al. (Decomposition Procedures in Inorganic Analysis) [3] have appeared. 25.2.2 Books including chapters, sections on or discussing sample treatment, decomposition Chapters or sections dealing with speciﬁc treatment of foods are found in Refs. [4 –12]. Those covering other biological and organic materials, which can easily ﬁnd applications to foods, are in Refs. [13– 19]. The Handbook of Reference Methods for Plant Analysis [20] devotes several chapters to the preparation of plant tissue [21–25]. More or less general discussions are presented by Cantle [26]; Koch and Koch-Dedic in their Handbuch der Spurenanlyse [27]; Stoeppler [28]; Jackwerth and Wu¨rfels [29]; Begerow and Dunemann [30]; Ostapczuk [31]; Butcher and Sneddon [32]; Lim and Jackson [33]; and Kurfu¨rst [34]. Methods for the element selenium (but also of broader applicability) have been offered by Ihnat [35], Shamberger [36], Watkinson [37] and Cooper [38]. Methods for nitrogen have been published by Drew [39], Bradstreet [40] and Gustin and Ogg [41]. 25.2.3 Reviews on sample treatment, decomposition Review articles on sample treatment and speciﬁc aspects and applications have been written by Lamble and Hill [42]; deBono et al. [43]; Matusiewicz and Sturgeon [44]; Sansoni and Panday [45]; Smith and Arsenault [46]; Ure et al. [47]; To¨lg [48]; Bendicho and de Loos-Vollebregt [49]; Hoenig [50]; Sulcek et al. [51]; Ure [52]; Burguera and Burguera [53]; Griepink and To¨lg [54]; Jackwerth and Gomiscek [55]; and Sansoni and Iyengar [56].

767

M. Ihnat

25.2.4 Other reviews including coverage of sample treatment, decomposition Other review articles on other principal topics but including sample treatment have appeared, including the most useful and comprehensive reviews on foods in Analytical Chemistry, and on clinical and biological materials, foods and beverages in the Journal of Analytical Atomic Spectrometry, the most recent being by Chang et al. on Food [57] and by Taylor et al., Atomic Spectrometry Update—Clinical and Biological Materials, Food and Beverages [58,59]; Horwitz, review of tin analysis [60]; Talmi and Feldman, review of arsenic [61]; Penrose’s coverage of arsenic [62]; Langmyhr and Wibetoe, on the direct analysis of solids [63]; Carey and Caruso’s coverage of electrothermal vaporization for sample introduction in plasma source spectrometry [64]; and Jacobs’ article addressing the determination of nitrogen in biological materials [65]. 25.2.5 Papers, publications, containing (major) writing on sample treatment Publications, which can essentially be classiﬁed as research articles, containing major contributions to sample treatment are those by: To¨lg (Selenium— analysis in biological materials) [66]; To¨lg (Problems and trends in extreme trace analysis for the elements) [67]; Olson et al. (Determination of selenium in biological materials) [68]; Stoeppler (Analytical aspects of sample collection, sample storage and sample treatment) [69]; Robards and Worsfold (Cadmium: toxicology and analysis) [70]; Ferrari and Cantanzaro (Nitrogen analysis by a continuous digestion system) [71]; Gorsuch (Radiochemical investigations in the recovery for analysis of trace elements in organic and biological materials) [72] and Food and Agriculture Organization of the United Nations (Manuals of food quality control, 2. Additives contaminants techniques) [73].

25.3

PRETREATMENT

Most often, some manipulation and preparation of the food sample is required prior to the actual sample treatment/decomposition phase, in order to bring it to suitable conﬁguration for that procedure. All necessary and desired steps taken after collection and prior to sample treatment are pooled under the term pretreatment. This generally includes washing to reduce extraneous soil and other contamination from plant materials, grinding, milling, drying and mixing. Many other processing steps and related terminology taken from the literature (see also Ref. [12]) used as sources for sample decomposition information are presented in Tables 25.2 –25.9 and are included here to provide the reader with some idea of the many and varied steps performed before the food material is attacked in the decomposition phase of the analytical method.

768

Sample preparation for food analysis

First, in a small number of instances, depending on the nature of the starting material and the analysis technique, no pretreatment is required and the sample is used as received, carefully removed from the original container. This applies, for example, to materials such as liquid milk, clear noncarbonated beverages and mineral waters, and to determinative techniques such as instrumental neutron activation analysis (INAA) and solid sampling electrothermal vaporization atomic absorption spectrometry (SS-ET-AAS). Closely following these are instances requiring only minimal pretreatment, such as the decarbonating of beer and carbonated beverages, dispersion of dry milk powder with water, the addition of ethoxy nonylphenol aqueous solution as emulsifying agent to milk followed by homogenization using ultrasonic agitation and the pressing of solids into pellets for INAA or X-ray ﬂuorescence spectrometry (XRF). A step up from this are activities as simple as washing the agricultural crop and separating it into leaves, stems and other components, draining to remove brine/sauce and preserving liquids from a canned product, and defrosting frozen fruits and vegetables. Analysis of fresh, used Brewer’s yeast requires removal of excess beer by vacuum ﬁltration; margarine is melted and the clear oil is decanted for further processing. Fruits and vegetables are washed and peeled as required, and otherwise prepared as for eating to give edible portions “as normally consumed”. With meat products, extraneous tissues, such as skin, cartilage, grit, excess fat and other non-edible parts, are trimmed away, and the meat is deboned and otherwise dissected, leaving the edible portion. Flesh is removed from shelled animals. Cutting may be done with quartz, titanium or PVC utensils to minimize contamination. The food may be cooked if the product is normally consumed cooked; diet components can be composited. Various drying procedures can follow the foregoing preparations, if required, to give dry forms for convenience in handling and for increased stability. Meat can simply be patted dry on a paper wrapper; usually, materials are air-, oven- or freeze-dried. Homogenization and grinding can be applied to fresh or dried materials. Fresh materials can be homogenized with a blender (equipped with titanium blades if mandatory for ultratrace level work), minced or subjected to ultrasonic homogenization. Edible ﬁsh ﬂesh can be ground with a household wisk. Dried products can be crushed, or ﬁnely pulverized, milled or ground with, for example, a Wiley mill, tungsten carbide mill or Tecator Cyclotec sample mill, in a microdismembrator or in an agate ball-mill. Additional grinding can be accomplished using ceramic motor and piston or a mortar and pestle. The milled or powdered product can then be mixed/blended to increase homogeneity. Materials can be bottled and stored frozen if needed and thawed before decomposition. Spiking with internal standards or analytes for recovery testing is usually done immediately prior to decomposition. Additional speciﬁc examples are provided in some of the recommended methods below and details are included in ofﬁcial methods of analysis and Ref. [12].

769

M. Ihnat

25.4

CLASSIFICATION OF SAMPLE TREATMENT METHODS

The most heavily investigated phase of sample preparation for elemental determinations has been, and continues to be, sample treatment, with a great variety of decomposition approaches and nomenclature presented in the literature. Comprehensive discussions are found in the references listed in a preceding section. The detailed classiﬁcation scheme published by Sansoni and Panday [45] was augmented and updated by extracting decomposition schemes from other publications discussing sample treatment and is presented in Table 25.1 [1,2,4–6,8,10,13–18,27,32,36,41 –44,48,50,52,53,56,58,63 –65,68, 72–80]. Over the past two or three decades, the most commonly applied decomposition procedures for foods and related biological materials have been conventional wet ashing (digestion) and dry ashing, with increased current interest in microwave-assisted wet digestion and slurry preparation. Subsets of these and other techniques used for speciﬁc applications, of more limited applicability and used more sporadically include: no treatment or direct use of liquid and solid materials (as for INAA, SS-ET-AAS), dissolution/solubilization, oxygen ﬂask combustion, dry ashing with streaming oxygen/air and conventional wet digestion in closed vessels under pressure. Additional discussion is included in sections dealing with speciﬁc entries in Tables 25.2–25.9, which provide examples of sample treatment and decomposition procedures applied to foods and related materials.

25.5

COMPILATION OF SAMPLE TREATMENT METHODS FOR FOODS

Examples of some of the principal sample treatment procedures used for preparation of foods, feedstuffs and related biological materials prior to elemental determinations are presented in Tables 25.2 –25.9. A sampling of publications over the past three decades was used for compilation of this information, including some very recent publications as well as a few of the more classical papers. Only examples are included and this is not by any means a comprehensive compilation as sample treatment appears as the main topic, or most frequently as a necessary prelude to measurement, in thousands of publications. Each table includes information related to the matrix in question, elements determined, decomposition procedures used, instrumental technique and reference (with year of publication). Entries are arranged in decreasing order of date, beginning with the most recent. A detailed listing of matrices is included to which the decomposition and analytical techniques were applied as reported, with occasional condensation. The products and materials listed include all varieties of foods, botanical materials, feedstuffs and components, human milk, food additives, human and animal tissues and ﬂuids, organic compounds, in-house control materials, interlaboratory and proﬁciency food/feed/plant materials and check samples,

770

Sample preparation for food analysis TABLE 25.1 Classiﬁcation of sample treatment methods for foods, feeds and related biological and organic materials for determination of major, minor and trace elementsa 1. No treatment or minimal treatment 1.1. No treatment, direct use of liquid and solid samples 1.2. Minimal treatment, dilution, dissolution 2. Extraction 2.1. Extraction, with water, acids, or other extractants 2.2. Deproteination, haemolysis 3. Dry ashing/(dissolution) 3.1. High temperature 3.1.1. Combustion with air/oxygen 3.1.1.1. Stationary system A. Open mufﬂe furnace, air under atmospheric pressure, without ashing aid B. Open mufﬂe furnace, air under atmospheric pressure, with ashing aid C. Combination dry ashing and wet ashing (digestion) procedures, open mufﬂe furnace air under atmospheric pressure, with or without ashing aid, followed by more extensive acid treatment D. Closed (low or high pressure), without or with ashing aid (mufﬂe furnace, hot plate, ﬂame) E. Oxygen ﬂask, normal (atmospheric) pressure F. Oxygen bomb, high pressure 3.1.1.2. Streaming system A. Oxygen/air stream or other oxidizing gases in combustion tube B. H2/O2-ﬂame in a closed, cooled system C. Electrothermal vaporization (as a means of sample introduction) 3.1.2. Pyrolytic decomposition A. Heating under inert gas, e.g. Ar, N2 B. Reduction with H2 at high temperature 3.2. Low temperatures 3.2.1. Oxygen gas plasma at 100– 1258C A. Radiofrequency electrical ﬁelds B. Microwave electrical ﬁelds 3.2.2. Laser light (laser ablation) 4. Wet ashing (digestion), conventional 4.1. High temperatures 4.1.1. Normal (atmospheric) pressure 4.1.1.1. Oxidizing mineral acids A. Nitric acid B. Sulphuric acid C. Perchloric acid D. Nitric acid/sulphuric acid E. Nitric acid/perchloric acid F. Nitric acid/perchloric/sulphuric acid continued

771

M. Ihnat TABLE 25.1 (Continuation) G. Other acid and reagent mixtures H. Kjeldahl’s method 4.1.1.2. Hydrogen peroxide A. 30% hydrogen peroxide alone but usually as an additional reagent with oxidizing acids B. 50% hydrogen peroxide/sulphuric acid 4.1.1.3. Bases (high or low temperatures) 4.1.2. Low pressure 4.1.3. Moderate pressure 4.1.4. High pressure 4.1.4.1. Oxidizing acids in Teﬂon bomb, sealed quartz vessels/tubes, other vessels 4.1.4.2. Hydrogen peroxide in bomb 4.2. Low temperatures 4.2.1. OH radicals from hydrogen peroxide/Fe2þ at 100–2208C (Fenton’s reagent) A. Open beaker B. Closed system 5. Wet ashing (digestion), high temperatures, microwave-assisted 5.1. Open vessels at normal (atmospheric pressure) 5.2. Closed vessels at moderate pressure 5.3. Closed vessels at high pressure 5.4. On-line microwave digestion techniques 6. Slurry preparation 7. Others 7.1. High temperature 7.1.1. Oxidative fusion 7.1.2. Oxidation with nitric acid vapour 7.1.3. Oxidation with ozone 7.1.4. Halogenation 7.1.5. Reduction 7.2. Low temperature 7.2.1. Enzymatic/hydrolytic decomposition 7.2.2. Radiative decomposition (glow discharge, photolytic, UV, pulsed discharge lamp), photochemical decomposition Not all decomposition procedures are commonly used for decomposition of foods and related biological materials. a Based on the classiﬁcation scheme of Sansoni and Panday [45] with additions from their text and other sources: [1, 2, 5, 6, 8, 10, 13– 18, 32, 36, 41–44, 47– 50, 52, 53, 56, 58, 63–65, 68, 72–76, 79, 294].

miscellaneous products and reference materials. Increasingly, studies include reference materials, with some method developmental work being conducted solely with such control materials. Reference materials are deﬁned by material name as well as by supplier code, and include the following sources: NIST

772

Sample preparation for food analysis TABLE 25.2 Examples of sample treatment procedures involving no treatment or minimal treatment, including dissolution, extraction (not slurry preparation) for foods, feedstuffs and related biological materials Matrix

Elements determined

Decomposition procedure

Determinative technique

Rat diets, apple leaves NIST-SRM 1515, bovine liver NIST-SRM 1577b, non-fat milk powder NIST-SRM 1549, corn bran NIST-RM 8433, milk powder IAEA-A-11, wheat bran ﬂour CSRM 12-2-04, cocoa, milk powder, ﬁsh tissue (all 3 from NIPH) Meat homogenate NIST-SRM 1546, pork/chicken LCGCRM 7002, non-fat milk powder NISTSRM 1549, orchard leaves NIST-SRM 1571 Formula milk, UHT cow milk, human milk, infant formula, natural skim milk powder BCR-CRM 063R

I

INAA, PAA Pellets sealed in PE capsules (short-time neutron and photon irradiation, INAA, photon activation); pellets sealed in silica ampoules (long-time irradiation, INAA)

H, B, C, N, Na, Cl, K, S

Seal in Teﬂon foil bag for neutron irradiation

Sr, Cd, Hg, Pb, Na, Al, Ca, Mg, Cr, Mn, Fe, Ni, Cu, Zn, Se

Wines

Pb

ICP-MS To obtain milk whey: adjust pH to 4.6 w HCl, centrifuge to ppt and separate fat and casein micelles from the whey; milk whey diluted w water Dilute w matrixET-AAS matching solution, introduce into EAAS; examples of diluents [ethanol, glucose, fructose, NaCl, HNO3, NH4H2PO4, Mg(NO3)2]

PGAA

Reference [118] (2001)

[119] (2000)

[120] (2000)

[121] (1997)

continued

773

M. Ihnat

TABLE 25.2 (continuation) Matrix

Elements determined

Decomposition procedure

CRM Riverine Waters JAC 0031, 0032 (Committee of Reference Materials of the Japan Society for Analytical Chemistry) Instant powdered milk, liquid milk, mineral water

Mg, Al, Mn, Ni, Cu, Zn, As, Cd, Pb, Li, V, Co, Rb, Sr, Mo, Sb, Cs, Ba, W, U

Direct introduction ICP-MS via conventional concentric nebulizer

Spinach NIST-SRM 1570

Determinative technique

F-AAS Prepare dispersions w ultrasound, Triton X-100 to milk and La nitrate to waters before measurement; online dilution w custom-made equipment for F-AAS (diluent Triton X-100) (Zn in milk); La nitrate (Ca, Mg, Na, K in water) INAA Al, V, Ca, Mg, Heat seal in PE Mn, Na, K, La, foil bags, neutron Ba, Co, Cr, Cs, irradiation Eu, Fe, Rb, Sb, Sc, Se, Sr, Th, Zn Na, Mg, Al, Si, Pressure shape in XRF P, S, Cl, K, Ca, a jig in a press Ti, Mn, Fe, Ni, Cu, Zn, Br, Rb, Sr, Pb

Ca, Mg, K, Na, Zn, Fe

Various plants (moor vegetation), citrus leaves NIST-SRM 1572, pine needles NIST-SRM 1575 Human milk, non-fat Ca, P, Mg, Fe, milk powder NISTCu, Zn SRM 1549 Animal feed (not Cu further described)

Dilute w HNO3, ICP-OES directly aspirate emulsiﬁed samples Directly introduce ET-AAS 1– 4 mg samples into GF

Reference [122] (1997)

[123] (1996)

[124] (1995)

[125] (1995)

[126] (1994) [127] (1994)

continued

774

Sample preparation for food analysis

TABLE 25.2 (continuation) Matrix

Elements determined

Decomposition procedure

Determinative technique

Reference

Various wines

Pb

Sample into dilution tube, HNO3, shake

[128] (1994)

Sunﬂower oil, olive oil

Al, Cr, Cu, Fe, Ni, Pb

ET-AAS (matrix modiﬁer added directly to ET-AAS) ET-AAS

All types of crude or reﬁned edible oils and fats

Pb

Butter

Cu

Foods, fruits, vegetables, beverages, miscellaneous products, spinach NIST-SRM 1570, orchard leaves NIST-SRM 1571 Bovine liver BCRCRM 185, bovine liver NIST-SRM 1577a, pork liver GBW-08551, pig kidney BCR-CRM 186, potato powder ARC/CL, spinach powder IAEA

H, B, Cl, K, Na, S, Ca, Cd

Dissolve/dilute w MIBK; MIBK w egg lecithin as matrix modiﬁer (Pb) ET-AAS Place sample in oven at 608C, shake, weigh w equilibrated matrix modiﬁer (lecithin in cyclohexane; added prior to introduction to EAAS), mix, inject Heat sample at 508C, add mixture of n-butylamine/ water (85/15), homogenize, inject Seal in Teﬂon bags PGAA fabricated in laboratory

Cd, Cu, Pb, Zn

Directly introduce sample into GF (no details)

ET-AAS

[129] (1994)

[130] (1994)

[131] (1993)

[132] (1993)

[133] (1992)

continued

775

M. Ihnat

TABLE 25.2 (continuation) Matrix

Elements determined

Decomposition procedure

Determinative technique

Reference

Corn leaves

Mg, P, S, K, Ca, Mn, Fe, Cu, Zn

XRF

[134] (1992)

Wheat ﬂour ARC/CL-WF

P

K, Ca, Na, Co, Cr, Fe, Zn, Br, Sb, Mo, Cs, Rb, Sc, Ce, Sm, La, U, Eu, Tb, Yb Zn

Anion ion exchange or ferric- precipitation, ICP-OES INAA

[135] (1991)

11 vegetable samples proposed as European reference standards

Pellets made from discs punched from leaves and also from ground leaves by pressure in a die in a Carver laboratory press Sample into decanter, H2SO4, cover ﬂask, mix at room temp, centrifuge, ﬁlter, refrigerate Sample placed in PE vial

Human milk, non-fat milk powder NIST-SRM 1549 Pineapple juice Si (polydimethylsiloxane)

Edible oils and fats, Cu, Fe, Ni soybean oil, groundnut oil, cocoa butter B 8 plant species [7 from samples from international plant sample exchange program (ISPEP) (peas,citrus,lucerne, red beet, spinach, Desmodium intortum)], high boron maize, lettuce, spinach

Dilute w Triton ET-AAS X-100 before injection into GF F-AAS Bottle, 4-methyl2-pentanone/ HCl, shake, centrifuge, collect upper layer; aspirate Direct introduction ET-AAS into GF Extract dry mater- ICP-OES ial w HF/HCl/water (5 þ 2 þ 18) in polycarbonate or PE vessels at RT on shaker, ﬁlter; extract fresh material w HF/HCl/water (5 þ 2 þ 18) in polycarbonate or PE vessels at RT on shaker, centrifuge

[136] (1991)

[137] (1991) [138] (1990)

[139] (1990) [140] (1987)

continued

776

Sample preparation for food analysis

TABLE 25.2 (continuation) Matrix

Elements determined

Decomposition procedure

Determinative technique

Reference

Cooking oils, margarines, solid shortenings Milk

Ni, Cr, Cu, Fe

ET-AAS

[141] (1983)

Fats, oils (no details)

Fe, Cu, Ni, Mn

Cox’s Orange Pippin apples

Ca

20 diff crops incl stems, leaves, heads, tops, vines of alfalfa, vegetables (asparagus, beans, beets, Brussel sprout, cabbage, corn, hops, lettuce, pea, potato, rhubarb, sorghum, spinach, Swiss chard), wheat Various plant tissues incl turnip greens, pine needles, sorghum, corn grain, clover, fruit tree leaves

Ca, Mg, K, Na, P, Zn, Mn, Cu, Fe

Disperse in MIBK at 50% w/v, warm for determination Homogenize milk, water, clearing agent zinc acetate, acetic acid, water; or potassium ferrocyanide), hexane; this removes fats and ppts proteins, centrifuge, remove aqueous layer, ﬁlter if needed Sample in volumetric ﬂask, MIBK (Cu, Mn); MIBK/HNO3 (Fe, Ni); (no/minimal treatment, dissolution/dilution) 1:1.5 apple to water mixture, conical ﬂask, HCl, boil, transfer to centrifuge tube, shake, centrifuge Sample into PE bottle, activated C, extracting solution (TCA; HOAc; NH4EDTA), shake (heat at 608C if required), ﬁlter

I

Ca, Mg, K, Mn, Cu, Zn

Sample into container, NH4EDTA, shake, let stand, shake, ﬁlter, store ﬁltrate in PP

[142] (1977)

ET-AAS

[143] (1976)

F-AES

[144] (1974)

F-AAS (Cu, Mn, Fe, Mg, K, Na, Zn); LAS (P)

[145] (1973)

F-AAS

[146] (1967)

777

778

TABLE 25.3 Examples of dry ashing/dissolution procedures without ashing aids used for foods, feedstuffs and related biological materials Elements determined

Decomposition procedure

Determinative technique

Reference

28 different foods from Western Africa (vegetables, cereal grains and products, tubers, roots and products, dried legumes, nut, seed) Foods, animal protein sources in Nigeria (shelled animals)

Ca

Porcelain crucible, ash at 4508C, HNO3, reash at 4508C, diss HCl, dil water

ICP-OES

[147] (2001)

P, Ca, Mg, Fe, Zn, Ni, Pb, Mn, Cu, Cd, Co, Cr

AAS (Ca, Mg, Fe, Zn, Ni, Pb, Mn, Cu, Cd, Co, Cr), LAS (P)

[148] (2000)

Fruits (apple, avocado, grapes, jicama, lime, lemon, mamey, mango, melon, orange, papaya, pineapple, plantain, plum, soursop), vegetables (beans, beetroot, broccoli, cactus, carrot, cauliﬂower, corn, courgette, chilli, chocho, quelte de Huaunzontle, leek, onion, pea, pepper, potato, Swiss chard, tomato, watercress, yam), herbs (camomile, coriander, goosefoot, marjoram, mint, parsley)

Na, K, Ca, P, Fe, Mg, Fe, Cu, Zn, Mn, Co, Cr, Cd, Pb

Crucible, ash to 5508C, heat until ash free of all visible traces of C, diss HCl, concentrate on steam bath, dil water Beaker, ash at 4758C, diss HCl, ﬁlter

F-AES (Na, K); LAS (Ca, P, Fe, Mg); F-AAS (Fe, Cu, Zn, Mn, Co, Cr, Cd, Pb)

[149] (1998)

M. Ihnat

Matrix

TABLE 25.3 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Tea, ground coffee, mixed human diet IAEA-H-9 Various wheat, wheat ﬂour NIST-SRM 1567a

Al

Ash at 4508C, diss HCl

ET-AAS

[150] (1997)

Fe, Mn, Pb, Cu, Co, Ni, Zn, Mg, Ca, Ba, Ba, Li, Na, K, Mg, Ca Pb, Cd, Cr, Ni, Co, Zn, Cu, Mn

Ash to 6008C to constant wt, diss HCl, ﬁlter

F-AAS (Fe, Mn, Cu, Zn, Mg, Na), ET-AAS (Ba, Co, Pb, Ni), IC (Ba, Li, Na, K, Mg, Ca)

[151] (1997)

Pt crucible, ash to 4508C, diss HNO3

F-AAS, ET-AAS

[152] (1995)

Evap dish, ash at 470– 5008C, diss HNO3/HClO4, dil water, preconc on C Crucible, ash in mufﬂe furnace to 4508C, water, evaporate, reash at 4508C, repeat until ash white/grey, HCl, evap, dissolve HNO3 Ash to 4508C, add HNO3 if required, evap, reash to get white ash

F-AAS

[153] (1995)

F-AAS, ET-AAS

[154] (1993)

F-AAS

[155] (1992)

Various edible fungi (mushrooms), new CRM (Cantharellus tubaeformis) and potato CRM from LIVSVER, bovine liver NIST-SRM 1577, wheat ﬂour NIST-SRM 1567 Vegetables (spinach, cabbage, lettuce, carrots, potato, cucumber, tomato) Liver paste, apple sauce, ﬁsh, wheat bran, milk powder, diets

Vegetables (artichokes, green beans, peas)

Co

Pb, Cd, Zn, Cu, Fe, Cr, Ni

Ca, Cu, Mg, Mn, Ni, Zn

779

continued

Sample preparation for food analysis

Matrix

780

TABLE 25.3 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Cheeses (American, mozzarella, cheddar, Romano, Parmesan), non-fat milk powder NIST-SRM 1549 Feed grains (feeds, forages, grass), rice ﬂour NIST-SRM 1568 Canned meat products (pork, ham, liver paste)

Ca, Mg, P

Vycor or Pt crucible, dry, char, ash at 5258C, diss HNO3, dil water

F-AAS (Ca, Mg) LAS (P)

[156] (1991)

Co

Silica crucible, ash to 6008C, diss HNO3

ET-AAS

[157] (1990)

Cd, Pb, Cu, Zn, Fe, Ni, Mn Al

Ash at 4508C, diss HCl, ﬁlter, dil water

F-AAS

[158] (1990)

Tall form beaker, precalcination over ﬂame, ash to 4908C, diss HCl, dil water

ICP-OES

[159] (1990)

Milk powder, additives (glucose, maltodextrin, proteins, lecithin, etc.) 756 vegetable samples in Poland Cucurbit (squash) embryos, other lipid rich seeds (various nuts, sunﬂower, castor bean, soybean) orchard leaves NIST-SRM 1571, tomato leaves NIST-SRM 1573, wheat ﬂour NIST-SRM 1567 Animal feeding stuffs

Pb, Cd Ca

Ca, Cu, Fe, Mg, Mn, K, Na, Zn

Porcelain crucible, ash at 4008C, F-AAS diss HCl, reash at 4008C, diss HCl, ﬁlter Porcelain crucible, char on hot plate, F-AAS ash to 6508C, diss HCl

[160] (1990)

Incineration dish, carbonize over ﬂame, ash to 5508C, diss HCl, heat, HCl, dil water

[162] (1986)

F-AAS

[161] (1986)

M. Ihnat

Matrix

TABLE 25.3 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Oyster NBS-SRM 1566, wheat ﬂour NBS-SRM 1567, rice ﬂour NBS-SRM 1568, orchard leaves NBS-SRM 1571, citrus leaves NBS-SRM 1572 Edible oils

K, Mg, Ca, Zn, Mn, Cu, Fe

Al foil cup, ash at 5508C, diss HCl, evap, HCl, evap, HCl

F-AAS

[163] (1986)

Cu, Fe, K, Na, Ni, Zn Cu, Mn, Zn, P

Quartz crucible, char, ash to 5008C, diss HNO3, dil HNO3 Ash defatted samples at 7508C, diss HCl/LaCl3 Ni crucible, ash to 5008C, diss HCl, ﬁlter Ash at 5008C, diss HNO3, ﬁlter, boil to hydrolyze pyrophosphate, dil water Pt, porcelain or Vycor crucible, dry, ash to 500/7008C (experimental), diss HNO3, dil water Ash at 5008C, diss HNO3/HCl

ET-AAS

[164] (1983)

F-AAS (Cu, Mn, Zn); LAS (P) F-AAS

[165] (1983)

Ovine bone, meat-bone meal, macademia nuts Vegetables (cabbage, squash, potato) Feeding stuffs and their base materials (no details) Rat tissue

Plant tissues (alfalfa, citrus leaves NBS-SRM 1572, orchard leaves NBS-SRM 1571, pine needles NBS-SRM 1575, tomato leaves NBS-SRM 1573)

Na, K P, Ca

Cr, Fe, Cd, Zn

Al, Ba, B, Ca, Cu, Fe, Mg, Mn, Mo, P, K, Na, Sr, Zn

[166] (1982)

LAS (P), F-AAS (Ca)

[167] (1980)

Gamma counting

[168] (1976)

Spark excitation direct reading emission spectroscopy

[169] (1975)

continued

Sample preparation for food analysis

Matrix

781

782 TABLE 25.3 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Very wide variety of New Zealand foods (including cereals and products, eggs, fats/oils, ﬁsh, fruits, meat, milk and products, vegetables, sugars) Corn leaf, alfalfa, wheat plant, orchard leaves NBS-SRM 1571, citrus leaves (Kenworthy)

Cr, Mn, Cu, Zn, Cd

Silica crucible, ash at 450– 5008C, diss HCl, ﬁlter

F-AAS

[170] (1975)

Ca, Fe, K, Mg, Mn, Zn, Al, B, Ba, Ca, Cu, Fe, K, Mg, Mn, Mo, P, Sr, Zn

Vycor crucible, ash to 400, 500 (satisfactory), 600, 7008C, diss HNO3, dil water; for emission evap, diss in buffer soln

Plants (mixed herbage, grass, barley, grains, hay, pine needles, Rhododendron, Swedes)

Co

Vitrosil evap dish, char on electric heater, ash at 4608C, HNO3, evap to dryness under IR, reash, diss HCl, repeat reash w HCl, diss HCl, ﬁlter

[171] (1972) F-AAS (Ca, Fe, K, Mg, Mn, Zn), direct reading arc/spark emission spectroscopy (Al, B, Ba, Ca, Cu, Fe, K, Mg, Mn, Mo, P, Sr, Zn) F-AAS [172] (1972)

M. Ihnat

Matrix

TABLE 25.4 Examples of dry ashing/dissolution procedures incorporating ashing aids used for foods, feedstuffs and related biological materials Elements determined

Decomposition procedure

Determinative technique

Reference

Daily diets

Cr, Mn, Cu, Zn, Ca, Mg

Moisten w HNO3, ash at 4508C, diss HCl

[173] (1998)

Meat

Fe, Zn

[174] (1997)

Many various foods, mixed diet NIST-RM 8431 Various edible fungi (mushrooms), new RM (Cantharellus tubaeformis) and potato CRM from LIVSVER, bovine liver NIST-SRM 1577, wheat ﬂour NISTSRM 1567 Orange juice

Cr

Preash, ash 3 £ in quartz crucible (add NH4NO3 before 2nd, 3rd), diss HCl, dil water Silanized quartz tube, MgNO3, ash to 4808C, diss HNO3, reash as many times as needed to ash completely Pt crucible, Mg(NO3)2/MgO, ash to 4508C, diss NA

F-AAS (Mn, Cu, Zn, Ca, Mg), ET-AAS (Cr) F-AAS

ET-AAS

[175] (1996)

HG-AAS

[152] (1995)

HG-AAS

[176] (1994)

Se

As

Borosilicate glass beaker, MgNO3/MgO, HNO3, dry, ash to 4508C, diss HNO3, dry, reash at 4508C, diss HCl, ﬁlter, dil HCl

continued

Sample preparation for food analysis

Matrix

783

784

TABLE 25.4 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Edible tissue of livestock and poultry

Cd, Co, Cu, Fe, Pb, Mn, Ni, Zn Cr

Mg(NO3)2, dry, ash to 500–5508C, reash to remove organic residue, diss HCl Sample or slurried cereals/grains in quartz tube, Mg(NO3)2, dry, ash to 4808C, diss NA, heat, reash at 4808C, repeat until white ash, diss HNO3

F-AAS

[177] (1992)

ET-AAS

[178] (1992)

HG-AAS, ET-AAS, ICP-OES ICP-OES

[179] (1992)

[180] (1990)

HG-AAS

[181] (1989)

ICP-MS

[182] (1988)

Foods, cola beverages, vegetable juices, corn chips, cereal, citrus leaves NIST-SRM 1572, mixed diet NIST-RM 8431 Seafood, mussel products, oyster tissue NIST-SRM 1566a Human milk, skim milk powder BCR-CRM 063, skim milk powder BCR-CRM 150, skim milk powder BCR-CRM 151 Beer, tomato leaves NIST-SRM 1573

Lettuce, peanuts, potatoes, soybeans, sweet corn, wheat, spinach NBS-SRM 1570, wheat ﬂour NBS-SRM 1567

HNO3/Mg(NO3)2/MgO, dry, ash to 4508C, HNO3, dry, reash, diss HNO3, ﬁlter, dil water Al, Ba, Cd, Cr, Quartz cup, dry, HNO3, char on hot Cu, Fe, Li, Mg, plate, ash to 4008C, repeat char/ash Mn, Ni, Pb, Zn to get white ash, diss HNO3, dil water As

As

Na, Mg, P, K, Ca, Al, Cr, Zn, Mo, Cd, Pb

Beaker, beer þ Mg(NO3)2 /MgO/water, ash to 4508C, HNO3, cool, dry, reash at 4508C to white ash, diss HCl, ﬁlter, dil HCl Ash w H2SO4 at 5008C, HNO3, reash at 5008C, acidify, diss, dil water

M. Ihnat

Matrix

TABLE 25.4 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Rat thyroid, citrus leaves NBS-SRM 1572, spinach NBS-SRM 1570, ﬁsh homogenate IAEA-MA-A2, skim milk powder BCR-CRM-63, skim milk powder BCR-CRM 150 Canned chick peas

I

Glassy carbon crucible, ZnSO4/KOH, dry by IR, heat in mufﬂe in N2 atmos to 6008C, diss water/HNO3/NaCl, centrifuge

XRF

[183] (1988)

Ca, Cl, Cu, Fe, Mg, Mn, P, K, Na, Zn

For Ca, Cu, Fe, Mg, Mn, K, Na, Zn, P: Erlenmeyer ﬂask, HNO3, boil, ﬁlter, dil water; For Cl: moisten sample w Na2CO3 soln, dry, ash at 4508C, diss water, ﬁlter, ash ﬁlter paper at 4508C, diss HNO3, pool washings, dil water Pt dish, char, ash at 500 –6008C, fuse at 10008C w Na2CO3 – Na borate mixture, diss HNO3, dil water

F-AAS (Ca, Cu, Fe, Mg, Mn, P, K, Na, Zn); Indirect F-AAS (Cl); LAS (P)

[184] (1988)

ET-AAS

[185] (1987)

Ash at 4508C, fuse w K2S2O7 in ﬂame, diss H2SO4, dil water Quartz dish, Mg(NO3)2/ethanol, dry, ash to 4508C, if C present add water/HNO3, dry, reash at 3508C, diss water/HNO3

ICP-OES

[186] (1986)

ET-AAS

[187] (1984)

Various foods (including rice, ham, apple sauce, ﬁsh, chicken, eggs, cabbage, peas, green beans, cauliﬂower, tomato, potato, beef, turkey) Milk powders Chicken, eggs, milk powder

Al

Ti Pb, Cd

continued

Sample preparation for food analysis

Matrix

785

786

TABLE 25.4 (continuation) Matrix

Elements determined

Decomposition procedure

Determinative technique

Reference

Raw agricultural crops (lettuce, corn, soybeans, potatoes, wheat, peanuts)

Pb, Cd

Quartz tall form beaker, H2SO4, evap, ash to 5008C, diss HNO3, evap to dryness, reash at 4508C, repeat if needed, diss HNO3, dil water, let insoluble matter settle Quartz or Vycor or Pyrex beaker, K2SO4/HNO3, dry, ash at 500– 5508C, add HNO3 if ash non-white due to C and dry, reash, repeat HNO3 treatment/ashing until white ash, diss HNO3, dil water, allow ppt to settle

DPASV

[188] (1982)

Pyrex tall form beaker, HNO3/H2SO4, char on hot plate, ash to 5608C, diss HCl/HNO3, dil water

F-AAS

[190] (1982)

KHSO4 into Pyrex ashing vessel w PTFE-lined screw cap, add sample, add more KHSO4, ash to 5508C w ducted O2, HNO3/H2SO4, reash, diss HCl, dil water, let silica settle, decant

LAS

[191] (1981)

DPASV [189] (1982) (at HMDE), LSASV (at a composite mercury graphite electrode) M. Ihnat

Pb, Cd Foods (green beans, apple juice, apple sauce, infant formula, ﬁsh, cereal, baby beef) bovine liver NBS-SRM 1577, rice ﬂour NBS-SRM 1568, wheat ﬂour NBS-SRM 1567, oyster tissue NBS-SRM 1566, tomato leaves NBS-SRM 1573, pine needles NBS-SRM 1575, spinach NBS-SRM 1570, tuna NBS-RM 50 Pb Vegetables (kale, lettuce, Swiss chard), spinach NBS-SRM 1570, orchard leaves NBS-SRM 1571, tomato leaves NBS-SRM 1573, pine needles NBS-SRM 1575 Co, Mo Plants (barley glumes, barley straw, lucerne, hay, clover), Bowen’s kale

TABLE 25.4 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Vegetables, dairy products, meat, ﬁsh, ﬁsh meal

Pb, Cd, Cu

F-AAS

[192] (1980)

US diets

Cr

ET-AAS

[193] (1979)

Whole milk, peas, wheat ﬂour, meat, orange juice, potatoes, cheese, sugar, lettuce, apple sauce, oyster Marine ﬁsh, crab

Cd, Cu, Pb, Zn

(a) Quartz crucible, preliminary oxidation w H2SO4, ash to 4508C, complimentary oxidation w HNO3, 2nd calcination at 7508C, diss HCl/NA (b) quartz crucible, preliminary oxidation w H2SO4, calcination at 7508C, diss HCl/HNO3; both with Herman-Moritz AV fast asher Porcelain crucible, ash to 5008C, H2SO4/H2O2, evap to dryness, reash 5008C, repeat until white ash, diss HCl Quartz tall form beaker, H2SO4, mix, dry/char, ash to 4758C, water/HNO3, evap to dryness, reash to 3008C, repeat ashing w HNO3, diss HNO3, dil water, let residue settle (do not ﬁlter) Porcelain crucible, Mg(NO3)2, preash under IR, ash to 5008C, diss HCl, dil water Pt dish, ash at 350/5008C, grind in sapphire mortar, reash, regrind ash, further steps: place ash in Pt dish w Al2(SO4)3/H2SO4, boil to diss (plus further steps to get puriﬁed ash for spectrochemical detn)

DPASV (Cd, Cu, Pb); CSDPV (Zn)

[194] (1977)

LAS

[195] (1973)

DC arc emission spectrography

[196] (1952)

Beef, rib steaks

As Al, B, Ca, Cr, Cu, Co, Fe, Pb, Mg, Mn, Mo, Ni, P, K, Si, Ag, Na, Sn, Zn

Sample preparation for food analysis

Matrix

787

788

TABLE 25.5 Examples of dry ashing procedures utilizing closed and open oxygen combustion and low temperature ashing for foods, feedstuffs and related biological materials Elements determined

Decomposition procedure

Determinative technique

Reference

Various I-containing organic compounds

I

VOL

[197] (1998)

City waste incineration ash BCR-CRM 176, estuarine sediment BCR-CRM 277, rye grass BCR-CRM 281, calcareous loam soil BCR-CRM 141, sewage sludge of industrial origin BCR-CRM 146, zinc ore concentrate BCR-CRM 108, coal ﬂy ash NISR-SRM 1633a, pine needles NIST-SRM 1575

Hg

Oxygen ﬂask, absorption in KBrO3/KBr/NaOH or (1,3-dibromo-5,5-dimethyl-2,4imidazolidenedione)/NaOH; [closed, oxygen ﬂask at atmospheric pressure] Quartz tube, into solid state burner (Wickbold combustion apparatus), heat, introduce combustion products into oxyhydrogen ﬂame, ﬂush oven w N2 or O2 w CCl4 if needed, absorb in water, dil water; [open, combustion with streaming air/oxygen]

FIAS-CVAAS

[198] (1994) M. Ihnat

Matrix

TABLE 25.5 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Mixed diets, powdered cereals, non-fat milk powder NIST-SRM 1549, apple leaves NIST-SRM 1515, peach leaves NIST-SRM 1547, citrus leaves NIST-SRM 1572, tomato leaves NIST-SRM 1573a, mixed total diet NIST-SRM 1548, mixed diet IAEA-H-9, mixed cereals ASREM-1A, ASREM-1B Fish meal, grass, hair, tobacco, sludge, milk powder, cellulose, orchard leaves NIST-SRM 1571, pine needles NIST-SRM 1575, tomato leaves NIST-SRM 1573, bovine liver NIST-SRM 1577, spinach leaves NIST-SRM 1570, oyster tissue NISR-RM 50

I

Quartz tube in train (custom-made apparatus), movable furnace, 2 ﬂowing streams of O2, puriﬁcation, trap I2 on activated charcoal, seal quartz tube w I2 under vacuum for irradation in reactor; [open, combustion with streaming air/oxygen]

Gamma counting after NAA of separated I2

[199] (1994)

Cd, Hg, Zn, Pb, Cu

(Simultaneous introduction system into ﬂame or electrically heated portion of T-tube for AAS) sample on graphite platform, into T-tube combustion system, ignite w ﬂash from lamp, sweep w air into absorption cell portion of T-tube; [open, combustion with zstreaming air/oxygen]

ET-AAS

[200] (1990)

continued

Sample preparation for food analysis

Matrix

789

790

TABLE 25.5 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Various plants, alfalfa, grass, sagebrush, lichen, tomato, wheat grain, citrus leaves NBS-SRM 1572, wheat ﬂour NBS-SRM 1567, rice ﬂour NBS-SRM 1568, spinach leaves NBS-SRM 1570, orchard leaves NBS-SRM 1571, tomato leaves NBS-SRM 1573, pine needles NBS-SRM 1575 Coffee, coal, plastics, sugar, graphite, organic compounds, tobacco, coal NBS-SRM 1632a, orchard leaves NBS-SRM 1571, bovine liver NBS-SRM 1577, pine needles NBS-SRM 1575, tomato leaves NBS-SRM 1573, spinach NBS-SRM 1570

S

Combust in stream of pure O2 at 13708C(Leco combustionIR sulfur analyzer SC-132 w balance, resistance furnace,solid-state IR detector); [open, combustion with streaming air/oxygen]

CEA

[89] (1985)

Al, As, Cd, Cr, Cu, Fe, Mn, Pb, Se, Zn, Co, Sb, Ni, V

Dry, pulverized sample in quartz decomposition vessel [quartz vessel w integrated cooling ﬁnger (Cool Plasma Asher, Anton Paar)], ash, place decomposition vessel w cooling ﬁnger on reﬂux heater, add acid or acid mixture,reﬂux (all elements, volatile and not are dissolved); [closed, low temperature, low pressure, combustion with streaming plasma of oxygen gas]

ET-AAS, F-AAS, HG-AAS (depending on element and range)

[201] (1983)

M. Ihnat

Matrix

TABLE 25.5 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Rubber, plastics, fats, wood, cellulose, grass, blood, milk powder, ﬂour, coffee, tea, protein powder, wheat ﬂour NBS-SRM 1567, rice ﬂour NBS-SRM 1568, spinach NBS-SRM 1570, orchard leaves NBS-SRM 1571, tomato leaves NBS-SRM 1573, pine needles NBS-SRM 1575, bovine liver NBS-SRM 1577 Dairy products, meat, ﬁsh, poultry, grain, cereal products, potatoes, leafy vegetables, legume vegetables, fruits, sugar products, spinach NBS-SRM 1570, oyster NBS-SRM 1566, rice ﬂour NBS-SRM 1568, bovine liver NBS-SRM 1577

B, Cr, Cu, Fe, Mn, Zn, Cd, Pb, Hg, As, Se

Cycle of one incineration (Quartz Trace-O-mat, Anton Paar): position sample in burning chamber, ﬁll cooling unit w liq N2, ignite w IR radiators and combust in stream of O2, thaw combustion products, accumulate released elements by reﬂuxing acid (HNO3), rinse and dry combustion unit; [open, combustion with streaming air/oxygen]

ICP-OES (B, Cr, Cu, Fe, Mn, Zn); ET-AAS (Cd, Pb); CV-AAS (Hg); HG-AAS (As); XRF (Se)

[202] (1981)

I

Combust sample after irradation in sealed quartz vial (custom-made quartz tube combustion/separation apparatus, tubular furnace): transfer to combustion boat, insert boat þ irradiated quartz vial into combustion tube, O2, heat w furnace at 10008C, trap I2 in hydrated manganese oxide trap, heat to expel I2 and trap I2 in silver wool trap (proceed to counting gamma); [open, combustion with streaming air/oxygen] Tin capsule, WO3, place in instrument (Carlo Erba Elemental Analyzer 1106 w modiﬁcations); [open, combustion with streaming air/oxygen]

NAA (count pure I2 in silver wool trap)

[203] (1981)

GC (EC) determination in same instrument

[204] (1979)

Inorganic and organic cpds

C, H, N, S

791

continued

Sample preparation for food analysis

Matrix

792

TABLE 25.5 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

83 species of mushrooms/fungi

Se

GC (EC)

[205] (1977)

Vegetable oils (corn, linseed, olive, peanut, safﬂower, soybean)

V

Catalytic method (LAS)

[206] (1975)

Organic and inorganic compounds

S

VOL

[207] (1975)

Reﬁned sugar, unreﬁned sugar, brown sugar, molasses

Cr

HG-AAS

[208] (1974)

Fish, ﬁsh meal

Hg

SS-MS

[209] (1973)

Chicken breast muscle tissue

Hg

Wrap in paper, ignite, combust in Schoniger O2 ﬂask, absorb HCl; [closed, oxygen ﬂask at atmospheric pressure] Combust in Scho¨niger O2 ﬂask w cellulose added to facilitate combustion, absorb in HCl; [closed, oxygen ﬂask at atmospheric pressure] Paper carrier, 500 ml thick wall combustion ﬂask, NaOH/H2O2, O2, combust, diss, dil water; [closed, oxygen ﬂask at atmospheric pressure] Ash in activated oxygen plasma low temperature asher (LTA-505, LFE Corp.), diss HCl; complete ashing if needed in furnace at 4508C; [closed, low temperature, low pressure, combustion with streaming plasma of oxygen gas] Combust in Scho¨niger O2 ﬂask, absorb in HCl; extract Hg, adsorb on graphite, dry, prepare electrode; [closed, oxygen ﬂask at atmospheric pressure] Sample in black paper wrapper, dry, combust in Erlenmeyer combustion ﬂask, absorb in HCl, ﬁlter, dil water; [closed, oxygen ﬂask at atmospheric pressure]

CV-AAS (amalgamation)

[210] (1972)

M. Ihnat

Matrix

TABLE 25.5 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Cereals, oilseeds, human hair, tobacco, coal, petroleum oils

Hg

CV-AAS

[211] (1972)

Organic cpds, Teﬂon

F

ISE

[212] (1970)

Cellulose, ﬁsh solubles, sawdust, animal blood/ serum, corn ﬂour, wheat ﬂour, grass, water, kale]

Hg

Hastalloy SS combustion crucible (Parr 43A4), Parr oxygen bomb assembly (1108), ceric ammonium sulfate, O2, combust, washings to ﬂask; [closed, oxygen bomb at elevated pressure] PE or PP bottle, combust in O2, mix w benzoic acid if high F, absorb combustion products in NaOH, dil water; [closed, oxygen ﬂask at atmospheric pressure] Irradiated sample into silica boat w carrier (custom-made combustion/separation tube, train), turn on furnaces around slivered quartz wool and Se paper, pass air/O2, adsorb volatilized Hg on Se paper; [open, combustion with streaming air/oxygen]

RNAA

[213] (1969)

continued

Sample preparation for food analysis

Matrix

793

794

TABLE 25.5 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Kale powder (Bowen)

Ag, Mo, Hg, Au, Cr, Co, Se, Fe, Zn

RNAA

[214] (1967)

Oats

Se

Silica tube, irradiated in reactor, then chemically processed; wrap in ﬁlter paper, thick walled pyrex ﬂask, O2, combust, absorb in water, rinse Pt wire w HNO3/H2O2, heat,evap to near ryness, rediss in HF, multiple ion exchange; [closed, oxygen ﬂask at atmospheric pressure] Pellet into Pt holder, purge w O2, combust in O2 ﬂask w balloon, absorb in water, dil water; [closed, oxygen ﬂask at atmospheric pressure]

LAS

[215] (1961)

M. Ihnat

Matrix

TABLE 25.6 Examples of conventional wet digestion procedures utilizing open systems and oxidizing acids (and hydrogen peroxide) for foods, feedstuffs and related biological materials Elements determined

Decomposition procedure

Determinative technique

Reference

Wheat CRM GBW 08503, hair 09101, stream sediment GBW 07312, bone meal NIST-SRM 1486, total diet NIST-SRM 1548, bovine muscle powder NIST-RM 8414, pineneedles NISR-SRM 1575, oyster NIST-SRM 1566a, coal ﬂy ash NIST-SRM 1633a Sardines in soybean oil or tomato sauce, dogﬁsh muscle NRCC-DORM-2

La, Ce, Nd, Sm, Eu, Tb, Yb, Lu in Cs, Sr, Th, U

Aluminium foil for neutron irradation; after decay dissolve in HNO3/HClO4

RNAA (La, Ce, Nd, Sm, Eu, Tb, Yb, Lu in 08503 and 09101; Cs, Sr, Th, U in 5 NIST SRMs)

[216] (2001)

Cu, Fe, Mn, Zn, Sn, Cd, Cr, Pb

For Cu, Fe, Mn, Zn, Cd, Cr, Pb: digestion tube, HNO3, predigest at 1358C until clear, HNO3/HClO4, heat at 1358C until liquor colourless, ﬁlter, dil water; for Sn: use Dabeka procedure, predigest w HNO3 at 1008C, HNO3/HClO4, heat at 1008C, HCl, heat at 1008C, ﬁlter, dil water/ KCl as ionization suppresser Beaker, HNO3/HClO4, predigest overnight at RT, heat on hot plate to 160 –1708C to HClO4 fumes, transfer, dil water

F-AAS

[217] (2001)

F-AAS (Fe, Cu, Zn, Ca, Mg); F-AES (Na, K)

[218] (2001)

Fresh cow’s milk, sterilized (UHT) cow’s milk

Fe, Cu, Zn, Na, K, Ca, Mg

795

continued

Sample preparation for food analysis

Matrix

796

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Various foods incl confectoonary products, doughnuts, cheese, chewing gum, nondairy creamer, salad dressing

Ti

ICP-OES

[219] (2000)

McIntosh apple seedling leaf and stem tissues, durum wheat ﬂour NISTRM 8436, corn bran NIST-RM 8433 Bowen’s kale, hay powder IAEA-V-10, spinach NISTSRM 1570, orchard leaves NIST-SRM 1571, apple leaves NIST- SRM 1515, spinach leaves NIST-SRM 1570a, tomato leaves NIST-SRM 1573a, oriental tobacco leaves ICHTJ CTA-OTL-1, Virginia tobacco leaves ICHTJ CTAVTL-2 Tomato leaves, corn, soybean, mixed weeds, sunﬂower, rye, vetch, apple leaves in-house control, tomato leaves NIST-SRM 1573

Fe, K, Mg, Mn, Na, Zn, Ca, Ba, Cd, Cu, Mo, Ni, Pb, Rb, Sr, Zn

Borosilicate glass tube (Tecator Digestion System), H2SO4, heat 2508C, trans into PP cent tube, dil H2SO4, if ppt forms centrifuge or rediss by shaking before detn Volumetric ﬂask dedicated to digestion, HNO3, heat to reduce vol by 1/2, HNO3/HClO4, heat to fumes of HClO4, water, clear colourless digest, dil water After irradiation, wet ash samples by methods C1, C2: C1: HNO3/HClO4 open vessel on hot plate; C2: HNO3/HClO4/HF open vessel on hot plate

F-AAS (Fe, K, Mg, Mn, Na, Zn, Ca); ICP-MS (Ba, Cd, Cu, Mn, Mo, Ni, Pb, Rb, Sr, Zn)

[116] (2000)

INAA, RNAA

[220] (2000)

Borosilicate glass digestion tube, Na2SO4/CuSO4·5H2O/ H2SO4, predigest overnight at RT, heat on hot plate until clear soln, water, dil water

VOL (N); LAS (P); F-AAS (Ca, Mg); F-AES (K)

[221] (1999)

Co

N, P, Ca, Mg, K

M. Ihnat

Matrix

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

AAFCO check samples 9531 (Gilt ﬁnisher), 9623 (cattle feed), 9724 (pet food), 9728 (horse feed)

Ca, Cu, Fe, Mg, Mn, P, K, Zn

ICP-MS

[222] (1998)

Spinach leaves and stems (22)

Mg, Ti, Cr, Mn, Al, Fe, Cu, Co, Ni, Zn, Na, K, Ba, Sr, Ca, P

Volumetric ﬂask, HNO3, boil on hot plate, dil to vol water, shake, ﬁlter if required, trans aliquot to autosampler tubes, dil HNO3 HCl/HNO3, digest, HClO4, heat to white fumes and clear soln, water, repeat digestion, dil water

[223] (1998)

Cod muscle, freezedried animal blood

Cu, Mn, Ni

ICP-OES, simultaneous (Mg, Ti, Cr, Mn, Al, Fe, Cu, Co, Ni, Zn); ICP-OES, sequential (Na, K, Ba, Sr); F-AAS (Ca); not mentioned (P) FIAS-ET-AAS

Peat, straw, raw manure, composted manure, durum wheat ﬂour NIST-RM 8436, corn bran NIST-RM 8433

Al, Ba, Ca, Cd, Cr, Cu, Fe, K, Mg, Mn, Mo, Na, Ni, P, Pb, Rb, S, Sr, Ti, V, Zn

797

Erlenmeyer ﬂask, HNO3, heat on hot plate, HClO4, heat until moist residue, diss water, dil water For F-AAS, ICP-MS: volumetric ﬂask dedicated to digestion, HNO3, heat to reduce vol by 1/2 , HNO3/ HClO4, heat to fumes of HClO4, water, clear colourless digest w traces of white siliceous residue, dil water; for ICP-AES: digestion w HNO3/ HClO4/HCl

F-AAS (Cu, Fe, K, Mg, Mn, Na, Zn, Ca); ICP-MS (Ba, Cd, Cu, Mn, Mo, Ni, Pb, Rb, Sr, Zn); ICP-OES (Al, Ba, Ca, Cr, Cu, Fe, Mg, Mn, Ni, P, S, Sr, Ti, V, Zn)

[224] (1998)

[115] (1996)

continued

Sample preparation for food analysis

Matrix

798

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

32 different fruits

Ca, Cu, Fe, Mg, Mn, K, Na, Zn

F-AAS (Ca, Cu, Fe, Mg, Mn, Zn); F-AES (K, Na)

[225] (1996)

Beech leaves BCR-CRM 100, spruce needles BCR-CRM 101, hay powder BCR-CRM 129, citrus leaves NIST-SRM 1572, tomato leaves NIST-SRM 1573, pine needles NIST-SRM 1575

N, P, K, Ca, Cu, Mg, Fe, Mn, Zn

ICP-OES

[226] (1995)

Hospital diets including rice, wheat, ﬁsh and products, meat and products, milk and egg products, fruits, miso, shoyu, sugar, salt, bovine liver NIST-SRM 1577

Be, Cr, As, Cd, Sn, Sb, Ce, Hg, Pb

Kimax test tube, HNO3, heat at 808C overnight, 50% H2O2, heat at 1008C, repeat H2O2 until digests clear, heat at 808C overnight, HCl, heat, ﬁlter, dil water, store in PE tubes Digest powdered plants by the Kjeldahl method modiﬁed by Olivier, sample in cigarette paper into Pyrex tube, H2SO4 followed by H2O2, take to about 4008C progressively on a heating block (Gerhardt Kjeldatherm 40), dil water Wet ash freeze-dried sample w HNO3/H2O2 to give clear digest, dil water

ICP-MS

[227] (1994)

M. Ihnat

Matrix

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Fish homogenate, horse kidney, bovine liver NIST-SRM 1577

Hg

CV-AAS

[228] (1994)

Nine plant samples from the International Plant– Analytical Exchange, Wageningen Agricultural University, citrus leaves NIST-SRM 1572, wheat ﬂour NIST-SRM 1567, rapeseed standards BCR RM190, RM366, RM367 Nine plant samples from the International Plant– Analytical Exchange, Wageningen Agricultural University, citrus leaves NIST-SRM 1572, wheat ﬂour NIST-SRM 1567, rapeseed standards BCR RM190, RM366, RM367

S, P, K, Ca, Mg, Fe, Zn, Cu, Mn

Erlenmeyer ﬂask, (four digestion procedures: M1: HNO3; M2: HNO3/H2SO4; M3: HNO3/HClO4; M4: HNO3/H2O2), heat at 908C, dil water; for ﬁsh homogenate digest ﬁrst w HNO3 before adding HClO4 for M3 or H2O2 for M4 Pyrex tube, fuming HNO3, predigest at RT, KNO3, digest on heating block to 4008C, HCl, mix, rewarm at 808C, water, rewarm at 808C, dil water

ICP-OES

[229] (1994)

Pyrex tube, NaOBr, evap to dryness on heating block at 2508C, continue heating, formic acid/HCl, rewarm at 808C, dil water

ICP-OES

[229] (1994)

S, P, K, Ca, Mg, Fe, Zn, Cu, Mn

799

continued

Sample preparation for food analysis

Matrix

800

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Nine plant samples from the International Plant– Analytical Exchange, Wageningen Agricultural University, citrus leaves NIST-SRM 1572, wheat ﬂour NIST-SRM 1567, rapeseed standards BCR RM190, RM366, RM367 14 ref plant samples (alfalfa CII, whole meal ﬂour BCR-CRM 189, aquatic plant BCR60, aquatic plant BCR-CRM 061,olive leaf BCR-CRM 062, sugar beet leaf CII, oak leaf CII, Lucerne ﬂour CII, hay CII, hay IAEA v-10, lettuce CII, CL NIST 1572, dianthus straw CII,ray grass CII); 2 substrates (pine bark CII, peat CII)

S, P, K, Ca, Mg, Fe, Zn, Cu, Mn

Pyrex tube, HNO3/HClO4, predigest at RT, digest on heating block to 1908C, HCl, mix, rewarm at 808C, water, rewarm at 808C, dil water

ICP-OES

[229] (1994)

Al, B, Ca, Cd, Cu, Fe, K, Mg, Mn, Na, P, Pb, S, Zn

PTFE pot (custom-made), HNO3/HF, cover, heat at 1208C overnight on heating block, H2O2, heat, repeat H2O2 twice, HNO3, heat, evap to dryness, diss HNO3, heat, dil water

ICP-OES (Al, B, Ca, Cu, Fe, K, Mg, Mn, Na, P, S, Zn); ET-AAS (Cd, Pb)

[230] (1993)

M. Ihnat

Matrix

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Forages I and II (in-house stds), cotton (AR-4401), peach (AR-4402), citrus leaves NIST-SRM 1572, wheat ﬂour NIST-SRM 1567, rice ﬂour NISTSRM 1568 Food samples (15 from 4 diff food groups), low fat milk, ice cream, cream cheese, chicken, pork, beef, bread, breakfast cereals, broccoli, carrots, peas, peaches, pineapple, apple sauce, spinach NIST-SRM 1570, bovine liver NIST-SRM 1577 Liver from rats administered radiolabelled elements

S

Digestion tube, HNO3/HClO4, digest in programmable Al block up to 3508C, evap to dryness, diss water, mix, let residue settle, take supernatant, ﬁlter

HPLC

[231] (1991)

Ca, Cu, Fe, K, Mg, Mn, Na, P, Zn

Erlenmeyer ﬂask, HNO3, predigest to 1458C on hot plate, HNO3/HClO4, heat to reﬂux, repeat addition of HNO3/HClO4 heating to complete digestion, a nearly dry translucent material remains, diss HNO3

ICP-OES

[82] (1990)

Co, Cu, Se, Zn

Digestion ﬂask, HNO3/H2SO4 (Sjo¨strand type of wet oxidation reﬂux apparatus), reﬂux procedure, to white fumes of SO3, repeat once or twice for 2 or 3 cycle digesting processes

Electrophoresis, scintillation counting

[232] (1990)

continued

Sample preparation for food analysis

Matrix

801

802

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Shell ﬁsh (clams, oysters, mussels), wheat ﬂour NBS-SRM 1567, orchard leaves NBS-SRM 1571a,b, tomato leaves NBS-SRM 1573

Pb, Cd, Cu, Cr, Ni

F-AAS, ICP-OES

[233] (1988)

Spinach NBS-SRM 1570, orchard leaves NBSSRM 1571, tomato leaves NBS- SRM 1573, citrus leaves NBSSRM 1572

Na, K, Mg, Ca, Mn, Fe, Cu, Zn

Beaker, HNO3 (þ V as internal standard), predigest overnight at RT, heat on steam bath or hot plate, evap to near dryness, do not char, HNO3, heat to 5 ml, water, heat, ﬁlter dil water, split for F-AAS/ICP-AES Sampledecomposition: Kjeldahl ﬂask, HNO3, predigest overnight at RT, heat to reduce vol by 1/2, HNO3/HClO4, heat to fumes of HCLO4, rectify any darkening w HNO3, water, heat to 3 ml, dil water, ﬁlter residue on polycarbonate ﬁlter, quantitatively recover and dil ﬁltrate for detns on both ﬁltrate and res. Residuedecomposition: residue into (polytetraﬂuoroethylene) Teﬂon beaker, HClO4/HF, reﬂux, HF, repeat reﬂux, reduce vol to 0.5 ml, dil water

F-AAS

[105] (1988) M. Ihnat

Matrix

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Plants, tomato leaves, wheat ﬂour NBS-SRM 1567, rice ﬂour NBS-SRM 1568, estuarine sediment NBS-SRM 1646

Se

FLR

[234] (1987)

A variety of plants incl barley, clover, medic, pea straw, wheat ﬂour, wheat straw, Orchard leaves NIST-SRM 1571

P, K, S, Ca, Mg, Na, Al, Cu, Zn, Mn, Fe, Co, Mo, B

Digestion tube, HNO3/H3PO4, heat on aluminium block at 508C, reﬂux at 145–1508C, H2O2, stand, MnSO4, heat to reduce vol and colour of digest turns purple, repeat HNO3/H2O2 if required, (further preparation for Se detn: add HOAc/HCl, heat…) Tecator digestion system (1) HNO3, predigest overnight at RT, heat at 1208C then 1408C, continue until 1 ml acid remaining, dil HNO3; (2) HNO3, heat at 908C, then at 1408C continue until 1 ml acid remaining, dil HNO3; (3) HNO3, heat at 1408C, continue until 1 ml acid remaining, dil HNO3; (4) HNO3/HClO4 predigest overnight at RT, heat at 1208C then at 1758C, if char add HNO3, continue until 2 ml acid remaining, heat at 2258C to ensure complete dissolution, dil HNO3. In all digestions let silica settle, decant

ICP-OES

[83] (1987)

803

continued

Sample preparation for food analysis

Matrix

804

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Lobster hepatopancreas NRCC-TORT-1, marine sediment NRCC-BCSS-1, marine sediment NRCC-MESS-1

Sn

ET-AAS; ET-AAS; ID-ICP-MS

[85] (1987)

Spinach NBS-SRM 1570, orchard leaves NBS-SRM 1571, tomato leaves NBS-SRM 1573, citrus leaves NBSSRM 1572

Na, K, Mg, Ca, Mn, Fe, Cu, Zn

(1) (TORT-1) sample into PFA beaker, HNO3, heat on hot plate at 1908C to reﬂux, HF/HClO4, evap to near dryness, HNO3/HClO4, repeat to near dryness, HClO4, again heat to near dryness, diss HCl, dil water Sampledecomposition: Kjeldahl ﬂask, HNO3, predigest overnight at RT, heat to reduce vol by 1/2, HNO3/HClO4, heat to fumes of HCLO4, rectify any darkening w HNO3, water, heat to 3 ml, dil water, ﬁl residue on polycarbonate ﬁlter, quantitatively recover and dil ﬁltrate for detns on both ﬁltrate and res. Residuedecomposition: residue into (polytetraﬂuoroethylene) Teﬂon beaker, HClO4/HF, reﬂux, HF, repeat reﬂux, reduce vol to 0.5 ml, dil water

F-AAS

[104] (1987) M. Ihnat

Matrix

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Baby food samples: baby rice, baby grits, baby grits with apple, baby mix with vegetables, baby grits with chocolate and nuts, baby mix with vegetable and chicken, milk powder for infants, wheat ﬂour NBS-SRM 1567, rice ﬂour NBS-SRM 1568 Many various foods incl grain products, vegetables, dried fruits, seeds/nuts, ﬁsh/shellﬁsh, cocoa, beverages, supplement tablets Oyster NBS-SRM 1566, bovine liver NBS-SRM SRM 1577a, curly kale (in-house collaborative material)

Pb, Cd, Cu

Silica vessel (custom altered), HNO3, heat in block at 608C w reﬂux column, HClO4, heat at 2108C, cool to 608C, cap vessel, evap w vac/IR, diss HCl, heat, dil water

ASV

[235] (1987)

Ni

Wet digest HNO3/H2O2

ET-AAS

[236] (1986)

Cd, Pb, Mo, V

Boiling tube, HNO3/H2SO4, char on aluminium heating block at 1808C, H2O2, heat to remove HNO3/organic matter, dil water (add matrix modiﬁers for E-AAS and internal standards for ICP-MS)

ICP-MS (Cd, Mo, Pb, V); ET-AAS (Cd, Pb)

[237] (1986)

continued

Sample preparation for food analysis

Matrix

805

806

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Cereals, meat, ﬁsh, fats, fruit, root vegetables, other vegetables, beverages, milk, apple, potato, dried milk, tuna NBS-RM 50, bovine liver NBS-SRM 1577, wheat ﬂour NBS-SRM 1567, oyster tissue NBS-SRM 1566, spinach NBS-SRM 1570, Bowen’s kale Wheat ﬂour NBS-SRM 1567, bovine liver NBS-SRM 1577a, oyster tissue NBS-SRM 1566, spinach NBS-SRM 1570

Li, Rb, Sr

Wet digest to destroy org matter w HNO3/ H2SO4

F-AAS

[238] (1985)

Fe

Beaker, HNO3/H2SO4, boil on hot plate until charring, H2O2, heat to clear soln, dil HCl, proceed w extraction for ID-TIMS Thick-walled Pyrex digestion tube, HNO3/HClO4/H2SO4, predigest at RT, heat in digestor (Tecator digestion system) w program to 2008C, dil water

ID-TIMS

[239] (1985)

F-AAS (Cu, Zn); HG-AAS (As, Se); CV-AAS (Hg); ET-AAS (Cd, Pb)

[240] (1985)

Animal liver, kidney, bovine liver NBS-SRM 1577a

As, Cd, Cu, Hg, Pb, Se, Zn

M. Ihnat

Matrix

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

6 infant formulas: 2 powders, 2 ready-to-feed, 2 concentrates

Ca, Cu, Fe, Mg, Mn, P, K, Na, Zn

ICP-OES

[241] (1984)

Spinach NBS-SRM 1570, orchard leaves NBS-SRM 1571, tomato leaves NBS-SRM 1573

Na, K, Mg, Ca, Mn, Fe, Cu, Zn

Kjeldahl ﬂask, HNO3/HClO4, predigest overnight, heat on heating mantle at low T, boil, boil-off HNO3/water, heat, if charring cool, HNO3, heat on high heat (not to dryness), dil water Sample decomposition: Kjeldahl ﬂask, HNO3, predigest overnight at RT, heat to reduce vol by 1/2, HNO3/HClO4, heat to fumes of HCLO4, rectify any darkening w HNO3, water, heat to 3 ml, dil water, ﬁl residue on polycarbonate ﬁlter, quantitatively recover and dil ﬁltrate for detns on both ﬁltrate and res. Residue decomposition: residue into (polytetraﬂuoroethylene) Teﬂon beaker, HClO4/HF, reﬂux, HF, repeat reﬂux, reduce vol to 0.5 ml, dil water

F-AAS

[102] (1982)

continued

Sample preparation for food analysis

Matrix

807

808

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Orchard leaves NBSSRM 1571, tomato leaves NBS-SRM 1573, spinach NBS-SRM 1570, pine needles NBS-SRM 1575, Brewer’s yeast NBS-SRM 1569, oyster tissue NBS-SRM 1566, bovine liver NBS-SRM 1577, tuna NBS-RM 50, wheat ﬂour NBS-SRM 1567, rice ﬂour NBS-SRM 1568 Fresh vegetables (peas), canned goods (bamboo shoots, plums, baked beans), fresh seafood (ﬁsh) Fish, ﬁsh liver, ﬁsh kidney, orchard leaves NBSSRM 1571, bovine liver NBS-SRM 1577, ﬁsh ﬂesh IAEA-MA-M-2

Cd, Co, Cr, Mo, Ni, V, Cu, Mn, Pb, Zn, Al, Fe, As, Se, Sb

(1) Kjeldahl ﬂask, HNO3/HClO4, digest to past HClO4 stage, dil water (2) Kjeldahl ﬂask, HNO3/HClO4/H2SO4, digest to past HClO4 stage, distill-off HClO4, heat to fumes of SO3, dil water; postdigestion treatment of insol residue w HF done in some cases

ICP-OES (Cd, Co, Cr, Mo, Ni, V, Cu, Mn, Pb, Zn, Al, Fe); HG-AAS (As, Se, Sb)

[242] (1982)

Pb, Hg, Cd, Tl

Evaporating dish, HNO3/H2SO4, heat to destroy all organic matter, water, evap to 10 ml, neut w NH4OH for extraction Calibrated digestion tube, HNO3/H2SO4, predigest at RT, heat in aluminium hot block on hot plate at 608C, HNO3, heat to 120/1508C, H2O2, heat, repeat H2O2/heat until clear, dil water Kjeldahl ﬂask, H2SO4 and acid-washed sand, predigest at RT, heat to boil, boil, HClO4/ H2SO4, heat to clear digest, dil water

Amperometric titration (Pb, Cd, Hg, Tl); F-AAS (Pb, Cd, Pb, Tl) F-AAS

[243] (1982)

IUFRO plant samples used for interlaboratory comparison, standard Bowen’s kale, pine needles

Cr, Cu, Zn, Cd, Ni, Pb

N, P, K, Ca, Mg, Fe, Mn (Na)

VOL (N); F-AAS (Mg, Ca); F-AES (K); LAS (P)

[244] (1980)

[245] (1979)

M. Ihnat

Matrix

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

7 diff bakery products, various foods, ﬂounder, orchard leaves NBS-SRM 1571, bovine liver NBS-SRM 1577, tuna NBS-RM 50, Bowen’s kale, ﬂour IAEAV-2/1, oyster IAEA-MA-M-1

As, Se

HG-AAS

[246] (1977)

Haddock, perch, ﬂounder, cod, eggs, chicken liver, orchard leaves NBS-SRM 1571, bovine liver NBS-SRM 1577, tuna NBS-RM 50 Variety of foods representative of the 6 major classes: vegetable, fruit, cereal, dairy products, meat, ﬁsh including skim milk powder, corn cereal, meat

As, Se, Sb, Te

Kjeldahl ﬂask, HNO3/HClO4, predigest overnight at RT or over low heat, heat to reduce vol by 1/2, HNO3/H2SO4, heat, avoid charring by addition of HNO3 as needed, heat to white fumes of HClO4, heat to white fumes of SO3 and soln becomes clear/colourless, trans w water/HCl, dil water Kjeldahl ﬂask, HNO3/H2SO4/ HClO4, heat gently, boil, if charring add HNO3, boil-off excess HNO3, heat to fumes SO3, dil HCl/water Kjeldahl ﬂask containing water, HNO3, predigest overnight at RT, heat, HClO4/H2SO4, heat, avoid charring, HNO3 as needed to reduce charring, heat to white fumes, H2O2, heat to white fumes, repeat H2O2 2 £ , water, transfer w water for further determinative steps V2O5 þ sample into digestion ﬂask, assemble apparatus (custom-made), HNO3, heat, H2SO4 dropwise, HNO3 dropwise, 50% H2O2, ﬁlter

HG-AAS

[247] (1976)

FLR

[81] (1974)

CV-AAS

[248] (1974)

Beef liver, beef kidney, porcine liver, porcine kidney, animal feeds, ﬁsh

Se

Hg

809

continued

Sample preparation for food analysis

Matrix

810

TABLE 25.6 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Dried ﬁsh, spinach, oyster, lettuce, potatoes, dried corn, apples, milk

Zn, Pb, Cd, Cu, Co, Mn, Ni

F-AAS

[249] (1973)

Brewer’s yeast (beer-yeast slurry)

Cu, Fe, Zn, Ca, Mg, K, Na

F-AAS

[250] (1971)

Various animal tissues and products (liver, bone, brain, milk), various vegetable tissues and products (apple, cocoa powder, pepper, oil)

Elements determined ‘not reported’

Beaker, V2O5/HNO3, predigest at RT, heat on hot plate, add HNO3 and repeat if required until sample is completely in solution, HNO3/H2SO4, evap until charring begins, 50% H2O2, dil water (proceed w separation for detn) Heavy wall test tube, H2SO4/H2O2, multiple additions of H2O2, heat to fumes of SO3, dil water Pyrex glass vessel (lipless beaker which can be covered closed), water/HNO3/H2SO4, heat on hot plate at 310 –3508C until sample has dispersed, evap, HNO3 to moisten, cover and leave on hot plate until after residue becomes dry, repeat HNO3 addition heating to past dryness until residue is whitish with dark patches, fuming HNO3 in place of HNO3 and continue until white ash, dil water

Determinative method ‘not reported’

[99] (1954)

M. Ihnat

Matrix

Sample preparation for food analysis

(National Institute of Standards and Technology, Standard Reference Materials Programme, USA), IAEA (International Atomic Energy Agency), LGC (Laboratory of the Government Chemist), IRMM/BCR (Institute for Reference Materials and Measurements, Belgium), NIES (National Institute for Environmental Studies, Japan), ARC/CL (Agricultural Research Centre of Finland), ICHTJ (Institute of Nuclear Chemistry and Technology, Poland), NRCC (National Research Council of Canada), GBW (National Research Centre for Certiﬁed Reference Materials, China), CII (Le Comite InterInstituts d’Etude des Techniques Analytiques de Diagnostic Foliaire), CSRM (Pb-anal, Kosice, Slovakia), CZIM (Slovak Institute of Metrology), NIPH (Czech National Institute of Public Health), LIVSVER (Swedish National Food Administration), JAC (Committee of Reference Materials of the Japan Society for Analytical Chemistry). The column depicting elements determined is self-explanatory. A point form, abbreviated format summary of the decomposition procedure, including speciﬁc details, appears in the decomposition procedure column. Occasionally, additional details of the decomposition procedure (e.g. open, atmospheric pressure; closed, moderate pressure) are included to differentiate the procedure from others in the same table and under the same major decomposition heading. Although most such entries should be self-explanatory, several examples may be instructive. As examples, (1) if the entry reads: dilution tube, HNO3, shake, this means: introduce the weighed sample into a dilution tube, add HNO3 (speciﬁed volume of speciﬁed concentration), shake; (2) if the entry reads: 1:1.5 apple to water mixture, conical ﬂask, HCl, boil, transfer to centrifuge tube, shake, centrifuge, this means: prepare a 1:1.5 apple to water mixture, introduce a weighed sample into a conical ﬂask, add HCl (speciﬁed volume of speciﬁed concentration), boil, transfer contents to a centrifuge tube, shake, centrifuge; (3) if the entry reads: porcelain crucible, ash to 5008C, H2SO4/H2O2, evap to dryness, reash 5008C, repeat until white ash, diss HCl, this means: place a weighed sample into a porcelain crucible, ash (generally) in a mufﬂe furnace (under a program raising temperature to) 5008C, add H2SO4/H2O2 (speciﬁed volume of speciﬁed concentration), evaporate to dryness, reash at 5008C, repeat reagent addition and ashing until a white ash is obtained, dissolve the ash in HCl (speciﬁed volume of speciﬁed concentration) usually by heating; (4) if the entry reads: High pressure asher, HPA (Anton Parr), 70 ml HPA quartz vessel, HNO3, evap to dryness on hot plate at 1008C, HNO3, digest in HPA according to T –P program, this means: the weighed sample is placed into 70 ml high pressure asher, HPA (Anton Parr) quartz vessel, HNO3 is added (speciﬁed volume of speciﬁed concentration), evaporate to dryness on a hot plate at 1008C, add HNO3 (speciﬁed volume of speciﬁed concentration), digest in HPA according to a speciﬁed temperature –pressure program; (5) if the entry reads: medium pressure Teﬂon vessel (Milestone MV), HNO3/H2O2, heat in microwave oven (Milestone MLS 1200 microwave oven), trans to digestion tube, heat on block

811

812

TABLE 25.7 Examples of conventional pressure wet digestion procedures utilizing closed systems and oxidizing acids (including hydrogen peroxide) for foods, feedstuffs and related biological materials Elements determined

Decomposition procedure

Determinative technique

Reference

Human colostrum

Fe, Cu, Zn, Cr

T-XRF

[84] (2002)

Total diet samples (Swiss), non-fat milk powder NIST-SRM 1549, whole milk powder NIST-RM 8435, whole egg powder NISTSRM 8415, total diet NIST-SRM 1548, peach leaves NIST-SRM 1547, pine needles NIST-SRM 1575, durum wheat ﬂour NISTRM 8436, hay powder BCR-CRM 129, single cell protein BCR-CRM 274 3 common ﬁsh species, ulva lactuca BCR-CRM 279

I

Polyethylene vial, HNO3/H2O2, heat at 608C in closed/sealed vials for 5 days, allow to dry, diss HNO3, (further treatment for T-XRF); [closed, low pressure] Dry sample into 15 ml quartz vessel, water, 129I, HNO3, seal w PTFE tape and close w lid, place in high pressure asher autoclave (high pressure asher autoclave, Ku¨rner), autoclave (21 vessels) close and ﬁll w N2 at 100 bar, external autoclave pressure 130 bar, heat according to program to 2308C, dil water; [closed, high pressure]

ICP-MS

[251] (2000)

F-AAS

[252] (1999)

T-XRF

[86] (1998)

39 different teas, GBW 09505 tea from Research Center for Eco-Environmental Sciences, Academia Sinica, Beijing)

Cu, Cr, Ni, Zn, Fe, Mn P, S, K, Ca, Ti, Mn, Fe, Ni, Cu, Zn, Se, Rb, Sr, Ba, Pb

Teﬂon vessel, HNO3, heat under pressure at 1208C; [closed, moderate pressure] Quartz tube, HNO3/HCl, seal tube, place in HPA (HPA, Hans Ku¨rner Analysentechnik), heat to 2608C (pressure increases to max 13 MPa), cool, add internal standards; [closed, high pressure]

M. Ihnat

Matrix

TABLE 25.7 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Plankton, periphyton (plants)

Ti, Mn, Fe, Ni, Cu, Zn, Se, Rb, Sr, Pb

T-XRF

[253] (1998)

Hay powder BCR-CRM 129, milk powder BCR-CRM 150, pig kidney BCR-CRM 186, cod muscle BCRCRM 422, spinach NIST-SRM 1570, citrus leaves NIST-SRM 1572, oyster tissue NIST-SRM 1566a

I

Periphyton disc into tube [50 ml Oak Ridge centrifuge tubes of ﬂuorinated ethylene –propylene copolymer (Tefzel FEP) with screw closures of ethylene-tetraﬂuoroethylene copolymer (Tefzel EFTE)], HNO3, tube with loosely ﬁtted closure heated in water bath at 1008C, H2O2, heat at 100C… other operations…transfer to microcentrifuge tube, freeze; [open/closed, normal pressure] PTFE liner of bomb [High pressure steel bomb, model DAE II, with PTFE liners 50 ml (Berghof)], HNO3/HClO4, close, heat at 2608C, dil water; [closed, high pressure]

ICP-MS

[254] (1997)

continued

Sample preparation for food analysis

Matrix

813

814

TABLE 25.7 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Human milk, bovine milk, non-fat milk powder NIST-SRM 1549

Li, Be, B, Al, Sc, Ti, V, Cr, Mn, Fe, Co, Ni, Cu, Zn, Ga, As, Se, Rb, Sr, Mo, Cd, Cs, Ba, Pb, Th, U

ICP-MS (Li, Be, B, Al, Sc, Ti, V, Cr, Mn, Fe, Co, Ni, Cu, Zn, Ga, As, Se, Rb, Sr, Mo, Cd, Cs, Ba, Pb, Th, U); ICP-OES (P, Mg, Ca, Na, K, Zn, Fe)

[255] (1997)

Tuna ﬁsh

As, Hg, Cu, Cd, Pb, Ni, Cr

ET-AAS (As, Cu, Cd, Pb, Ni, Cr); CV-AAS (Hg)

[256] (1997)

Roots and tubercles (radish, celery, potato), bulbs (onion, leek), leaves and soft stalks (chard, spinach, lettuce, endive), cabbages (cauliﬂower, cabbage), fruits and garden produce (tomato, green pepper, artichoke, green bean, eggplant), Bovine liver NIST SRM 1577

Cr, Cu, Zn

High pressure asher, HPA (Anton Paar) (1) 70 ml HPA quartz vessel, HNO3, evap to dryness on hot plate at 1008C, HNO3, digest in HPA according to T – P program (2) 70 ml HPA quartz vessel, HNO3, predigest on hot plate at 708C, HNO3, digest in HPA according to T – P program (3) 70 ml HPA quartz vessel, HNO3, digest in HPA according to T – P program; [closed, high pressure] Custom-made PTFE-lined digestion bomb w inner quartz tube and lid, HNO3, close with a special seal of PTFE, heat in oven at 1808C, cool, dil water; [closed, moderate pressure] Teﬂon bomb, HNO3, heat at 1108C, dil water; [closed, moderate pressure]

ET-AAS (Cr); F-AAS (Zn); ICP-OES (Cu)

[257] (1993)

M. Ihnat

Matrix

TABLE 25.7 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Human whole blood, urine, canned tuna, tuna NIST-RM 50, pond sediment NIESCRM-02, control blood OSSD20/21 (Behring), urine control (Kaulson)

Hg

CV-AAS

[258] (1993)

Spinach NIST-SRM 1570, mussel NIES-CRM 6

Mg, Al, Sc, Mn, Co, Ni, Cu, Zn, Cr, Fe, As, Rb, Sr, Ag, Cd, Sb, La, Eu, Hg, Tl, Pb, Th, U Al, Cu, Mn, Mo, Sn

PTFE vessel, water/HNO3, cap, place crucible in SS body of the Parr 4745 bomb [Parr Instrument Co. (Moline IL)], close SS screw cap, heat at 1308C in gravity convection oven, HClO4, heat again at 1308C, dil water; [closed, high pressure] Quartz device, HNO3, digest in digestion bomb (Berghof), dil water; [closed, moderate pressure]

T-XRF; ICP-MS

[259] (1992)

ET-AAS

[260] (1992)

Canadian infant formulas, evaporated milk

Graduated polystyrene centrifuge tube, water, citric acid/HNO3 (citric acid prevents pptn of stannic acid in absence of organic matter), cap tubes snugly, heat 2/3 immersed in water bath at 558C, release pressure, dil water, shake, stand; [closed, normal pressure]

continued

Sample preparation for food analysis

Matrix

815

816

TABLE 25.7 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Cereals (wheat, corn, barley), legumes and oilseeds (snap beans, dry beans, adzuki beans, tepary beans, chick peas, ﬁeld beans, lentils, soybeans, sesame) Mussel NIES-CRM-6, algae NIES-CRM-9

Cr

PTFE container, HNO3, heat at 1508C in an autoclave, water, dil water; [closed, moderate pressure]

ET-AAS

[261] (1990)

Zn, Cd, Pb, Cu

Quartz vessel, HNO3, heat at 3208C [high pressure asher (HPA), Anton Paar], cool, clear solns, evap to dryness at 1308C in small quartz vessel to remove nitrous oxide, HNO3/water, heat, dil water; [closed, high pressure] (Sediments) sample into LORRAN Teﬂon pressure decomposition vessel (H.K. Morrison and Sons Ltd, NS), HNO3/HF/HClO4, heated submerged in water bath at 1008C, trans to PFA beaker, evap on hot plate and IR lamp to incipient dryness, HF/HClO4, again to near dryness, HCl/HNO3, warm, dil water, complete solubilization; [closed, moderate pressure]

ASV

[93] (1989)

ET-AAS; HG-AAS; ID-ICP-MS

[85] (1987)

Lobster hepatopancreas NRCC-TORT-1, marine sediment sediment NRCC-BCSS-1, marine sediment NRCC-MESS-1

Sn

M. Ihnat

Matrix

TABLE 25.7 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Whole blood, urine, grass, leaves, earth worms, wine, beer, spirits, eggs, milk, cheese, butter, oil, liver, sewage sludge

Hg

CV-AAS

[262] (1984)

Fish, meat, blood, water

Be, Se, I, Hg

(1) Custom-made semi-closed quartz tubes of 10 and 40 ml w quartz bars (boiling aid) and head pieces w polished connections; (2) closed quartz vessels of 50 ml and 250 ml w ﬁnely polished connections and covers w hooks/springs (Normschliff Gera¨tebau) under slight overpressure (max 4 bar): sample into vessel, HNO3/HClO4 or only HNO3 for difﬁcult materials, predigest, heat to max 2008C or lower depending on system/material (care w alcoholic beverages; avoid HClO4 for fatty materials); [partly closed, about normal pressure; closed, moderate pressure] Teﬂon tube vessel (custom-made high pressure digestion device), HNO3 or HNO3/HF or HF/HClO4, heat at 1708C; [closed, high pressure]

Determinative technique ‘not reported’

[263] (1972)

Sample preparation for food analysis

Matrix

817

818

TABLE 25.8 Examples of microwave-assisted wet digestion procedures for foods, feedstuffs and related biological materials Elements determined

Decomposition procedure

Determinative technique

Reference

Evaporated milk, cheese, bacon, tuna, eggs, peanut butter, corn, bread, pancakes, breakfast cereal, prune juice, lemonade, broccoli, potato, spaghetti/meat balls, mayonnaise, beer, beef, haddock, canned pears, rice ﬂour NIST-SRM 1568a, total diet NIST-SRM 1548, whole egg powder NIST-RM 8415, dogﬁsh muscle NRCC-DORM-1, oyster tissue NIST-SRM 1566, mussel NIES-CRM-06, bovine liver NIST-SRM 1577a Rice ﬂour NIST-SRM 1568, apple leaves NIST-SRM 1515, wheat gluten NIST-RM 8418, spinach NIST-RM 1570a, lucerne 12-2-03 SMI Slovakia, lichen BCR-CRM 482

Al, As, B, Ba, Cd, Ca, Cr, Co, Cu, Fe, Pb, Mg, Mn, Mo, Ni, P, K, Se, Na, Sr, Tl, V, Zn

TFM Teﬂon-lined heavy duty vessel (limit 2008C, 600 psig), HNO3, seal, heat in microwave oven (MDS-2000 microwave system, CEM), cool, dil water (digestions are judged complete if temp reaches 2008C and clear to light yellow solns are produced; incompletely digested ones are discarded and dign repeated w smaller mass); [closed, high pressure]

ICP-OES

[95] (2002)

Zn

PTFE vessel, water, HNO3, heat microwave oven (microwave decomposition unit UniClever, Plazmatronika-Service, Poland), cool, dil water; [closed, high pressure]

ICP-MS

[264] (2001)

M. Ihnat

Matrix

TABLE 25.8 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Soil, ﬁsh ﬁllet ash, grass ash (weed, ornamental grasses)

U, Am, Cm

RNAA

[265] (2001)

Food spices (18 species including ginger, garlic, onion, turmeric, cinnamon, bay laurel, red pepper, cumin, caraway, mustard, etc.), pulses (8 species incl soybean, adzuki bean, red bean, kidney bean, pea, chick pea, mung bean, black gram, red gram), Bowen’s kale, tomato leaves NIST-SRM 1573, tea leaves NIES-CRM-07

Na, Mg, Al, Cl, K, Ca, Sc, V, Mn, Fe, Co, Zn, Br, Rb, Sb, Cs, Ba, La, Ce, Sm, Eu, Dy, Yb, Lu

Dry ashing/microwave digestion: dry ash at 5508C, reash if required at 6008C. Grass (two step): place ash into Teﬂon decomposition vessel (Floyd double wall 90 ml, limits 2508C, 200 psig), HNO3, heat in microwave oven (Floyd microwave RMS-850 system) acc to program, trans to PP tube, centrifuge, recentrifuge, trans solid residue back into dig vessel, HF, heat in micro-oven, ﬁlter, HNO3, evap to incipient dryness, pool w previous soln, use for RNAA. Fish (one step): ash into vessel, HNO3, heat in microwave oven, ﬁlter use ﬁltrate for RNAA; [closed, high pressure] After neutron irradiation: irradiated sample into Teﬂon vessel, HNO3 þ carriers, cap, heat in domestic microwave oven, cool, transfer beaker, HClO4, heat on hot plate to near dryness, H2O2, heat to near dryness, diss HCl…; [closed, moderate pressure, with subsequent hot plate dign]

INAA/RNAA (Na, Mg, Al, Cl, K, Ca, Sc, V, Mn, Fe, Co, Zn, Br, Rb, Sb, Cs, Ba, La, Ce, Sm, Eu, Dy, Yb, Lu); PAA (Na, Mg,Cl,Ca,Mn, Ni, Zn, Rb, Sr)

[266] (2000)

819

continued

Sample preparation for food analysis

Matrix

820

TABLE 25.8 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Bowen’s kale, hay powder IAEAV-10, spinach NIST-SRM 1570, orchard leaves 1571, apple leaves NIST-SRM 1515, spinach leaves NIST-SRM 1570a, tomato leaves NIST-SRM 1573a, oriental tobacco leaves ICHTJ CTA-OTL-1, Virginia tobacco leaves ICHTJ CTA-VTL-2

Co

INAA, RNAA

[220] (2000)

Apple leaves NIST-SRM 1515, peach leaves NIST-SRM 1547, tomato leaves NIST-SRM 1573, pine needles NIST-SRM 1575, wheat ﬂour NIST-SRM 1567a, rice ﬂour NIST-SRM 1568a

Al, As, B, Ba, Ca, Cd, Co, Cr, Cu, Fe, Hg, K, Mg, Mn, Mo, Na, Ni, P, Pb, S, Sb, Se, Si, Sr, Th, Ti, Tl, U, V, Zn

After irradiation, wet ash samples by 3 methods: MD1: one-step microwave digestion in closed system w HNO3 (Plazmatronica, Poland); MD2: one-step microwave digestion in closed system w HNO3/H2O2 (Plazmatronica, Poland); MD3: one-step microwave digestion in closed system w HNO3/H2O2/HF (Plazmatronica, Poland); [all methods closed, high pressure] (1) PFA inner liner, HNO3/HF, predigest at RT, heat microwave oven [QWAVE-1000 microwave sample preparation system, Questron, T limit 1758C, pressure limit 15.2 bar (220 psig)], cool, dil water (2) PFA inner liner, HNO3/HF, predigest at RT, heat microwave oven (QWAVE-1000), H3BO3, warm to 1008C in closed vessel, cool, dil water; [closed, moderate pressure]

ICP-MS

[267] (1999)

M. Ihnat

Matrix

TABLE 25.8 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Human milk and colustrum, milkbased infant formula, soy-based infant formula, non-fat milk powder NIST-SRM 1549

Se

HG-AAS

[268] (1999)

Rice, rice ﬂour NIES-CRM-10

P, K, Mg, Ca, Mn, Zn, Fe, Cu, Al, Na, As, Mo, Cd

Medium pressure Teﬂon vessel (Milestone MV), HNO3/H2O2, heat in microwave oven (Milestone MLS 1200 microwave oven), trans to digestion tube, heat on block digestion system (Kjeldatherm Gerhardt block digestor) to evap to 1 ml and obtain clear digest, reduce Se w HCl, dil water; [closed, moderate pressure] Advanced composite Teﬂon PFA digestion vessel (CEM), HNO3/H2O2, heat in microwave oven (CEM MDS-2000), cool, dil water; [closed, moderate pressure]

[269] (1999)

Mussel, lobster hepatopancreas NRCC TORT-1

Cd

ICP-OES (P, K, Mg, Ca, Fe, Mn, Zn, Cu, Al); ICP-MS (Ni, Mo, As, Cd); F-AAS (Na) F-AAS

Pulverized sample into Teﬂon bomb (Parr 4782 microwave digestion bomb designed speciﬁcally for microwave heating from high strength microwave-transparent material to enclose a chemically inert Teﬂon sample cup of 45 ml), HNO3, heat in domestic microwave oven, cool, adjust pH, dil water; [closed, moderate/high]

[270] (1998)

821

continued

Sample preparation for food analysis

Matrix

822

TABLE 25.8 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Peas, wheat ﬂour NIST-SRM 1567a

Ag, Al, Au, B, Ba, Be, Bi, Ca, Cd, Ce, Co, Cr, Cu, Dy, Er, Eu, Fe, Ga, Gd, Ge, Hf, Ho, Ir, La, Lu, Mn, Mo, Nb, Nd, P, Pb, Pd, Pr, Pt, Re, Rh, Sb, Sc, Se, Si, Sm, St, Ta, Tb, Te, Th, Ti, Tl, Tm, U, V, Y, Yb, Zn, Zr Cr, Fe, Cd, Pb

PTFE PFA vessel (CEM), HNO3, heat in microwave oven (CEM MDS 2000), dil clear residue-free digest dil w water; [closed, moderate pressure]

HR-ICP-MS

[271] (1998)

Teﬂon vessel (Milestone), HNO3/spike, heat in microwave oven (Milestone microwave digestion system MLS-1200 MEGA), cool, HF/HClO4, heat at 1008C to incipient dryness, HNO3, evap to near dryness, dil water; [closed, moderate or high pressure]

ID-ICP-MS

[272] (1997)

Unpolished rice

M. Ihnat

Matrix

TABLE 25.8 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

CRMs: brown bread BCR-CRM 061, whole meal ﬂour BCR-CRM 189, rice ﬂour NIES-CRM-10a/b/c, corn bran NIST-RM 8433, durum wheat ﬂour NIST-RM 8436, hard red spring wheat ﬂour NIST-RM 8437, soft winter wheat ﬂour NIST-RM 8438, rice ﬂour NIST-SRM 1568a, wheat ﬂour NIST-SRM 1567a, wheat ﬂour NRCCRM, rice ﬂour NRCCRM (Beijing, PRC), whole wheat CGC-QC-4, CGC-QC5 (Canadian Grain Commission, Winnipeg)

Cd, Cu, Pb, Se

ET-AAS

[273] (1997)

Fresh eggs, pork liver GBW 08551

Cu, Zn, Ca, Mg, Fe

(1) Closed vessel: advanced composite vessel (CEM), HNO3, seal, heat microwave oven (microwave digestion system CEM MDS-2000), cool, dil water, sample solns are clear/light yellow (2) Open vessel: vessel (focused open-vessel digestion system, Prolabo Microdigest A301 w PTFE pumps, borosilicate vessels and operating software), cap w reﬂux column, load into A301 hood, HNO3, heat/digest, cool, dil water, solns are light yellow/ clear; [closed, moderate/high pressure; open, normal pressure] (Eggs) sample into glass beaker, HNO3/H2O2, heat at 180 –1908C on hot plate, dry, diss water, trans to PTFE calibrated ﬂask for F-AAS of Cu, Fe, Ca, Mg. Another sample into PTFE bomb, HNO3/H2O2, heat in microwave oven, cool, dil water for Zn; [open, normal pressure; closed, moderate pressure]

F-AAS

[252] (1997)

continued

Sample preparation for food analysis

Matrix

823

824

TABLE 25.8 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Whole milk, skim milk, UHT milk, non-fat milk powder NIST-SRM 1549

Mn, Zn

ET-AAS

[274] (1995)

Noodles

B

(1) Dry ashing: porcelain crucible, dry on hot plate, char, ash at 4508C, HNO3 if required, heat in furnace, diss HNO3, dil HNO3 (2) Microwave digestion: vessel (custom-made PTFE reactors to withstand high P/T), HNO3, seal, heat in domestic microwave oven [MDS-2000 Microwave oven (CEM) used for monitoring pressure and temperature during mineralization], cool, dil yellowish and transparent soln w water; [closed, high pressure] PTFE TFM digestion vessel (Milestone), HNO3/H2O2, heat in microwave oven (Model MLS 1200 MEGA, Milestone, Italy), cool, dil water, ﬁlter store in plastic tube; [closed, moderate or high pressure]

ICP-OES

[275] (1995)

M. Ihnat

Matrix

TABLE 25.8 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

18 RMs, tobacco, peat, pine bark, ryegrass, apple, rice, vine leaves, cotton, hay, lettuce (CII), whole meal ﬂour BCR-CRM 189, skim milk powder-spiked BCR-CRM 151, platihypnidium ripariodes BCRCRM 61, lagarosiphon major BCR-CRM 60, mussel NIES-CRM06, pepperbush NIES-CRM-01, total diet NIST-SRM 1548, tomato leaves NIST-SRM 1573a.

P, B, Cu, Fe, Mn, Zn, Cd, Pb

(1) Botanical sample into vessel (microwave acid digestion bomb for pressure up to 8 £ 106 Pa, Parr, Moline, IL), HNO3, heat in Moulinex domestic microwave oven (Moulinex microwave oven model 430), cool, dil water, settle/centrifuge/ ﬁlter; [closed, high pressure] (2) Botanical sample into vessel (microwave acid digestion bomb for pressure up to 8 £ 106 Pa, Parr, Moline, IL), HNO3, heat in Moulinex domestic microwave oven, cool, HF, heat on hot plate, diss NA, dil water, settle/centrifuge/ﬁlter; [closed, high pressure] (3) Milk sample into vessel (Parr or Milestone), HNO3/H2O2, heat in Moulinex domestic microwave oven or Milestone microwave oven (Milestone microwave system MLS-1200 MEGA, Milestone, Sorisole, Italy), cool, dil water, settle/centrifuge/ﬁlter; [closed, high pressure]

F-AAS (Fe, Cu, Mn, Zn); ET-AAS (Cu, Mn, Cd, Pb)

[276] (1993)

Sample preparation for food analysis

Matrix

825

continued

826

TABLE 25.8 (continuation) Matrix

Elements determined

Na, K, Mg, Fe, Ca, Zn, Mn

Lobster hepatopancreas NRCC-TORT-1

As, Cd, Co, Cr, Cu, Fe, Mn, Mo, Ni, Pb, Se, Sr, V, Zn

(4) Milk sample into vessel [Teﬂon vessel (MRD 1000/6/100/110 designed for pressures up to 11 £ 106 Pa)], HNO3/HClO4, heat Milestone microwave oven, cool, dil water, settle/centrifuge/ﬁlter; [closed, high pressure] Digestion tube, place on carousel of microwave digestion apparatus (Microdigest A-300, Rhone Poulenc, Prolabo Div.), run program suited to matrix w HNO3/H2O2; [open, normal pressure] (1) Sample into vessel (PFA-Teﬂon pressure-relief digestion vessel 120Ml (CEM) [All PTFE materials are made from tetraﬂuoroethylene with a fully ﬂuorinated alkoxy side chain (i.e., PFA-Teﬂon)] the pressure relief valve opens at 6.8 atm), HNO3 and 30 or 50% H2O2, cap tightly, heat in microwave oven (CEM model MDS-81), cool, dil water, store in PP bottles; [closed, moderate pressure]

Determinative technique

Reference

F-AAS

[277] (1991)

ET-AAS; FAAS

[88] (1989)

M. Ihnat

Breakfast cereal, baked goods, feeds, AAFCO feed No. 1, blind feed No. 2, pasta, bovine liver NIST-SRM 1577a, citrus leaves NIST-SRM 1572

Decomposition procedure

TABLE 25.8 (continuation) Matrix

As, Cd, Co, Cr, Cu, Fe, Mn, Ni, Pb, Se, Zn

Decomposition procedure (2) Sample into vessel [Parr 4781 microwave digestion bomb w a sealable 23 ml PTFE cup w cover contained in a polymer resin shell (pressure to 82 atm)], HNO3/H2O2, close tightly, heat in microwave oven (CEM model MDS-81), cool, dil water, store in PP bottles; [closed, high pressure] (3) Sample into vessel (Berghof PTFE digestion vessel no. 14029 100 ml, max pres 24.7 atm), HNO3 and 30 or 50% H2O2, close tightly, heat in microwave oven (CEM model MDS-81), cool, dil water, store in PP bottles; [closed, high pressure] Teﬂon PFA CCT or PRT vessel, HNO3/HClO4, tightly seal, heat in microwave oven to 70–75 psig [Model MDS-81 microwave oven, CEM w PFA 120 ml vessels (completely closed type CCT and pressure-relief type PRT)], cool, dil water, store in PP bottles; [closed, moderate pressure]

Determinative technique

Reference

ET-AAS

[278] (1988)

827

continued

Sample preparation for food analysis

Lobster hepatopancreas NRCC-TORT-1, marine sediment NRCC-MESS-1

Elements determined

828

TABLE 25.8 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Oyster tissue NIST-SRM 1566, rice ﬂour NIST-SRM 1568, wheat ﬂour NIST-SRM 1567, bovine liver NISTSRM 1577a, human urine NISTSRM 2670

None reported

Determinative technique ‘not reported’

[279] (1986)

Bovine liver NIST-SRM 1577, orchard leaves NIST-SRM 1571

Zn, Cu, Pb, As, Se, Co, Cr, Ni

60 ml Teﬂon PFA vessel (Savillex Corp.), HNO3, predigest on hot plate until nearly dry, HNO3, seal, heat microwave oven (microwave digestion system MDA-81, CEM Corp.); [closed, moderate pressure] Dried plant or animal sample into 125 ml Erlenmeyer ﬂask, HNO3/HClO4, heat in microwave oven (modiﬁed domestic microwave oven) until fumes of HCLO4; for F-AAS; also postirradiation digestion for RNAA; [open, normal pressure]

F-AAS (Zn, Cu, Pb); RNAA (As, Se, Co, Cr, Ni)

[87] (1975) M. Ihnat

Matrix

TABLE 25.9 Examples of slurry preparation and online digestion procedures for foods, feedstuffs and related biological materials Elements determined

Decomposition procedure

Determinative technique

Reference

Sewage sludge, wheat ﬂour NIST-SRM 1567, spinach NIST-SRM 1570, orchard leaves NIST-SRM 1571, tomato leaves NIST-SRM 1573, bovine liver NISTSRM 1577, coal ﬂy ash NIST-SRM 1633a, sewage sludge BCR-CRM 145R, pig kidney BCR-CRM 186, sewage sludge CRM PR9472 Spinach NIST-SRM 1570, orchard leaves NIST-SRM 1571, citrus leaves NIST-SRM 1572, tomato leaves NISTSRM 1573, bovine liver NIST-SRM 1577, pig kidney BCR-CRM 186 Wood from balsam ﬁr, black spruce, jack pine

Cd, Cu, Mn, Pb, Zn

Add sample to HNO3 w stirring

F-AAS

[280] (2002)

Cd, Cu, Pb

Beaker, HNO3/TX-100, ultrasonicate (ultrasonic bath, Branson)

TS-FF-AAS

[281] (2002)

Cu, Fe, Mn

(1) Sample (milled w Tecator Cyclotec sample mill), wet/swell w water/ ZnCl2, dil ethanol/water, process w modiﬁed homogeniser with pressure to 80 psig (Emulsifex model C5 homogeniser, Avestin Inc.); (2) sample, suspend in ethanol/water/TMAH, homogenise as in (1); (3) sample, suspend/swell w DMSO or N,Ndimethylacetamide/LiCl or MMNO (N,N-dimethylacetamide,4-methylmorpholine-N-oxide)

ET-AAS

[282] (2001)

829

continued

Sample preparation for food analysis

Matrix

830

TABLE 25.9 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Various colour additives, ﬂy ash NISTSRM 1633a

As, Cd, Pb

ET-AAS

[283] (2001)

Wheat ﬂour samples from pilot plant, whole meal ﬂour BSR-CRM 189

Ni, Cr, Co

ET-AAS

[284] (1999)

Fish, copepod homogenate IAEAMA-M-1, ﬁsh ﬂesh homogenate IAEA-MA-A-2 Cabbage GBW 08504, oyster tissue NIST-SRM 1566a, orchard leaves NIST-SRM 1571 Human hair, human hair BCR-CRM 397

Cd

ET-AAS

[285] (1999)

ET-AAS

[286] (1998)

F-AAS; F-AES

[287] (1998)

Fish, oyster tissue NIST-SRM 1566a, ﬁsh homogenate IAEA MA-A-2 1062/TM, mussel NIES-CRM-6

Fe

Plastic vessel, TX-100/HNO3/H2O2/ matrix modiﬁer (Ni or NH4H2PO4), silicone antifoam, sonicate (ultrasonic bath, Branson), take aliquots from stirred suspension for EAAS PTFE tube, HNO3/H2O2, homogenize by ultrasonic agitation (ultrasonic bath, Bandelin), before analysis Calibrated glass ﬂask, dil to vol w TX-100/HNO3, homo in ultrasonic bath (Thornton) PFA container w screw cap, HNO3 or glycerol, homogenize by stirring w magnetic stirrer Sample (Retsch vibrating ball-mill w zirconia cups and balls), polyethylene vial, water, 48C, slurry into volumetric ﬂask w HNO3, dil as required, stir ultrasonically before aspiration HNO3/H2O2, ultrasonic bath (Branson), dil water

[288] (1998)

Fish, ﬁsh homogenate IAEA MA-A-2 1062/TM

Se

LAS (SS/ online micro wave diges tion/LAS) ET-AAS

Cd, Cu, Mn, Pb Ca, Cu, K, Mg, Na, Zn

HNO3/TX-100, ultrasonic agitation (ultrasonic bath, Thornton)

[289] (1997)

M. Ihnat

Matrix

TABLE 25.9 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Animal diet mixture (w variety of plant and animal materials), dogﬁsh muscle NRCC-DORM-1, dogﬁsh muscle NRCC-DORM-2, bovine muscle powder NIST-RM 8414, oyster tissue NISTSRM 1566, lobster hepatopancreas NRCC-TORT-1, B-4 liver, B-4 kidney, B-10 liver, B-10 kidney, pine needles NIST-SRM 1575, apple leaves NISTSRM 1515, corn stalk NIST-RM 8412, corn bran NIST-RM 8433 Rice, spinach, tomato leaves, soil, rice ﬂour NIST-SRM 1568, estuarine sediment NIST-SRM 1646, coal ﬂy ash NIST-SRM 1633, Buffalo River sediment NIST-SRM 2704, spinach NISTSRM 1570, coal NIST-SRM 1632a, tomato leaves NIST-SRM 1573 Cauliﬂower stem and leaves, bean leaves, grapefruit leaves, citrus leaves NIST-SRM 1572, apple leaves NISTSRM 1515

Cr, Cu, Fe, Mn, Ni, Se

Add sample to ethanol/water/TMAH in beaker, macerate/blend in a Tissuemizer (SDT Tissuemizer, Tekmar), process through homogenizer at 138 MPa (Emulsiﬂex model EF-B3, Avestin)

ET-AAS

[290] (1996)

Pb, Cr, Mn, Zn, Cu

Teﬂon autosampler cup, water/TX100 or HNO3/TX-100 (no details) (larger slurry vols prepared in PE tt or bottles and vortex mixing)

ET-AAS

[291] (1994)

Ca, Mg, Fe, Mn

Suspend sample ash w glycerol/HCl, ultrasonic agitation, stir

FI-F-AAS

[292] (1993)

continued

Sample preparation for food analysis

Matrix

831

832

TABLE 25.9 (continuation) Elements determined

Decomposition procedure

Determinative technique

Reference

Tomato leaves NIST-SRM 1573, pine needles NIST-SRM 1575, citrus leaves NIST SRM 1572, aquatic plant BCRCRM 60, aquatic moss BCR-CRM 61

Pb

ET-AAS

[80] (1990)

Spinach NIST-SRM 1570, wheat ﬂour NIST-SRM 1567, bovine liver NISTSRM 1577, tomato leaves NIST-SRM 1573

Mn, Fe, Cr, Cu, Al

ET-AAS

[186] (1990)

Mussel, lettuce, tomato, whole diet IAEA-H-9

Pb

Sample (grind w vibrational-type microdismembrator, B. Braun, w Teﬂon “egg” cylinder and ball or in agate ball-mill), into 20 ml sterile container, water, shake mechanically, transfer to vol ﬂask w water, Viscalex HV30/antifoam/NH3, shake, dil water PP tube or PE bottle, add zirconium oxide beads, water or HNO3, wet grind by agitating (wet grinder w zirconium oxide beads, Glen mills), coarse ﬁlter to remove beads, vortex mix before withdrawing aliquot for EAAS (ultrasonic mixer, Kontes) PE bottle w zirconia spheres (Glen Creston), TX-100, shake on a ﬂask shaker

HG-AAS

[293] (1989)

M. Ihnat

Matrix

Sample preparation for food analysis

digestion system (Kjeldatherm Gerhardt block digestor) to evap to 1 ml and obtain clear digest, reduce Se w HCl, dil water, this means: place a weighed sample into a medium pressure Teﬂon vessel (Milestone MV), add HNO3/H2O2 (speciﬁed volume/concentration), heat in a microwave oven (Milestone MLS 1200 microwave oven) under speciﬁed conditions, transfer to a digestion tube, heat on a block digestion system (Kjeldatherm Gerhardt block digestor) to evaporate to 1 ml and obtain clear digest, reduce Se with HCl (speciﬁed volume/concentration), dilute with water. Analytical techniques applied to quantitation after treatment are also listed; those without qualiﬁcation imply that generally all techniques were used for all the elements listed in column 2, while speciﬁc elements are indicated in those cases for element-speciﬁc measurements. Abbreviations for instrumental (or other) analytical techniques used for quantiﬁcation are to be found in the master list at the end of this book. 25.5.1 No treatment Table 25.2 presents examples of sample treatment procedures involving no treatment or minimal treatment, including dissolution and extraction, but not slurry sample preparation, which is a preparative technique of current interest and worthy of stand-alone consideration. All these procedures fall under the classiﬁcations (listed in Table 25.1): (1) no treatment or minimal treatment and (2) extraction. This ﬁrst tabulation concerns instances where no treatment or only minimal treatment is required in preparing the sample for the quantitation step and the material is used more or less directly as received. Such approaches are used sporadically and only when the starting material is a homogeneous liquid or solid and the instrumental/determinative technique can accept it directly. Placement of the sample into ampoules, vials, bags or other containers for techniques such as neutron irradiation for INAA, and pressing into pellets for XRF and near infra-red spectroscopy (NIR) are examples of “no treatment”. On a microscale, removal of milligram or submilligram quantities for introduction into a graphite furnace for SS-ET-AAS is an example of an interesting direct solid sampling technique. Procedures involving minimal treatment include steps such as dilution of liquids with water or other reagents, dissolution/solubilization (of fats, oils) in organic solvents and preparation of emulsions or suspensions by shaking or ultrasonic dispersion. A step up from these procedures are those approaches involving extraction and leaching with agents such as water, chelating agents such as EDTA and acids such as HCl or HNO3 at ambient or higher temperatures, generating ﬁltrates or supernatant solutions. Some of these extraction procedures may be classiﬁed as partial digestion and/or selective digestion and may provide a means for analyte speciation or phase analysis.

833

M. Ihnat

Deproteination, precipitation of protein and haemolysis also fall into this category. 25.5.2 Dry ashing Examples of sample treatment procedures involving three major classes of dry ashing/dissolution approaches are presented in Tables 25.3–25.5. Procedures are divided into those employing dry ashing/dissolution without ashing aids (Table 25.3), those incorporating ashing aids (Table 25.4) and those utilizing closed and open oxygen combustion and low temperature ashing (Table 25.5). The dry ashing procedure usually denoted just that, but the more complete nomenclature is dry ashing/dissolution, as most often the dry ashing stage is followed by dissolution of the ash in aqueous acidic solution. This technique has been covered in detail by Hoenig in Chapter 7. Dry ashing/dissolution procedures listed in Tables 25.3 and 25.4 fall under the classiﬁcation: 3.1.1.1 Dry ashing/(dissolution), high temperature, combustion with air/oxygen, stationary system, and include (A) open mufﬂe furnace, in air under atmospheric pressure, without ashing aid and (B) open mufﬂe furnace, in air under atmospheric pressure, with ashing aid. Discussion is not included for the less common case, (C) combination dry ashing and wet ashing (digestion) procedures, open mufﬂe furnace in air under atmospheric pressure, with or without ashing aid, followed by more extensive acid treatment. The principal difference between the combination dry ashing and wet ashing (digestion) procedures and the typical dry ashing/dissolution procedures is the much more extensive (wet) manipulations conducted in the former. Rather than simple dissolution in acid by heating, these more involved operations include acid heating/digestions before or after dry ashing and repeated acid addition/boil-down/evaporation prior to a ﬁnal take-up in acid. In addition, no consideration is included in this chapter of methods (D) closed (low or high pressure), without or with ashing aid (mufﬂe furnace, hot plate, ﬂame). The common dry ashing procedure is referred to by various terms, such as dry ashing, classical dry ashing, oxidation, combustion, mineralization, destruction high temperature ashing, calcination and acid digestion of the ash. These are typically conducted with an open vessel (crucible, beaker) in a temperature programmable or temperature controlled mufﬂe furnace in air under atmospheric pressure, with or without ashing aid/additive; occasionally a hot plate or ﬂame is the source of heat. Dry ashing/dissolution procedures summarized in Table 25.5 utilizing closed oxygen combustion fall under the classiﬁcation scheme: 3.1.1.1 Dry ashing/(dissolution), high temperature, combustion with air/oxygen, stationary system, (E) oxygen ﬂask, normal (atmospheric) pressure, (F) oxygen bomb, high pressure. Those utilizing open oxygen combustion fall under: 3.1.1.2 Dry ashing/(dissolution), high temperature, combustion with air/oxygen, streaming

834

Sample preparation for food analysis

system, (A) oxygen/air stream or other oxidizing gases in combustion tube and (B) H2/O2-ﬂame in a closed, cooled system. Procedures listed in Table 25.5 utilizing low temperature ashing fall under the classiﬁcation: 3.2.1 Dry ashing/(dissolution), low temperatures, oxygen gas plasma at 100 –1258C, (A) radiofrequency electrical ﬁelds or (B) microwave electrical ﬁelds. Indications of classiﬁcation are included under the decomposition procedure column as: [closed, oxygen ﬂask at atmospheric pressure]; [closed, oxygen bomb at elevated pressure]; [open, combustion with streaming air/oxygen]; [closed, low temperature low pressure combustion with streaming plasma of oxygen gas]. Oxygen ﬂask combustion at atmospheric pressure is based on developments by Hempel followed by Scho¨niger. In oxygen ﬂask combustion, the sample, wrapped in paper or as a pellet, is afﬁxed to a holder. It is ignited (prior to introduction or after introduction by remote ignition) and inserted into a glass or plastic ﬂask ﬁlled with oxygen and containing a suitable absorbing solution. Combustion occurs in the immediately closed ﬂask at normal, atmospheric or only slightly raised pressure, and gaseous combustion products are absorbed in the solution, which is then used for analysis or processed further, as the need may be. Combustion in closed oxygen vessels under higher pressures is commonly denoted an oxygen bomb. High pressure combustion typically utilizes a sealed metal bomb ﬁlled with oxygen under pressure and ignited remotely. Examples include a Parr bomb with oxygen at 15 atm pressure; Parr bomb with Na2O2/KNO3/sugar (peroxide bomb technique); other devices using Na2O2; and a Berthelot vessel with external ignition and oxygen at 2.5– 4 MPa. Combustion in a ﬂowing stream of oxygen/air or other oxidizing gases, including Cl2, takes place in a combustion tube with the gaseous oxidants passing over the sample in the externally heated tube. Sample oxidation/pyrolysis of the sample produces volatile products and species of the sought-for element, which are collected by absorption, dissolution, amalgamation (Hg), or directed into a measuring device for determination. Reagents can be added to separate the element by distillation. The Dumas method, adapted by Pregl, and also known as the Pregl – Dumas method, is an excellent example of a combustion tube procedure used successfully in microanalysis for C, H, N and S determination. This principle forms the basis of modern automated combustion/elemental determination instruments. The To¨lg/Knapp combustion tube modiﬁcation (Trace-O-Mat, Anton Parr and Hans Ku¨rner) employs a combustion chamber and a condenser in which a sample pellet is burned in a stream of oxygen at atmospheric pressure. Oxygen –hydrogen or oxy/hydrogen ﬂame combustion in a closed and cooled system is often accomplished in specialized apparatuses (e.g. by Kunkel, Heraeus Quarzschmelze GmbH, Hanau, Germany, with version 4 said to be especially suitable for solid biological materials, and the Wickbold combustion apparatus).

835

M. Ihnat

Low temperature dry ashing is done with a radiofrequency plasma (excited oxygen plasma produced by electric glow discharge or radiofrequency electric ﬁelds) with the ash typically dissolved in HCl. An early low temperature asher (LTA) instrument was placed on the market by Tracer lab (now International Plasma Corp., USA; LFE Corp., Process Control Div. and Technics GmbH, Munchen). It uses oxygen at 2 torr and 27.12 MHz power. Low temperature oxygen –ﬂuorine ashing using atomic F produced by attack of atomic oxygen on PTFE crucibles holding the sample has been reported. Other, more specialized, dry ashing techniques such as electrothermal vaporization (as a means of sample introduction), pyrolytic decomposition under hydrogen or inert gases or laser ablation are not widely used in food analysis and are thus not included here for discussion. 25.5.3 Wet digestion—conventional Examples of sample treatment procedures involving two major, but different, conventional wet digestion approaches are presented in Tables 25.6 and 25.7. Procedures are divided into those utilizing open systems under atmospheric pressure (Table 25.6) and closed systems (Table 25.7), the common element in both cases being the use of oxidizing acids (and occasionally H2O2). The lengths of the tables reﬂect the fact that wet digestion is the most widely used and most popular sample decomposition procedure for biological materials. Classiﬁcations (according to Table 25.1) of the wet digestion procedures include wet ashing (digestion) conventional, high temperatures, normal pressure, oxidizing mineral acids, hydrogen peroxide (Table 25.6), and wet ashing (digestion) conventional, high temperatures, low, moderate or high pressure, oxidizing mineral acids, hydrogen peroxide (Table 25.7). The term conventional is included to indicate use of ordinary convective heating, in order to differentiate such procedures from the more recent applications of microwave heating for wet decomposition. Conventional acid digestion, also known as wet ashing, decomposition, destruction, mineralization or oxidative acid digestion, occurs in open vessels at atmospheric pressure using a variety of single or mixtures of oxidizing acids; it is the single most commonly used decomposition procedure for biological materials. The most commonly used acids and combinations include: nitric acid, sulphuric acid, perchloric acid, nitric acid/sulphuric acid, nitric acid/perchloric acid, nitric acid/perchloric/sulphuric, followed by other acid and reagent mixtures. Hydrogen peroxide, usually 30% but at times 50%, is a valuable additional reagent. Some automated versions of techniques for atmospheric pressure wet digestion are available (e.g. VAO automatic wet digestion device, Hans Ku¨rner Analysentechnik, Germany). Kjeldahl’s method (Kjeldahl digestion, the Kjeldahl –Wilforth – Gunning method for organic nitrogen), usually associated with the determination of nitrogen, may be mentioned

836

Sample preparation for food analysis

separately as an important case. It employs sulfuric acid with K2SO4, to raise the digestion temperature, and various catalysts and is usually conducted in open Kjeldahl ﬂasks or tubes. Many modiﬁcations are in use and closed tube variants are known. A fairly common example of a conventional, open vessel wet digestion procedure listed in Table 25.6 is as follows [81]: nitric acid is added to a weighed sample in a Kjeldahl ﬂask (this is a non-Kjeldahl method even though the procedure incorporates the most utilitarian of the digestion vessels, a Kjeldahl ﬂask) and following overnight predigestion, the mixture is heated on an electric Kjeldahl unit. Perchloric and sulfuric acids are added, the mixture is heated and nitric acid is added as needed to minimize charring, and the mixture is ﬁnally taken to white fumes of SO3. In another example [82], nitric acid is added to the sample in an Erlenmeyer ﬂask and the mixture is predigested on a hot plate. Nitric and perchloric acids are added and the mixture is heated to reﬂux, followed by repetition of the binary acid additions and heating to complete digestion to yield a nearly dry translucent material which is dissolved in nitric acid. In a third example [83], predigestion and digestion with nitric acid is accomplished with a Tecator digestion system. Table 25.7 provides examples of conventional wet digestion procedures utilizing closed systems under pressure, with further differentiation of classiﬁcation subdivision under low, moderate or high pressure, as indicated. Pressure is provided by placing the sample and acids in a variety of closed or sealed vessels, such as plastic bottles, disposable and sealable rigid polystyrene containers, specially constructed moderate or high pressure PTFE vessels, Teﬂon closed-tubes, glassy carbon-lined steel bombs or sealed quartz tubes (sealed quartz tube wet ashing procedure of Carius). Heat is provided by a conventional oven, pressure cooker, hot water bath, or an autoclave (unusual approach). An example from those of Table 25.7 for low pressure digestion is as follows [84]: place the sample with nitric acid and hydrogen peroxide into a polyethylene vial, close and heat in a water bath at 608C for ﬁve days. Open, allow to dry, dissolve the residue in nitric acid and take through further processing for T-XRF measurements. An example of a moderate pressure digestion is [85]: place the sample with a mixture of nitric acid, hydroﬂuoric acid and perchloric acid into a LORRAN Teﬂon pressure decomposition vessel and heat submerged in a water bath at 1008C. Transfer the contents to a PFA (Teﬂon) beaker and evaporate on a hot plate and under an infra-red lamp to incipient dryness. Add hydroﬂuoric/perchloric acids, evaporate again to dryness and dissolve in HCl/HNO3. An example of an operation under high pressure is [86]: seal the sample, nitric acid and HCl in a quartz tube, place in a commercial high pressure asher (HPA, Hans Ku¨rner Analysentechnik), heat to 2608C (pressure increases to 13 Mpa) and cool.

837

M. Ihnat

25.5.4 Wet digestion—microwave-assisted Table 25.8 presents examples of microwave-assisted wet digestion procedures. The procedures therein are classiﬁed as: 5. Wet ashing (digestion), high temperatures, microwave-assisted with further subdivisions of 5.1. open vessels at normal (atmospheric pressure); 5.2. closed vessels at moderate pressure; 5.3. closed vessels at high pressure. They all have in common the use of the increasingly popular microwave source for providing heat. Microwave digestion techniques have progressed from the early application studies of AbuSamra et al. at the University of Missouri [87] using a modiﬁed domestic microwave oven and open Erlenmeyer ﬂasks. Very few publications report use of open vessels with virtually all studies and applications using closed vessels at high temperatures and usually at moderate but sometimes at high pressure, and usually in commercially available laboratory-dedicated microwave ovens. The principles of this technique have been detailed by King in Chapter 2.3. An example of a closed vessel, moderate pressure, microwave-assisted digestion is taken from the work of Matusiewicz et al.[88]: a sample of Lobster Hepatopancreas (NRCC-TORT-1 CRM) is placed into a PFA-Teﬂon pressurerelief digestion vessel (CEM Corp., maximum pressure 6.8 atm). Nitric acid and 50% H2O2 are added and the tightly capped vessel is heated in a microwave oven (CEM model MDS-81). After cooling, the vessel is opened, the solution diluted with water and stored in polypropylene bottles for analysis by F-AAS and ET-AAS. An example of a closed vessel, high pressure, microwave-assisted digestion is from the same publication [89]: a sample of TORT-1 is placed into a sealable PTFE cup contained in a microwave-transparent polymer resin shell (Parr 4781 microwave digestion bomb, maximum pressure 82 atm). Nitric acid and 50% H2O2 are added and the tightly capped vessel is heated in a microwave oven (CEM model MDS-81). After cooling, the vessel is opened, the solution diluted with water and stored in polypropylene bottles for analysis by F-AAS and ET-AAS. Microwave digestion is currently receiving much research and developmental attention with recent trends toward automation. Automated microwave digestion techniques utilize ﬂow injection systems for online mineralization with the slurried sample ﬂowing through a coil inserted in a microwave oven as an open system. Both stopped-ﬂow (discrete ﬂow) and continuous-ﬂow systems are being studied with the potential for online coupling to atomic absorption and inductively coupled plasma atomic emission spectrometers. 25.5.5 Slurry sample preparation As mentioned at the beginning of this chapter, the goal of sample treatment is the quantitative release, from the sample, of the element of interest, usually associated with the concept of total quantitative decomposition, providing the

838

Sample preparation for food analysis

element in an aqueous inorganic solution. Extraction techniques may not provide quantitative extractions, but such procedures can be considered as alternatives to total, complete decompositions and are of interest for quick preparation of samples in a form amenable to acceptable introduction into the measuring device (slurry sampling) and automation. Preparation of sample slurries is one such partial decomposition/extractive approach. Table 25.9 provides examples of slurry preparation and online digestion procedures (for which slurries are eminently suited). For preparation of slurry, one begins with a sample in ﬁnely ground form. This material is then suspended in an aqueous medium and subjected to high speed or ultrasonic blending, providing a suspension of the powdered material in aqueous or liquid medium. If acid is a component of the suspending medium, then some extraction of the analyte or predigestion may occur. Typically, the slurry must be remixed by magnetic agitation, vortex mixing, gas mixing or ultrasonic agitation immediately prior to sampling for analysis. Alternatively, the slurry may be stabilized by using a sufﬁciently ﬁne powder or by means of emulsiﬁers, stabilizing or thixotropic agents. 25.6

SPECIFIC CASES: METHODS, ELEMENTS, MATRICES

The choice of sample preparation procedure has an impact on the performance of the quantitation technique used, the behaviour of the sought-after element and the amenability of the sample matrix to proper decomposition. Choice of procedure depends on (1) the nature of the organic/inorganic material, (2) the element to be determined and (3) the method used for its quantitation. Brief discussions relating to selected, speciﬁc relationships of decomposition procedures and quantitation techniques, elements and matrices are presented. 25.6.1 Analytical method The different mineralization and sample solubilization procedures generally are of great beneﬁt in, and go a long way towards, presenting the matrix in a form inﬁnitely more suitable for the measuring device. The physical and chemical complexities, however, of the starting matrices and of the chemical decomposition process lead to various stages of complete/incomplete decomposition and yield ﬁnal solutions of varying inorganic/organic composition. These solutions may contain remnants of undigested starting material, undissolved residues, undigested refractory organic compounds, decomposition-converted organic compounds, remains or products of decomposition acids/reagents, additional constituents leached out from decomposition vessels and equipment and excessive quantities of inorganic ions and compounds. These may or may not impact on the performance of the analytical technique, depending on its robustness, but a generalization may be that the choice of sample preparation

839

M. Ihnat

procedure depends on the analysis technique used and vice versa. In-depth investigations were conducted by Wu¨rfels et al. [90–92] on residues from biological materials after pressure decomposition with nitric acid. Many organic compounds selected to identify with those occurring in biological materials were taken through conventional pressure digestion with nitric acid in PTFE crucibles designed for a maximum temperature of 2008C and pressures of 25 bar. The extent of decomposition was indicated by measurements of residual carbon. Although numerous compounds were completely decomposed, a number of them were refractory, suggesting that incomplete decomposition of natural biological materials will occur. Additionally, inorganic and organic reaction products were formed and characterized. In the third part of the study, these researchers demonstrated that such organic products, which frequently remain after pressure decomposition with nitric acid, lead to serious interferences during inverse square wave anodic stripping voltammetric trace metal determinations, for which quite clean solutions are required. Signals interfering with determination of Zn, Cd, Pb and Cu were caused by reduction of nitrated benzoic acids, which arise from the degradation of phenylalanine in the sample and also by oxidation products of tryptophan. In an earlier paper, Wu¨rfels [93] conducted conventional pressure digestions of two biological CRMs, Mussel NIES-CRM 6, and algae, NIES-CRM 9, using a high pressure asher rated up to a pressure of about 100 bar and temperature of 3208C. Zinc, Cd, Pb and Cu were again determined by square wave anodic stripping voltammetry. His observations were that the voltammograms from the digestion solutions were nearly free of interferences. On the other hand, solutions obtained by pressurized nitric acid decomposition at lower temperature (1808C, in a PTFE vessel with another apparatus) were not suitable. Voltammetric measurements were either inaccurate or impossible, due to a high background current or an interfering signal from nitrobenzoic acid arising from oxidation of phenylalanine. He states that although voltammetric determinations are not viable on such solutions, decomposition at 1808C is well suited for determinations by atomic spectrometric methods; electrochemically active oxidation products can be decomposed completely only at temperatures above 3008C. The necessity of employing different sample decomposition methods according to the sample matrix and the analytical technique being used was also demonstrated by Reid et al. [94]. They reported that for spectrometric techniques, such as ICP-OES, that are tolerant of undigested organic material or for analysis of relatively easily decomposable materials, a single stage digestion with nitric acid produces suitable solutions. Further digestion using hydrogen peroxide signiﬁcantly reduces the residual carbon content which may be important for techniques such as AAS or ICP-MS. Complete decomposition is, however, often needed for electrochemical analysis and this can be readily accomplished using postdigestion with perchloric acid. That fairly complete digestion is needed even for ICP-OES was mentioned by Dolan and Capar [95],

840

Sample preparation for food analysis

who observed that incompletely digested solutions of food products caused severe matrix effects and plasma instability. Electrothermal atomization atomic absorption spectrometry can be affected by interferences remaining after decomposition. One report [96] showed that the nature of the digestion procedure is fundamental to the accurate determination of trace elements such as Cd in complex foodstuffs. Residual nitric acid exhibited a non-speciﬁc absorption at the Cd wavelength; a sulfuric– nitric acid procedure with reﬂuxing was found best. The intensive developmental research by Dabeka indicated severe interferences in the ET-AAS determination of Cd, Co, Ni and Pb when using food digests. Precise and reliable graphite furnace measurements necessitated the separation/preconcentration of the analytes from the acid-digested matrix using ammonium pyrrolidine dithiocarbamate (all four elements) and additionally palladium/ ascorbic acid for lead [97,98]. 25.6.2 Elements As stated at the outset, the goal of sample treatment is the quantitative release of the element of interest from the matrix by total quantitative decomposition, destruction and dissolution of the organic and inorganic constituents of the sample. Perusal of the analytical literature touching on decomposition indicates that, in the main, quantitative or adequate recoveries of elements are reported by the application of a multitude of sample decomposition procedures. Other reports, however, targeting recovery and method performance studies, suggest that this is not always the case and some of the many elements occurring in foods are not quantitatively recovered. Losses of speciﬁc elements and especially volatile ones, such as As, Se, Sb and Hg, can occur during the use of different wet and dry decomposition procedures, but most likely during dry ashing. In-depth research was conducted by Gorsuch [1,72] on the recovery of trace elements in organic and biological materials using radiochemical methods and a cocoa matrix. In his experiments, eight oxidation systems were compared and losses of elements (Cu, Ag, Zn, Cd, Hg, Fe, Co, Cr, Mo, As, Sb, Se, Sr) by volatilization or retention were determined. The eight decomposition procedures were: (1) wet digestion with nitric and perchloric acids, (2) wet digestion with nitric, perchloric and sulfuric acids, (3) wet digestion with nitric and sulfuric acids, (4) the nitric acid method of Middleton and Stuckey [99], (5) dry oxidation, (6) dry oxidation with nitric acid as ashing aid, (7) dry oxidation with sulfuric acid as ashing aid and (8) dry oxidation with magnesium nitrate as ashing aid. In his conclusions, he states that the only conﬁdent generalisation is that wet oxidation of organic material with nitric and perchloric acids proved the most satisfactory for the recovery of all trace elements studied, with the exception of Hg. He adds that no such clear-cut recommendation was possible for dry oxidation.

841

M. Ihnat

Compilations of literature by Sansoni and Iyengar [56] concerning losses during dry ashing of biological materials indicate serious losses (sometimes only at unusually high temperatures) for As, Cd, Cr, Hg, K, Pb and Zn. The complexity of decomposition can be demonstrated with reference to arsenic. In his review of arsenic in the marine and aquatic environments [62], Penrose lists the following considerations for the consistently observed difﬁculties in As recovery: (1) As can be lost from digestion mixtures that are hot and/or have the appropriate composition. As(III) is volatile and oxidizing conditions would be expected to prevent this loss. Fluoride, chloride or bromide would be expected to enhance losses by formation of volatile trihalides, and arsine could be formed during charring of the organic matter. (2) All As is are probably in an organic form. (3) Organoarsenic compounds differ in their stability to oxidation, with methylarsonic and dimethylarsinic acids being refractory to wet digestion. (4) Recovery studies using inorganic As do not assure full release of the organically bound element. 25.6.3 Matrix and constituents The amenability of the sample matrix to proper decomposition is dependent on the interplay of sample composition and decomposition procedure. The major food classes, namely, cereal products, dairy products, eggs and egg products, meat and meat products, ﬁsh and marine products, vegetables, fruit and fruit products, fats and oils, nuts and nut products, sugar and sugar products, beverages, spices and condiments [12], exhibit an extremely wide range of matrix composition. Adding to this list the great variety of feedstuffs available for animal feeding as well as other related biological materials [feeds: hay, corncobs, cannery residues, citrus pulp, milk by-products, mineral premixes and supplements; plants: plant tissues, leaves, roots, wood, vegetation; animal tissues: liver; animal ﬂuids: blood serum; agricultural materials: fertilizers, cottonseed meal, fertilizers containing organic matter (tankage, corncobs), fertilizers containing fritted trace elements; clinical materials: urine; water: drinking water, surface water, saline water, seawater, ground water; organic wastes: wastewater, sewage sludge, manure, composted manure, biosolids, animal waste] expands the matrix composition list. The analyst is thus faced with the task of applying decomposition procedures to adequately deal with these varied matrices, destroying them and releasing the elements of interest. Major constituents having impacts on decomposition/determination are fat, protein, carbohydrate, ﬁbre and ash (including siliceous residue); the latter (ash), including inorganic elements at major or minor levels, at times gives rise to serious interferences in the processing and quantitation steps of the methods. Regarding destruction with nitric and sulfuric acids, Watson [16] indicates that applicability is to most organic materials, including, dyestuffs, medicinal compounds, rubber chemicals, biological materials and foodstuffs (including

842

Sample preparation for food analysis

beverages and fruit juices). The method is not recommended for organic materials containing appreciable amounts of alkaline-earth elements, which give rise to insoluble sulfates which, in term, absorb considerable proportions of trace metals, particularly lead. Five variants are provided for materials that are readily reactive, less reactive, that decompose quietly, that are liable to deﬂagrate violently and a general method applicable to many organic and biological materials. With respect to a nitric/perchloric method, applicability is stated for protein and carbohydrate materials; small amounts of fat are permissible but the method is not recommended for samples containing a high proportion of fat. Its three variants are suitable for use before the determination of most of the common elements, with the exception of Hg. The six variants given for dry ashing are applicable to a range of materials; some disadvantages include incomplete recoveries from some residues, insolubilization of tin compounds, and trapped carbon particles inaccessible to oxidation in certain ﬂour products. In the determination of Al in foods, incorporation of HF treatment into the nitric/perchloric wet digestion was found necessary for most foods and total diet samples. Use of microwave digestion with nitric acid did not improve dissolution of some Al-containing constituents. Peat and plant matrices, containing various amounts of siliceous mineral phases, require the addition of HF in the digestion mixture, or in a separate digestion, to dissolve silicates [100 –102].

25.7

EXAMPLES OF SPECIFIC, RECOMMENDED SAMPLE TREATMENT PROCEDURES

Examples of a few proposed or recommended food decomposition procedures, with experimental details, covering some of the major classes are provided in this section. 25.7.1 Conventional wet digestion with nitric and perchloric acids A procedure involving this acid mixture has been employed in the analytical laboratories of Agriculture and Agri-Food Canada (Ottawa) and by the author in the Ottawa and Summerland laboratories for decades. A summary of the experimental procedure for agricultural –biological, food and related materials, is provided below (mainly from Ref. [103]; also based on Refs. [102,104,105]; and used for RM certiﬁcation [106–112] and for other research by the author [113–116]). Details are included on the preparation of solutions speciﬁcally suitable for F-AAS. Minor variants of the described method have been applied in all of the author’s work and this method is one of those being taken forward for incorporation into an ofﬁcial AOAC International F-AAS/ICP-OES/ICP-MS method development and collaborative study.

843

M. Ihnat

25.7.1.1 Procedure Weigh 1.000 ^ 0.002 g of a dry biological material into a clean, dry 100 ml borosilicate glass volumetric ﬂask dedicated to digestions. Rinse the neck with a few millilitres of deionized water and introduce three 5 mm diameter, previously acid-boiled, pyrex glass beads (earlier digestions were in 100 ml borosilicate glass Kjeldahl ﬂasks, with a variation using custom-made 100 ml quartz Kjeldahl ﬂasks and quartz boiling beads when sodium was desired). Add, from a graduated pipette, 10.0 ml concentrated nitric acid (e.g. Fisher trace metal grade or George Frederick Smith double distilled) and swirl to moisten material. Carefully heat on a hot plate set at a low setting to initiate digestion; close observation and attention are required to prevent foamingover. Alternatively, allow the samples to predigest overnight at room temperature. Swirl occasionally to control foaming. Should excessive foaming occur, immediately remove the volumetric ﬂask from the hot plate, add a few millilitres of deionized water to break the foam, or quickly immerse the bulb of ﬂask under cold running water. Return to heat and continue gentle heating under observation. Increase temperature and boil gently to reduce the volume of nitric acid by approximately one-half. Remove from heat, add 10.0 ml nitric acid þ 5.0 ml (or another speciﬁed volume) of 70% concentrated perchloric acid (e.g. G.F. Smith high purity) or 15.0 ml of a 2 þ 1 mixture of HNO3 and HClO4 and replace on the hot plate. Continue digestion at increasing hot plate setting, observing closely at all times, especially at the nitric-to-perchloric transition stage characterized by rapid surface reaction. Should the solution darken at this stage, remove immediately from heat, allow to cool somewhat, add 2.0 ml nitric acid and continue with the digestion. The hot plate setting should be close to maximum near the end of the procedure to boil-off all the nitric acid and effect boiling of the perchloric acid digest. Bring to perchloric acid fumes and keep at high temperature for approximately 5 min past the initial appearance of fumes. Remove from heat, allow to partially cool, rinse down the neck of the warm ﬂask with about 10 ml of water. The result (at least for biologicals) should be a complete digestion, a water-white solution with perhaps some undigested white siliceous residue or crystals of potassium perchlorate. Voluminous residue can indicate insoluble (at high concentrations of K) potassium perchlorate, which normally will dissolve upon dilution. Allow the ﬂask and contents to cool in a fume hood and bring into laminar ﬂow hood for preparation of solutions. Preparation of solutions for F-AAS determinations: all solutions for F-AAS determinations contain 1000 mg Cs/ml as an ionization suppressant. Add 1.00 ml of 100,000 mg Cs/ml as CsCl solution, using a pipette dedicated to this purpose, to the digest in the 100 ml volumetric ﬂask and dilute to volume with water (solution E). Let stand to allow any suspended silica or other particulate matter to settle before transferring to test tubes. Remove 5.00 ml of solution E without disturbing settled residue, transfer to a 50 ml volumetric ﬂask and bring to volume with diluent 1000 mg Cs/ml in 0.3 N

844

Sample preparation for food analysis

HNO3 (solution I). Let stand to allow any suspended silica or other particulate matter to settle. Remove 1.00 ml of solution E without disturbing settled residue, transfer to a 100 ml volumetric ﬂask and bring to volume with diluent 1000 mg Cs/ml in 0.3 N HNO3 (solution M). Let stand to allow any suspended silica or other particulate matter to settle. Similarly, dilute 5.00 ml of solution M to 50 ml in a volumetric ﬂask with diluent 1000 mg Cs/ml in 0.3 N HNO3 (solution Q). Transfer, without disturbing settled silica, each solution into the required number of 16.5 ml (capped or screw-capped) polypropylene test tubes to provide sufﬁcient solution for all elemental determinations. 25.7.2 Dry ashing with or without ashing aid This procedure is based on those published in Ofﬁcial and Standardized Methods of Analysis 2nd [117] and 3rd editions [16] and by Gorsuch [1] and included in Ref. [12]. It should be suitable for the determination of trace metals in a variety of materials with small modiﬁcations, as necessary. Incorporation of ashing aids, such as H2SO4, MgO or Mg(NO3)2, may be required to facilitate ashing when samples have a low ash content. 25.7.2.1 Procedure Accurately weigh 5 –10 g of sample into a suitable silica or platinum crucible and spread thinly over the bottom. Add ashing aid if required. Dry and char using an infrared lamp, hot plate or burner, taking care not to cause ignition. Place into a cold mufﬂe furnace, raise the temperature slowly to 450 –5008C and heat overnight, again not allowing the contents to ignite. If unoxidized organic matter remains, moisten the residue with water or (1 þ 2) HNO3, evaporate to dryness on a water bath and heat again in the furnace for a further period. When a suitable ash has been obtained, cool, moisten with water, carefully add 10 ml of (1 þ 1) HCl and evaporate to dryness on a water bath. Dissolve the residue in (1 þ 9) HCl or another suitable solvent. 25.7.3 Microwave-assisted wet digestion In the research by Matusiewicz et al. [88] on trace analysis of NRCC biological certiﬁed reference material TORT-1, the following pressure digestion method utilizing nitric acid –hydrogen peroxide and microwave heating was followed: 0.25 g of a dry sample was placed in a PFA-Teﬂon pressure-relief vessel (designed to release pressure in excess of 6.8 atm) and 4 ml each of HNO3 and 30% H2O2 were added. After tightly capping the vessel, the sample carousel containing 12 bombs (one of which was used as a pressure monitor) was placed in the CEM model MDS-81 microwave oven

845

M. Ihnat

and the contents were heated rapidly at full power for 3 min (peak pressure of about 5.5 atm was reached). After completion of the heating cycle, the carousel was removed from the oven and the vessels were cooled in a water bath. They were uncapped, the contents were transferred into a 25 ml calibrated ﬂask and diluted to volume with double distilled water; solution storage was in 30 ml screw-capped polypropylene bottles prior to determination by ET-AAS.

25.8

CLOSING REMARKS

The vast number of publications on research, development and application of sample treatment strategies include a currently increasing number dealing with evaluating various procedures. They deal with assessment of recoveries of elements (by spiking or from certiﬁed reference materials), digestion efﬁciencies and completeness, including fundamental studies of the behaviours of speciﬁc, naturally occurring compounds, and impacts of digests on the performance of the ﬁnal quantitative ﬁnishes (techniques). Because neither sample decomposition nor elemental determination techniques are perfect (completely element speciﬁc and interference free), there is an interrelationship between decomposition and determinations, and the choice of sample preparation procedure depends on the analytical technique used and vice versa. Further critical evaluations of sample decomposition procedures, including evaluation of the extensive literature database and additional experimental studies, appear warranted. Under consideration by the author is a literature evaluation approach “A critical (literature) evaluation of sample decomposition procedures for food and related biological materials for the determination of major, minor and trace elements”. The following items/parameters are to form a basis for evaluation: materials analyzed, reference materials used, reagents used, elements determined, sample treatment method, pretreatment, equipment, automation, scale of operation, procedural details, decomposition completeness, solution preparation/dilution, separation/concentration, solution stability, determinative method and performance. Other supplementary items are: comments and conclusions (by author of publication and evaluator), optimization of sample treatment method, method status assignment, method classiﬁcation/subclassiﬁcation, technique comparisons, collaborative or interlaboratory studies. An experimental investigative phase is also underway of a limited number of targeted biological sample decomposition procedures. This work will contribute towards the development and validation, via interlaboratory collaborative studies, of three related topics dealing with: elements in foods, feeds and biological materials by (1) ﬂame atomic absorption spectrometry, (2) inductively coupled plasma atomic emission spectrometry and (3) inductively coupled plasma mass spectrometry, underway by the author.

846

Sample preparation for food analysis

REFERENCES 1 2 3

4 5

6

7 8 9 10

11 12

13

14

15 16 17

18

19

T.T. Gorsuch, The Destruction of Organic Matter. Pergamon, Oxford, 1970. R. Bock, A Handbook of Decomposition Methods in Analytical Chemistry. Wiley, New York, 1979. J. Dolezal, P. Povondra and Z. Sulcek, Decomposition Procedures in Inorganic Analysis, English translation edited by D.O. Hughes, P.A. Floyd and M.S. Barratt. Illife Books, London, 1986. M. Ihnat, in: W. Horwitz (Ed.), Ofﬁcial Methods of Analysis of AOAC International, 17th ed., AOAC International, Gaithersburg, MD, 2000, Ch. 9. M.A. Joslyn, in: M.A. Joslyn (Ed.), Methods in Food Analysis, Physical, Chemical, and Instrumental Methods of Analysis, 2nd ed., Academic Press, New York, 1970, pp. 109 –140, Ch. 5. K.W. Boyer and W. Horwitz, in: I.K. O’Neill, P. Schuller and L. Fishbein (Eds.), Environmental Carcinogens Selected Methods of Analysis, Vol 8—Some Metals: As, Be, Cd, Cr, Ni, Pb, Se, Zn, IARC Scientiﬁc Publ. No 71, International Agency for Research on Cancer, Lyon, France, 1986, pp. 191– 220, Ch. 12. D. Pearson, The Chemical Analysis of Foods, 7th edn., Churchill Livingstone, Edinburgh, 1976, Ch. 3, pp. 27–106. G.J. DeMenna, in: R.H. Wilson (Ed.), Spectroscopic Techniques for Food Analysis. VCH, Weinheim, 1994, Ch. 5, pp. 147– 179. Y. Pomeranz and C.E. Meloan, Food Analysis, Theory and Practice, Ash and Minerals, 3rd ed., Chapman and Hall, New York, 1994, Ch. 35, pp. 602 –624. B.L. Sharp, in: P.W.J.M. Boumans (Ed.), Inductively Coupled Plasma Emission Spectroscopy Part 2, Applications and Fundamentals, Chem. Anal. New York, Vol. 90. John Wiley and Sons, New York, 1987, pp. 65–99. A. Proctor and J.-F. Meullenet, in: S.S. Nielsen (Ed.), Food Analysis, 2nd ed., Aspen Publishers, Gaithersburg, 1998, pp. 71– 82, Ch. 5. M. Ihnat, in: J.E. Cantle (Ed.), Atomic Absorption Spectrometry, Techniques and Instrumentation in Analytical Chemistry, Vol. 5. Elsevier, Amsterdam, 1982, pp. 139–210, Ch. 4d. F.J.M.J. Maessen, in: P.W.J.M. Boumans (Ed.), Inductively Coupled Plasma Emission Spectroscopy, Part 2, Applications and Fundamentals, Chem. Anal. New York, Vol. 90. Wiley, New York, 1987, pp. 100–150. J. Laporte, G. Kovacsik and J. Bellanger, in: M. Pinta (Ed.), Atomic Absorption Spectrometry, translated by K. M. Greenland and F. Lawson. Halstead Press, New York, 1975, Ch. 9, pp. 240–275. J.C. van Loon, Selected Methods of Trace Analysis: Biological and Environmental Samples. Wiley, Toronto, 1985, Ch. 4, pp. 77– 111. In: C.A. Watson (Ed.), Ofﬁcial and Standardized Methods of Analysis, 3rd edn., Royal Society of Chemistry, Cambridge, 1994, pp. 446–474. E.C. Dunlop, in: I.M. Kolthoff and P.J. Elving (Eds.), Treatise on Analytical Chemistry, Part I, Vol. 2. Interscience Publishers, New York, 1961, pp. 1051–1093. T.T. Gorsuch, in: I.M. Kolthoff and P.J. Elving (Eds.), Treatise on Analytical Chemistry, Part II, Vol. 12. Interscience Publishers, New York, 1965, Section B-1, pp. 295–372. H.T. Delves, in: J.E. Cantle (Ed.), Atomic Absorption Spectrometry, Techniques and Instrumentation in Analytical Chemistry, Vol. 5, Elsevier, Amsterdam, 1982, pp. 341–380, Ch. 4k.

847

M. Ihnat 20 21 22 23 24 25 26

27 28 29

30

31 32

33

34 35

36 37 38 39 40

848

Y.P. Kalra (Ed.), Handbook of Reference Methods for Plant Analysis. 1998. CRC Press, Boca Raton, FL. C.R. Campbell and C.O. Plank, in: Y.P. Kalra (Ed.), Handbook of Reference Methods for Plant Analysis. CRC Press, Boca Raton, FL, Ch. 3, 1998, pp. 37– 49. R.O. Miller, in: Y.P. Kalra (Ed.), Handbook of Reference Methods for Plant Analysis. CRC Press, Boca Raton, FL, 1998, pp. 53–56, Ch. 5. R.O. Miller, in: Y.P. Kalra (Ed.), Handbook of Reference Methods for Plant Analysis. CRC Press, Boca Raton, FL, 1998, pp. 57–61, Ch. 6. Y.P. Kalra and D.G. Maynard, in: Y.P. Kalra (Ed.), Handbook of Reference Methods for Plant Analysis. CRC Press, Boca Raton, FL, 1998, pp. 63–67, Ch. 7. R.O. Miller, in: Y.P. Kalra (Ed.), Handbook of Reference Methods for Plant Analysis. CRC Press, Boca Raton, FL, 1998, pp. 69–73, Ch. 8. J.E. Cantle, in: J.E. Cantle (Ed.), Atomic Absorption Spectrometry, Techniques and Instrumentation in Analytical Chemistry, Vol. 5, Elsevier, Amsterdam, 1982, pp. 37– 66, Ch. 3. O.G. Koch and G.A. Koch-Dedic, Handbuch der Spurenanlyse. Springer-Verlag, Berlin, 1964. M. Stoeppler, in: M. Stoeppler (Ed.), Sampling and Sample Preparation: Practical Guide for Analytical Chemists. Springer, Berlin, 1997, pp. 132– 141, Ch. 11. E. Jackwerth and M. Wu¨rfels, translated by P.H.E. Gardiner, In: M. Stoeppler (Ed.), Sampling and Sample Preparation: Practical Guide for Analytical Chemists. Springer, Berlin, 1997, pp. 142–154, Ch. 12. J. Begerow and L. Dunemann, in: M. Stoeppler (Ed.), Sampling and Sample Preparation: Practical Guide for Analytical Chemists. Springer, Berlin, 1997, pp. 155–166, Ch. 13. P. Ostapczuk, in: M. Stoeppler (Ed.), Sampling and Sample Preparation: Practical Guide for Analytical Chemists. Springer, Berlin, 1997, pp. 167– 182, Ch. 14. D.J. Butcher and J. Sneddon, A Practical Guide to Graphite Furnace Atomic Absorption Spectrometry, Chem. Anal. New York, Vol. 149. Wiley, New York, 1998, Ch. 6, 119–174. C.H. Lim and M.L. Jackson, in: A.L. Page, R.H. Miller and D.R. Keeney (Eds.), Methods of Soil Analysis, Part 2, Chemical and Microbiological Properties, 2nd ed., American Society of Agronomy, Soil Science Society of America, Madison, 1982, pp. 1– 12, Ch. 1. U. Kurfu¨rst (Ed.), Solid Sample Analysis, Direct and Slurry Sampling Using GFAAS and ETV-ICP. Springer, Berlin, 1998. M. Ihnat, Selenium, In: M. Stoeppler (Ed.), Hazardous Metals in the Environment, Techniques and Instrumentation in Analytical Chemistry, Vol. 12, Elsevier Science, Amsterdam, 1992, pp. 475– 515, Ch. 16. R.J. Shamberger, in: R.J. Shamberger (Ed.), Biochemistry of Selenium. Plenum Press, New York, 1983, Ch. 10, pp. 311– 327. J.H. Watkinson, in: O.H. Muth, J.E. Oldﬁeld and P.H. Weswig (Eds.), Selenium in Biomedicine. Avi Publishing Co., Westport, 1967, pp. 97–117, Ch. 6. W.C. Cooper, in: R.A. Zingaro and W.C. Cooper (Eds.), Selenium. Van Nostrand Reinhold, New York, 1974, pp. 615–653, Ch. 10. H.D. Drew, in: C.A. Streuli and P.R. Averell (Eds.), Analytical Chemistry of Nitrogen and its Compounds, Part 1, 1970, pp. 1– 27, Ch. 1. R.B. Bradstreet, The Kjeldahl Method for Organic Nitrogen, Academic Press, (1965) pp. 1– 239.

Sample preparation for food analysis 41

42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73

74 75

G.M. Gustin and C.L. Ogg, in: I.M. Kolthoff and P.J. Elving (Eds.), Treatise on Analytical Chemistry, Part II, Vol. 11. Interscience Publishers, New York, 1965, Section B-1, pp. 405– 498. K.J. Lamble and S.J. Hill, Analyst, 123 (1998) 103R–133R. Z.A. deBono, M. Velosa, C. Ceccarelli, M. de la Guardia and A. Salvador, Fresenius J. Anal. Chem., 339 (1991) 235–239. H. Matusiewicz and R.E. Sturgeon, Prog. Anal. Spectrosc., 12 (1989) 21– 39. B. Sansoni and V.K. Panday, in: S. Facchetti (Ed.), Analytical Techniques for Heavy Metals in Biological Fluids. Elsevier, Amsterdam, 1983, pp. 91– 131. F.E. Smith and E.A. Arsenault, Talanta, 43 (1996) 1207–1268. A.M. Ure, L.R.P. Butler, R.O. Scott and R. Jenkins, Spectrochim. Acta B, 52 (1997) 409–420. G. To¨lg, Pure Appl. Chem., 55 (1983) 1989– 2006. C. Bendicho and M.T.C. de Loos-Vollebregt, J. Anal. At. Spectrom., 6 (1991) 353–374. M. Hoenig, Talanta, 54 (2001) 1021– 1038. Z. Sulcek, P. Povondra and J. Dolezal, CRC Crit. Rev. Anal. Chem., 6 (1977) 255–323. A.M. Ure, Anal. Chim. Acta, 76 (1975) 1– 26. J.L. Burguera and M. Burguera, Analyst, 123 (1998) 561– 569. B. Griepink and G. To¨lg, Pure Appl. Chem., 61 (1989) 1139–1146. E. Jackwerth and S. Gomiscek, Pure Appl. Chem., 56 (1984) 479–489. B. Sansoni and V. Iyengar, Spezielle Berichte Kernforschungsanlage Ju¨lich, JulSpez-13 (1978) 1– 79. S.K.C. Chang, E. Holm, J. Schwarz and P. Rayas-Duarte, Anal. Chem., 67 (1995) 127R–153R. A. Taylor, S. Branch, H.M. Crews, D.J. Halls, L.M.W. Owen and M. White, J. Anal. At. Spectrosc., 12 (1997) 119R– 221R. A. Taylor, S. Branch, D.J. Halls, L.M.W. Owen and M. White, J. Anal. At. Spectrosc., 15 (2000) 451– 487. W. Horwitz, J. Assoc. Off. Anal. Chem., 62 (1979) 1251–1264. Y. Talmi and C. Feldman, in: E.A. Woolson (Ed.), Arsenical Pesticides, ACS Symposium Series, No. 7, 1975, pp. 13– 34. W.R. Penrose, CRC Crit. Rev. Environ. Control, 4 (1974) 465– 482. F.J. Langmyhr and G. Wibetoe, Prog. Anal. At. Spectrosc., 8 (1985) 193–256. J.M. Carey and J.A. Caruso, CRC Crit. Rev. Anal. Chem., 23 (1992) 379– 439. S. Jacobs, CRC Crit. Rev. Anal. Chem., 7 (1978) 297– 322. G. To¨lg, Trace Elem. Anal. Chem. Biol. Med., 3 (1984) 95– 125. G. To¨lg, Anal. Chim. Acta, 283 (1993) 3 –18. O.E. Olson, I.S. Palmer and E.I. Whitehead, Met. Biochem. Anal., 21 (1973) 39–78. M. Stoeppler, Trace Elem. Anal. Chem. Med. Biol., 2 (1983) 909– 927. K. Robards and P. Worsfold, Analyst, 116 (1991) 549– 568. A. Ferrari and E. Cantanzaro, Ann. N. Y. Acad. Sci., 130 (1965) 602 –620. T.T. Gorsuch, Analyst, 84 (1959) 135– 173. Food and Agriculture Organization of the United Nations, Manuals of Food Quality Control, 2. Additives Contaminants Techniques, FAO Food and Nutrition Paper 14/2, Food and Agriculture Organization of the United Nations, Rome, 1980. E.V. Williams, J. Food Technol., 13 (1978) 367 –384. F. Baruthio, O. Guillard, J. Arnaud, F. Pierre, F.E. Smith, E.A. Arsenault and R. Zawissiak, Clin. Chem., 34 (1988) 227– 234.

849

M. Ihnat 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103

104 105 106

850

F.E. Smith and E.A. Arsenault, Talanta, 43 (1996) 1207– 1268. A.M. Ure, L.R.P. Butler, R.O. Scott and R. Jenkins, Spectrochim. Acta B, 52 (1997) 409 –420. C. Bendicho and M.T.C. de Loos-Vollebregt, J. Anal. At. Spectrom., 6 (1991) 353 –374. In: M. Stoeppler (Ed.), Sampling and Sample Preparation: Practical Guide for Analytical Chemists. Springer, Berlin, 1997, pp. 1– 202. Z. Yu, C. Vandecasteele, B. Desmet and R. Dams, Mikrochim. Acta, 1 (1990) 41–48. M. Ihnat, Fluorometric determination of selenium in foods, J. Assoc. Off. Anal. Chem., 57 (1974) 368 –372. R.L. Sims, L.M. Mullen and D.B. Milne, J. Food Compos. Anal., 3 (1990) 27–37. B.A. Zarcinas, B. Cartwright and L.R. Spouncer, Commun. Soil Sci. Plant Anal., 18 (1987) 131–146. R.S.S. da Costa, M. das Gracas Tavares do Carmo, C. Saunders, R.T. Lopes, E.F.O. de Jesus and S.M. Simabuco, J. Food Compos. Anal., 15 (2002) 27–33. R.E. Sturgeon, J.W. McLaren, S.N. Willie, D. Beauchemin and S.S. Berman, Can. J. Chem., 65 (1987) 961– 964. M. Xie, A. Von Bohlen, R. Klockenka¨mper, X. Jian and K. Gu¨nther, Z. Lebensm. Unters Forsch. A, 207 (1998) 31–38. A. Abu-Samra, J.S. Morris and S.R. Koirtyohann, Anal. Chem., 47 (1975) 1475–1477. H. Matusiewicz, R.E. Sturgeon and S.S. Berman, J. Anal. At. Spectrom., 4 (1989) 323 –327. L.L. Jackson, E.E. Engleman and J.L. Peard, Environ. Sci. Technol., 19 (1985) 437 –441. M. Wu¨rfels, E. Jackweth and M. Stoeppler, Anal. Chim. Acta, 226 (1989) 1– 16. M. Wu¨rfels, E. Jackweth and M. Stoeppler, Anal. Chim. Acta, 226 (1989) 17–30. M. Wu¨rfels, E. Jackweth and M. Stoeppler, Anal. Chim. Acta, 226 (1989) 31–41. M. Wu¨rfels, Mar. Chem., 28 (1989) 259 –264. H.J. Reid, S. Greenﬁeld and T.E. Edmonds, Analyst, 120 (1995) 1543–1548. S.P. Dolan and S.G. Capar, J. Food Compos. Anal., 15 (2002) 593 –615. M.T. Cabanis, G. Cassanas, J.C. Cabanis and S. Brun, J. Assoc. Off. Anal. Chem., 71 (1988) 1033– 1037. R.W. Dabeka, Sci. Total Environ., 89 (1989) 271–277. R.W. Dabeka, Final Program of the 17th Annual Meeting of the Federation of Analytical Chemistry and Spectroscopy Societies, 1990, p.102. G. Middleton and R.E. Stuckey, Analyst (Lond.), 79 (1954) 138–142; Z. Gu¨nther, Lebensm. Unters Forsch. A, 207 (1998) 31–38. M. Krachler, C. Mohl, H. Emons and W. Shotyk, J. Anal. At. Spectrom., 17 (2002) 844–851. C.S.E. Papp and T.F. Harms, Analyst, 110 (1985) 237 –242. M. Ihnat, Commun. Soil Sci. Plant Anal., 13 (1982) 969 –979. M. Ihnat, Technical Laboratory Information Report (TLIR-5), Paciﬁc Agri-Food Research Centre—Summerland, Agriculture and Agri-Food Canada, Summerland, BC, 2002, p. 9. M. Ihnat, Fresenius Z. Anal. Chem., 326 (1987) 739–741. M. Ihnat, J. Res. NBS, 93 (1988) 354– 358. M. Ihnat and W.R. Wolf, in: W.R. Wolf (Ed.), Biological Reference Materials: Availability, Uses and Need for Validation of Nutrient Measurement. Wiley, New York, NY, 1985, pp. 141 –165, Ch. 8.

Sample preparation for food analysis 107

108 109 110 111 112 113 114 115 116 117 118 119 120 121 122 123 124 125 126 127 128 129 130 131 132 133 134 135 136 137 138 139 140 141

M. Ihnat, Development of a new series of agricultural/food reference materials for analytical quality control of elemental determinations, J. AOAC Int., 77 (1994) 1605– 1627. M. Ihnat and M.S. Wolynetz, Fresenius J. Anal. Chem., 348 (1994) 452 –458. M. Ihnat, J. Anal. Chem., 348 (1994) 459–467. M. Ihnat, Fresenius J. Anal. Chem., 348 (1994) 474– 478. M. Ihnat, Fresenius J. Anal. Chem., 348 (1994) 468–473. M. Ihnat, J. Radioanal. Nucl. Chem., 245 (2000) 65– 72. M. Ivan, M. Ihnat and D.M. Veira, Anim. Feed Sci. Technol., 9 (1983) 131–142. M. Ivan, M. Ihnat and D.M. Veira, Can. J. Anim. Sci., 63 (1982) 163– 171. M. Ihnat and L. Fernandez, Biores. Technol., 57 (1996) 143– 156. M. Ihnat, G.H. Neilsen and E.J. Hogue, Commun. Soil Sci. Plant Anal., 31 (2000) 803–825. N.W. Hanson (Ed.), Ofﬁcial, Standardized and Recommended Methods of Analysis, 2nd ed., The Society for Analytical Chemistry, London, 1973. J. Kucera, Z. Randa and L. Soukal, J. Radioanal. Nucl. Chem., 249 (2001) 61–65. D.L. Anderson, J. Radioanal. Nucl. Chem., 244 (2000) 225– 229. F.A.R. Martino, M.L.F. Sanchez and A.S. Medel, J. Anal. At. Spectrom., 15 (2000) 163– 168. P.A. Brereton, P. Robb, C.M. Sargent, H.M. Crews and R. Wood, J. Assoc. Off. Anal. Chem. Int., 80 (1997) 1287–1297. T. Shimamura and M. Iwashita, Anal. Sci., 13 (1997) 177–192. I.L. Garcia, P. Vinas, N. Campillo and M.H. Cordoba, Fresenius J. Anal. Chem., 355 (1996) 57– 64. D.A. Becker, J. Radioanal. Nucl. Chem., 193 (1995) 25– 32. J. Omote, H. Kohno and K. Toda, Anal. Chim. Acta, 307 (1995) 117–126. N. Carrion, A. Itriago, M. Murillo, E. Eljuri and A. Fernandez, J. Anal. At. Spectrom., 9 (1994) 205–207. J.M. Anzano, M.P. Martinez-Garbayo, M.A. Belarra and J.R. Castillo, J. Anal. At. Spectrom., 9 (1994) 125–128. W.R. Mindak, J. Assoc. Off. Anal. Chem. Int., 77 (1994) 1023– 1030. M. Martin-Polvillo, T. Albi and A. Guinda, J. Am. Oil Chem. Soc., 71 (1994) 347– 353. D. Firestone, J. Assoc. Off. Anal. Chem. Int., 77 (1994) 951– 954. F. Dorey, D. Barillier, F. Bosque, V. Paraire and J.F. Le Querler, Analusis, 21 (1993) 49–52. D.L. Anderson, W.C. Cunningham and G.H. Alvarez, J. Radioanal. Nucl. Chem., 167 (1993) 139 –144. H. Schauenburg and P. Weigert, Fresenius J. Anal. Chem., 342 (1992) 950–956. K.D. Frank, J. Burch and J. Denning, Commun. Soil Sci. Plant Anal., 23 (1992) 2415– 2424. S. Plaami and J. Kumpulainen, J. Assoc. Off. Anal. Chem., 74 (1991) 32–36. A. Moauro and P.L. Carconi, J. Radioanal. Nucl. Chem., 151 (1991) 149– 157. J. Arnaud, A. Favier and J. Alary, J. Anal. At. Spectrom., 6 (1991) 647–652. R.D. Parker, J. Assoc. Off. Anal. Chem., 73 (1990) 721–723. S.G. Capar, J. Assoc. Off. Anal. Chem., 73 (1990) 320 –321. J.J. Van der Lee, I. Walinga, P.K. Manyeki, V.J.G. Houba and I. Novozamsky, Commun. Soil Sci. Plant Anal., 18 (1987) 789–802. A.M. Nash, T.l. Mounts and W.F. Kwolek, J. Am. Oil Chem. Soc., 60 (1983) 811 –814.

851

M. Ihnat 142 143 144 145 146 147 148 149

150 151 152 153 154 155 156 157 158 159 160 161 162 163 164 165 166 167 168 169 170 171 172 173 174 175 176 177 178 179 180

852

H.J. Bakker, J. Assoc. Off. Anal. Chem., 60 (1977) 1307–1309. J.T. Olejko, J. Am. Oil Chem. Soc., 53 (1976) 480– 484. M.A. Perring, J. Sci. Food Agric., 25 (1974) 237– 245. G.E. Leggett and D.T. Westermann, J. Agric. Food Chem., 21 (1973) 65– 69. J.H. Baker and T. Greweling, J. Agric. Food Chem., 15 (1967) 340– 344. I. Boukari, N.W. Shier, X.E. Fernandez, J. Frisch, B.A. Watkins, L. Pawloski and A.D. Fly, J. Food Compos. Anal., 14 (2001) 37– 42. A.P. Udoh, Talanta, 52 (2000) 749– 754. C.P. Sanchez-Castillo, P.J.S. Dewey, A. Aguirre, J.J. Lara, R. Vaca, P. Leon de la Barra, M. Ortiz, I. Escamilla and W.P.T. James, J. Food Compos. Anal., 11 (1998) 340–356. M. Mu¨ller, M. Anke and H. Illing-Gu¨nther, Z. Lebensm. Unters Forsch. A, 205 (1997) 170– 173. S. Anzalone, E. Bottari and M.R. Festa, Anal. Chim. Acta, 343 (1997) 241–249. L. Jorhem and B. Sundstro¨m, Z. Lebensm. Unters Forsch., 201 (1995) 311–316. M. Yaman and S. Gucer, Analusis, 23 (1995) 168–171. L. Jorhem, J. Assoc. Off. Anal. Chem. Int., 76 (1993) 798–813. M.V. Polo, M.J. Lagarda and R. Farre, J. Food Compos. Anal., 5 (1992) 77– 83. R.M. Pollman, J. Assoc. Off. Anal. Chem., 74 (1991) 27– 31. W.J. Blanchﬂower, A. Cannavan and D.G. Kennedy, Analyst, 115 (1990) 1323–1325. G. Brito, C. Diaz, L. Galindo, A. Hardisson, D. Santiago and F.G. Montelongo, Bull. Environ. Contam. Toxicol., 44 (1990) 309– 316. P. Thomas, Analusis, 18 (1990) 562– 564. J. Gzyl, Sci. Total Environ., 96 (1990) 199–209. I. Ockenden and J.N.A. Lott, Commun. Soil Sci. Plant Anal., 17 (1986) 627– 644. W.G. de Ruig, J. Assoc. Off. Anal. Chem., 69 (1986) 1009–1013. A. Yashi, H. Koizumi, T. Suzuki and C. Tsutsumi, Bunseki Kagaku, 35 (1986) T115– T119. R. Ooms and W. Van Pee, J. Am. Oil Chem. Soc., 60 (1983) 957 –960. T.-S. Koh, Anal. Chem., 55 (1983) 1814–1815. C.A. Rowan, O.T. Zajicek and E.J. Calabrese, Anal. Chem., 54 (1982) 149 –151. I. Sarudi Jr., Fresenius Z. Anal. Chem., 303 (1980) 197–199. S.R. Koirtyohann and C.A. Hopkins, Analyst, 101 (1976) 870 –875. J.B. Jones Jr., J. Assoc. Off. Anal. Chem., 58 (1975) 764 –769. B.E. Guthrie, New Zealand Med. J., 82 (1975) 418–424. R.A. Issac and B.J. Jones Jr., Commun. Soil Sci. Plant Anal., 3 (1972) 261– 269. A.L. Gelman, J. Sci. Food Agric., 23 (1972) 299–305. D. Pokorn, V. Stibilj, B. Gregoric, M. Dermelj and J. Stupar, J. Food Compos. Anal., 11 (1998) 47–53. M. Leonhardt and C. Wenk, J. Food Compos. Anal., 10 (1997) 218 –224. N.J. Miller-Ihli, J. Food Compos. Anal., 9 (1996) 290–300. M.L. Cervera, J.C. Lopez and R. Montoro, Microchem. J., 49 (1994) 20–26. M.E. Coleman, R.S. Elder, P. Basu and G.P. Koppenal, J. Assoc. Off. Anal. Chem. Int., 75 (1992) 615–625. N.J. Miller-Ihli and F.E. Greene, J. Assoc. Off. Anal. Chem. Int., 75 (1992) 354– 359. N. Ybanez, M.L. Cervera and R. Montoro, Anal. Chim. Acta, 258 (1992) 61– 71. E. Coni, A. Stacchini, S. Caroli and P. Falconieri, J. Anal. At. Spectrom., 5 (1990) 581–586.

Sample preparation for food analysis 181 182 183 184 185 186 187 188 189 190 191 192 193 194 195 196 197 198 199 200 201 202 203 204 205 206 207 208 209 210 211 212 213 214 215

M. Luisa, A. Navarro, R. Montoro, R. Catala and N. Ybanez, J. Assoc. Off. Anal. Chem., 72 (1989) 282– 285. R.D. Satzger, Anal. Chem., 60 (1988) 2500– 2504. J. Pavel, M. Fille and U. Frey, Fresenius Z. Anal. Chem., 332 (1988) 51– 54. A. Lopez and H.L. Williams, J. Food Prot., 51 (1988) 574– 576. D.M. Sullivan, D.F. Kehoe and R.L. Smith, J. Assoc. Off. Anal. Chem., 70 (1987) 118–120. A.M.G. Van Betteray-Kortekaas and G. Vos, Fresenius Z. Anal. Chem., 323 (1986) 492– 493. T. Muys, Analyst, 109 (1984) 119– 121. R.D. Satzger, C.S. Clow, E. Bonnin and F.L. Fricke, J. Assoc. Off. Anal. Chem., 65 (1982) 987– 991. R.J. Gajan, S.G. Capar, C.A. Subjoc and M. Sanders, J. Assoc. Off. Anal. Chem., 65 (1982) 970– 977. J.R. Preer, B.R. Stephens and C.W. Bland, J. Assoc. Off. Anal. Chem., 65 (1982) 1010– 1015. D.L. Heanes, Analyst, 106 (1981) 172 –181. M. Feinberg and C. Ducauze, Anal. Chem., 52 (1980) 207–209. J.T. Kumpulainen, W.R. Wolf, C. Veillon and W. Mertz, J. Agric. Food Chem., 27 (1979) 490– 494. J.W. Jones, R.J. Gajan, K.W. Boyer and J.A. Fiorino, J. Assoc. Off. Anal. Chem., 60 (1977) 826– 832. P.J. Leblanc and D.L. Jackson, J. Assoc. Off. Anal. Chem., 56 (1973) 383–386. A.J. Mitteldorf and D.O. Landon, Anal. Chem., 24 (1952) 469– 472. M. Hilp, Fresenius J. Anal. Chem., 360 (1998) 184 –191. D. Erber, L. Quick, F. Winter, J. Roth and K. Cammann, Fresenius J. Anal. Chem., 349 (1994) 502 –509. B.R. Norman and G.V. Iyengar, Fresenius J. Anal. Chem., 358 (1994) 430– 432. R.C. Campos, A.J. Curtius and H. Berndt, J. Anal. At. Spectrom., 5 (1990) 669– 673. S.E. Raptis, G. Knapp and A.P. Schalk, Fresenius Z. Anal. Chem., 316 (1983) 482– 487. G. Knapp, S.E. Raptis, G. Kaiser, G. To¨lg, P. Schramel and B. Schreiber, Fresenius Z. Anal. Chem., 308 (1981) 97– 103. M. Allegrini, K.W. Boyer and J.T. Tanner, J. Assoc. Off. Anal. Chem., 64 (1981) 1111 –1115. W.J. Kirsten, Anal. Chem., 51 (1979) 1173 –1179. T. Stijve, Z. Lebensm. Unters.-Forsch., 164 (1977) 201– 203. R.M. Welch and E.E. Cary, J. Agric. Food Chem., 23 (1975) 479–482. L.H. Scroggins, J. Assoc. Off. Anal. Chem., 58 (1975) 146–149. W. Wolf, W. Mertz and R. Masironi, J. Agric. Food Chem., 22 (1974) 1037– 1042. D.F. Ball, M. Barber and P.G.T. Vossen, Sci. Total Environ., 2 (1973) 101 –110. I. Okuno, R.A. Wilson and R.E. White, J. Assoc. Off. Anal. Chem., 55 (1972) 96–100. R. Tkachuk and F.D. Kuzina, J. Sci. Food Agric., 23 (1972) 1183–1195. D.A. Shearer and G.F. Morris, Microchem. J., 15 (1970) 199– 204. L. Kosta and A.R. Byrne, Talanta, 26 (1969) 1297– 1303. P. Van den Winkel, A. Speecke and J.J. Hoste, IAEA STI/PUB/155, 1967, pp. 159 –172 W.H. Gutenmann and D.J. Lisk, J. Agric. Food Chem., 9 (1961) 488– 489.

853

M. Ihnat 216 217 218 219 220 221 222 223 224 225 226 227 228 229 230 231 232 233 234 235 236 237 238 239 240 241 242 243 244 245 246 247 248 249 250

854

T. Weizhi, N. Bangfa, W. Pingsheng, N. Huiling, C. Lei and Z. Yangmei, J. Radioanal. Nucl. Chem., 249 (2001) 25– 28. C.R.T. Tarley, W.K.T. Coltro, M. Matsushita and N.E. de Souza, J. Food Compos. Anal., 14 (2001) 611– 617. E.M.R. Rodriguez, M.S. Alaejos and C.D. Romero, J. Food Comp. Anal., 14 (2001) 419–430. M.C.E. Lomer, R.P.H. Thompson, J. Commisso, C.l. Keen and J.J. Powell, Analyst (Lond.), 125 (2000) 2339– 2343. B. Danko, H. Polkowska-Motrenko and R. Dybczynski, J. Radioanal. Nucl. Chem., 246 (2000) 279– 283. Y. Hu and A.V. Barker, Commun. Soil Sci. Plant Anal., 30 (1999) 677 –687. J.S. McGuire and D.A. Hite, J. Assoc. Off. Anal. Chem. Int., 81 (1998) 923 –930. S. Bhattacharjee, P. Dasgupta, A.R. Paul, S. Ghosal, K.K. Pahdi and L.P. Pandey, J. Sci. Food Agric., 77 (1998) 456–458. E. Ivanova, K. Benkhedda and F. Adams, J. Anal. At. Spectrom., 13 (1998) 527–531. N.J. Miller-Ihli, J. Food Compos. Anal., 9 (1996) 301–311. P. Masson and J.M. Esvan, Analusis, 23 (1995) 437–440. H. Muto, T. Abe, Y. Takizawa, K. Kawabata, K. Yamaguchi and K. Saitoh, Sci. Total Environ., 144 (1994) 231–239. S.B. Adeloju, H.S. Dhindsa and R.K. Tandon, Anal. Chim. Acta, 285 (1994) 359–364. F. Zhao, S.P. McGrath and A.R. Crosland, Commun. Soil Sci. Plant Anal., 25 (1994) 407– 418. I. Novozamsky, V.J.G. Houba, J.J. Van der Lee, R. Van Eck and M.D. Mignorance, Commun. Soil Sci. Plant Anal., 24 (1993) 2595– 2605. A.A. Hafez, S.S. Goyal and D.W. Rains, Agron. J., 83 (1991) 148–153. J.Y. Yang, M.H. Yang and S.M. Lin, Anal. Chem., 62 (1990) 146 –150. W.A. Grant and P.C. Ellis, J. Anal. At. Spectrom., 3 (1988) 815– 820. A. Dong, V.V. Rendig, R.G. Burau and G.S. Besga, Anal. Chem., 59 (1987) 2728– 2730. B. Ogorevc, A. Krasna and V. Hudnik, Anal. Chim. Acta, 196 (1987) 183 –191. N.K. Veien and M.R. Andersen, Acta Dermato. Venereol., 66 (1986) 502–509. S. Munro, L. Ebdon and D.J. McWeeny, J. Anal. At. Spectrom., 1 (1986) 211– 219. W.H. Evans and J.I. Read, Analyst (Lond.), 110 (1985) 619– 623. A.A.-R. Gharaibeh, J. Eagles and R. Self, Biomed. Mass Spectrom., 12 (1985) 344–347. C.D. Salisbury and W. Chan, J. Assoc. Off. Anal. Chem., 68 (1985) 218– 219. R.F. Suddendorf and K.K. Cook, J. Assoc. Off. Anal. Chem., 67 (1984) 985 –992. J.W. Jones, S.G. Capar and T.C. O’Haver, Analyst (Lond.), 107 (1982) 353– 377. S.A.K. Hsieh, Y.S. Chong, J.F. Tan and T.S. Ma, Mikrochim. Acta, 2 (1982) 337–346. H. Agemian, D.P. Sturtevant and K.D. Austen, Analyst (Lond.), 105 (1980) 125–130. M.S. Cresser and J.W. Parsons, Anal. Chim. Acta, 109 (1979) 431– 436. M. Ihnat and H.J. Miller, J. Assoc. Off. Anal. Chem., 60 (1977) 813 –825. J.A. Fiorino, J.W. Jones and S.G. Capar, Anal. Chem., 48 (1976) 120– 125. M. Malaiyandi and J.P. Barrette, Arch. Environ. Contam., 2 (1974) 315– 330. R.A. Baetz and C.T. Kenner, J. Agric. Food Chem., 21 (1973) 436 –440. P.E. Bantz and D.W. Whitney, Proc. Am. Soc. Brew. Chem., (1971) 183–191.

Sample preparation for food analysis 251 252 253 254 255 256 257 258 259 260 261 262 263 264 265 266 267 268 269 270 271 272 273 274 275 276 277 278 279 280 281

M. Haldimann, A. Eastgate and B. Zimmerli, Analyst (Lond.), 125 (2000) 1977–1982. V.-A. Catsiki and E. Strogyloudi, Sci. Total Environ., 237/238 (1999) 387–400. R.P. Pettersson and M. Olsson, J. Anal. At. Spectrom., 13 (1998) 609–613. E.H. Larsen and M.B. Ludwigsen, J. Anal. At. Spectrom., 12 (1997) 435– 439. D. Amarasiriwardena, M. Kotrbai, A. Krushevska and R.M. Barnes, Can. J. Anal. Sci. Spectrosc., 42 (1997) 69–78. B. Gawlik, M. Druges, M. Bianchi, A. Bortoli, A. Kettrup and H. Muntau, Fresenius J. Anal. Chem., 358 (1997) 441–445. M. Schuhmacher, J.L. Domingo, J.M. Llobet and J. Corbella, Bull. Environ. Contam. Toxicol., 50 (1993) 514– 521. J.E. Tahan, V.A. Granadillo, J.M. Sanchez, H.S. Cubillan and R.A. Romero, J. Anal. At. Spectrom., 8 (1993) 1005–1010. K. Gu¨nther, A. Von Bohlen, G. Paprott and R. Klockenka¨mper, Fresenius J. Anal. Chem., 342 (1992) 444– 448. R.W. Dabeka and A.D. McKenzie, J. Assoc. Off. Anal. Chem., 75 (1992) 954 –963. M. Plessi and A. Monzani, J. Assoc. Off. Anal. Chem., 73 (1990) 798– 800. K. May and M. Stoeppler, Fresenius Z. Anal. Chem., 317 (1984) 248 –251. L. Kotz, G. Kaiser, P. Tscho¨pel and G. To¨lg, Fresenius Z. Anal. Chem., 260 (1972) 207– 209. O. Mestek, J. Kominkova, R. Koplik and M. Suchanek, Talanta, 54 (2001) 927– 934. R. Garcia and B. Kahn, J. Radioanal. Nucl. Chem., 250 (2001) 85– 91. Y. Miyamoto, A. Kajikawa, J.H. Zaidi, T. Nakanishi and K. Sakamoto, J. Radioanal. Nucl. Chem., 243 (2000) 747 –765. X. Feng, S. Wu, A. Wharmby and A. Wittmeier, J. Anal. At. Spectrom., 14 (1999) 939– 946. M.A. Torres, J. Verdoy, A. Alegria, R. Barbera, R. Farre and M.J. Lagarda, Sci. Total Environ., 228 (1999) 185–192. T.D. Phuong, P.V. Chuong, D.T. Khiem and S. Kokot, Analyst (Lond.), 124 (1999) 553– 560. M.F. Enriquez-Dominguez, M.C. Yebra-Biurrun and M.P. Bermejo-Barrera, Analyst (Lond.), 123 (1998) 105 –108. A. Bibak, A. Behrens, S. Stu¨rup, L. Knudsen and V. Gundersen, J. Agric. Food Chem., 46 (1998) 3146– 3149. C.J. Park and J.K. Suh, J. Anal. At. Spectrom., 12 (1997) 573– 577. E.J. Gawalko, T.W. Nowicki, J. Babb, R. Tkachuk and S. Wu, J. Assoc. Off. Anal. Chem., 80 (1997) 379– 387. M.A. de la Fuente, G. Guerrero and M. Juarez, J. Agric. Food Chem., 43 (1995) 2406– 2410. J. Ziaziaris and J.L. Kacprzak, J. Assoc. Off. Anal. Chem. Int., 78 (1995) 874– 877. M.D. Mingorance, M.L. Perez-Vazquez and M. Lachica, J. Anal. At. Spectrom., 8 (1993) 853– 858. P.J. Oles and W.M. Graham, J. Assoc. Off. Anal. Chem., 74 (1991) 812– 814. S. Nakashima, R.E. Sturgeon, S.N. Willie and S.S. Berman, Analyst (Lond.), 113 (1988) 159– 163. H.M. Kingston and L.B. Jassie, Anal. Chem., 58 (1986) 2534– 2541. P. Jacob and H. Berndt, J. Anal. At. Spectrom., 17 (2002) 1615–1620. E.R. Pereira-Filho, H. Berndt and M.A.Z. Arruda, J. Anal. At. Spectrom., 17 (2002) 1308– 1315.

855

M. Ihnat 282 283 284 285 286 287 288 289 290 291 292 293 294

856

S. Ehsan and W.D. Marshall, J. Anal. At. Spectrom., 16 (2001) 1180–1184. P. Vinas, M. Pardo-Martinez, I. Lopez-Garcia and M. Hernandez-Cordoba, J. Anal. At. Spectrom., 16 (2001) 1202–1205. M. Gonzalez, M. Gallego and M. Valcarel, Talanta, 48 (1999) 1051–1060. E.C. Lima, F.J. Krug, A.T. Ferreira and F. Barbossa Jr., J. Anal. At. Spectrom., 14 (1999) 269– 274. N.N. Meeravali and S.J. Kumar, J. Anal. At. Spectrom., 13 (1998) 647–652. P. Bermejo-Barrera, A. Moreda-Pineiro, J. Moreda-Pineiro and A. BermejoBarrera, Fresenius J. Anal. Chem., 360 (1998) 707– 711. E.R. Pereira, J.J.R. Rohwedder and M.A.Z. Arruda, Analyst (Lond.), 123 (1998) 1023– 1028. G.S.B. Jannuzi, F.J. Krug and M.A.Z. Arruda, J. Anal. At. Spectrom., 12 (1997) 375–378. Y. Tan, J.-S. Blais and W.D. Marshall, Analyst (Lond.), 121 (1996) 1419– 1424. N.J. Miller-Ihli, J. Anal. At. Spectrom., 9 (1994) 1129– 1134. P. Vinas, N. Campillo, I.L. Garcia and M.H. Cordoba, Anal. Chim. Acta, 283 (1993) 393–400. Y. Madrid, M. Bonilla and C. Camara, J. Anal. At. Spectrom., 4 (1989) 167–169. O.G. Koch and G.A. Koch-Dedic, Handbuch der Spurenanlyse. Springer, Berlin, 1964.

Chapter 26

The determination of trace elements in water Scott N. Willie

This chapter summarizes various methods used for the determination of trace elements in natural waters. Space limitations, and the vast number of publications on this subject, prevent a comprehensive review. However, appropriate methods, recent developments and future trends are highlighted. Several criteria were considered when selecting methods to be included here; the use of relevant certiﬁed reference materials (CRMs) to demonstrate validation, appropriate limits of detection (LODs) and novel approaches to analysis. Apologies are offered to those whose work was omitted. Many “recipe books” that are provided by instrument companies summarize established methods and regulatory and other international agencies also provide peer reviewed methods. The United States Environmental Protection Agency (US EPA) and the American Society for Testing and Materials (ASTM) web sites serve as examples of electronically compiled analytical methods. The International Organization for Standardization (ISO) also publishes analytical methods (ISO 71.040—Analytical Chemistry) although only a few are currently listed on the ISO web site; more are expected to be added in the years ahead. Recent developments are annually reviewed in Atomic Spectrometry Updates published in the Journal of Analytical Atomic Spectrometry. Atomic Spectrometry Updates covering Environmental Analysis [1 –3] and Advances in Atomic Emission, Absorption, and Fluorescence Spectrometry, and Related Techniques [4,5] are useful sources of information. As well, Analytical Chemistry provides application reviews, in alternate years, devoted to Water Analysis [6,7] and Environmental Analysis [8– 10].

26.1

DIRECT METHODS OF DETERMINATION

Telliard [11] published an overview of the EPA’s Analytical Methods Program and Performance Based Measurement System (PBMS). The US EPA provides a useful series of analytical methods that discuss sample preservation, reagents, Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

857

S.N. Willie

standards preparation, interferences and quality control. Method 200.9 outlines a procedure for the determination of dissolved and total recoverable elements by stabilized temperature platform ET-AAS in ground water, surface water, drinking water, storm runoff, and industrial and domestic waste water. Method 200.7 addresses trace elements in water, solids and biosolids by ICPAES and Method 200.8 is devoted to the determination of trace elements in waters and waste waters by ICP-MS [12]. ASTM D3919 and D4691 are general documents that outline the measurement of trace elements in water by ET-AAS and F-AAS, respectively [13]. AOAC International Ofﬁcial Method 993.14, Trace Elements in Water and Waste Water, outlines an ICP-MS procedure for 20 trace elements at concentrations between 0.8 and 200 mg l21 in ﬁnished drinking water, ground water and waste water [14]. The direct determination of trace analytes in natural waters is the most desirable approach as it involves minimal sample handling and pretreatment, thereby minimizing the risk of contamination. Direct determinations in nonsaline waters are relatively straightforward, provided the analyte concentrations are above instrumental detection limits and the matrix does not contain interfering components at sufﬁcient concentrations to cause severe problems. Sansoni [15] compared ICP-MS with ICP-OES, ET-AAS and ICP-AFS for the multielement analysis of ground and drinking water from a granitic region. More recently, a comparison of methods using ICP-MS and ICP-OES for 13 trace elements in municipal waters was performed by MillerIhli and Baker [16]. At concentrations above the quantitation limit, good agreement between methods was obtained; obviously the superior sensitivity of ICP-MS was able to provide results for several elements that were below the detection capability of ICP-OES. The direct determination of rare earth elements (REEs) in fresh water has also been reported. Halicz et al. [17] achieved detection limits between 0.005 and 0.05 ng l21 using ultrasonic nebulization (USN) sample introduction with ICP-MS detection. Problems encountered from the presence of total dissolved salts up to 0.05% were overcome with the use of added internal standards. The introduction of undiluted sea water into an ICP-MS can cause clogging of the nebulizer and cones, create space-charge effects and introduce signiﬁcant spectral interferences. However, for a select group of elements in sea water present at suitable concentrations, simple dilution can be performed. Roslund and Lund [18] introduced a small volume of the sea water CRM NASS-4 into an acidiﬁed water carrier stream using a ﬂow injection (FI) system. As a result of the transient response, the number of elements that could be determined using a quadrupole based ICP-MS was limited. Sector ﬁeld ICP-MS (SF-ICP-MS) spectrometers have the potential to revolutionize trace element determinations in sea water matrices as most spectral interferences arising from molecular ion interferences due to the presence of major elements such as chlorine, sodium, calcium, etc., can be eliminated. Unfortunately, the high cost of these instruments precludes their widespread availability. Microconcentric nebulization

858

The determination of trace elements in water

with aerosol desolvation combined with SF-ICP-MS was used by Field et al. [19] to determine V, Cr, Mn, Fe, Co, Ni, Cu, Zn, Cd and Pb in diluted sea water. Sample volumes of 50 ml were used and calibration was accomplished with a matrix-matched external calibration curve. Sources of contamination were discussed, which were derived primarily from the ICP-MS introduction system. A semiquantitative method for the determination of trace metals in water samples of low and high salinity was reported by Bayon et al. [20] using SFICP-MS. The use of both low and medium resolution was employed for a low salinity sample. Matrix effects were also evaluated using spiked sea water samples following a ﬁvefold dilution. Reaction or collision cell technology is a relatively new development that offers signiﬁcant potential for overcoming some traditional problems that currently limit quadrupole based ICP-MS. An interesting paper by Louie et al. [21] compared SF-ICP-MS with a dynamic reaction cell (DRC) ICP-MS and a traditional quadrupole ICP-MS instrument for the determination of trace elements in sea water. Results showed that DRC-ICP-MS with an ammonia reaction gas could be used for the direct determination of 51V, 52Cr and 55Mn and molecular interferences were reduced signiﬁcantly for 58Ni and 63Cu, especially if a matrix-matched blank was used. There are a number of papers that report on the direct determination of trace elements in sea water by ET-AAS. In some cases, these methods are suitable only for screening purposes or analysis of contaminated samples as the number of elements that can be determined directly in pristine water is limited. Nickel [22], Ag [23], As [24,25], Bi, In and Pb [26], Zn [27], Al [28], Se [29], Cd [30] and Pb[30], Cu and Mn [31], Cr, Cu, Mn [32] provide a few examples. Chemical modiﬁers are usually used to extend the thermal pretreatment temperature to release the volatile decomposition products and evaporate the maximum amount of matrix prior to atomization of the analyte. An alternative approach is the use of a releasing agent such as EDTA or citric acid for the selective volatilization of the analyte. For example, Guevremont has reported the direct determination of Cd [33] and Zn [34] in sea water with this novel approach. The coupling of electrothermal vaporization (ETV) sample introduction to an ICP-MS provides a number of signiﬁcant advantages, including high sensitivity, the use of chemical modiﬁers and appropriate volatilization temperatures to reduce or eliminate matrix interferences and the ability to process microliter volumes. Methods have been reported by: Gre´goire and Ballinas for As [35], Chapple and Byrne for Co, Cu, Mn, Ni, V and Hg [36], Chang and Jiang for Bi [37], and Byrne and Chapple for As, Pb, Sb, Sn and Tl [38]. Problems associated with analyte transport can be overcome using an isotope dilution (ID) calibration method [39]. Simple optical absorption spectrophotometric techniques for the determination of trace elements can be simple and selective but generally suffer from a lack of sensitivity. Recently, use of various long path length cells has overcome

859

S.N. Willie

this limitation. Zhang et al. [40] adapted a long liquid waveguide capillary ﬂow cell to a gas-segmented continuous ﬂow auto-analyzer for the sub nM (few ng l21) determination of Fe. Yao and Byrne [41] used a 5 m liquid core waveguide to extend the sensitivity of conventional absorbance spectroscopy for Cr and Mo. With both these methods no preconcentration was required. Table 26.1 summarizes a number of procedures that report the direct determination of trace elements, i.e., without analyte concentration or matrix separation. Methods using hydride generation (HG) or derivatization of the analyte to a volatile form could also be considered direct, but will be addressed separately in this chapter.

26.2

PRECONCENTRATION TECHNIQUES—MULTIELEMENT

The proliferation of plasma source mass spectrometry has revolutionized the multielement determination of trace elements. However, for complex samples such as sea water, some form of matrix separation and/or analyte preconcentration is often required and is the focus of this section. Various procedures using solvent extraction, coprecipitation and chelating resins have been utilized. It is evident that metal chelating resins and immobilized (adsorbed or bonded) chelating agents are the most popular for these applications, as methods based on their use are easy to automate and incorporate into FI systems. Chelating ion exchange resins based on an iminodiacetate (IDA) functionality to separate transition metals from alkaline earth ions have been widely studied since Riley and Taylor [52] ﬁrst reported a batch preconcentration procedure using Chelex 100 followed by atomic absorption detection. Alternative IDA products include MetPac CC-1 (Dionex, Sunneyvale, CA, USA) [53–56], Chelite-C (Serva, Heidelberg, Germany) [57], Muromac A-1 (Muromachi Chemical Co., Tokyo, Japan) [56,58], Prosep IDA (Bioprocessing, Consett, Durham, UK) [59] and Toyopearl AF-Chelate 650M (Tosohaas, Montgomeryville, PA, USA) [60 – 63]. Although all have the same IDA functionality, analytical performance amongst these materials varies slightly. Chelex 100, one of the original IDA chelating resins, swells and shrinks with changes in sample pH [59], which can be problematic when packed into a minicolumn. IDA resins based on more rigid supports are now popular: CheliteC was used by Taylor et al. [57] to determine Mn, Ni, Co, Cu, Zn, Cd, U and Pb in sea water using a low pressure manifold coupled to ICP-MS. Calibration was performed by standard additions and each run took 19 min to complete. The procedure required a wash of the column with 8 ml of buffer following sample loading. Ebdon et al. [55] reported that a buffer wash was also required to remove the matrix from a MetPac CC-1 column for the determination of Cu, Mo, Ni, U and Zn in sea water and brines. Similar results were reported by Bloxham et al. [53] using a MetPac CC-1 column.

860

The determination of trace elements in water TABLE 26.1 Direct determination Element

Instrumentation

Comments

Reference

36 elements

ICP-MS, ICP-OES, ICP-AFS, AAS ICP-OES, ICP-MS

Of the 36 elements attempted in ground water, 34 could be determined by ICP-MS, 14 by AAS, 13 by ICP-OES and 12 by ICP-AFS Results of a collaborative project to obtain trace element composition data for municipal waters sampled throughout the United States Spring water analyzed for REE concentrations between 0.02 and 13 ng l21. USN used to obtain sufﬁcient sensitivity Direct determination of trace metals in sea water by ICP-MS using FI and ETV. FI-ICP-MS was used for the determination of Mn, Mo and U in diluted sea water. ETV-ICP-MS was used for the determination of Cd and Pb Direct determination of trace metals in sea water using microconcentric nebulization with aerosol desolvation combined with SF-ICP-MS. Calibration using matrixmatched external calibration curve Semiquantitative method for trace metals in low and high salinity water The direct determination of several transition elements in sea water by reaction cell ICP-MS. Results show signiﬁcant improvement over a standard quadrupole instrument Determination of 15 trace metals in sea water. Discrepancies for Cd and Zn were explained by an unresolved MoO interference (Cd) and insufﬁcient internal standard correction (Zn). Present data indicate that SF-ICP-MS is applicable to the rapid and accurate multielement determination of trace metals at pg ml21 levels in saline water virtually free from spectral interferences

[15]

Ca, Cu, Fe, Mg, P, K, Na, Mn, Zn, Co, Cr, Ni and V REEs

ICP-MS

Mn, Mo, U, Cd and Pb

ICP-MS

V, Cr, Mn, Fe, Co, Ni, Cu, Zn, Cd and Pb

SF-ICP-MS

22 elements

SF-ICP-MS

Cu, Cr, Mn, Ni and V

ICP-RC-MS

15 elements

SF-ICP-MS

[16]

[17]

[18]

[19]

[20] [21]

[42]

continued

861

S.N. Willie

TABLE 26.1 (continuation) Element

Instrumentation

Comments

Reference

17 elements

ICP-MS

[43]

226

Ra, 230Th, Th, 233U, 237 Np, 238U and 241Am Ti, V, Cr, Mn, Fe, Co, Cu, Zn, Mo, Ag, Cd, Sb, Ba, Au, Pb, Bi and U

SF-ICP-MS

230

SF-ICP-MS

The determination of 17 trace metals in ultrapure water at 10 ng l21 and lower. A quick evaluation scheme is also presented for routine ultrapure water quality checks SF-ICP-MS with direct injection high efﬁciency nebulizer (DIHEN) used for analysis of radioactive environmental waste solutions Direct, simultaneous determination of trace elements in alpine snow and old ice at the ng g21 and pg g21 level. One milliliter samples were introduced using a microconcentric nebulizer. To avoid possible spectral interferences, measurements were made under both medium and low resolution modes Rapid determination of 230Th and 231 Pa in sea water using micro-nebulization SF-ICP-MS coupled with desolvation Chemical modiﬁcation and background correction systems were studied for the direct determination of several high and medium volatility and refractory metals in sea water. The inﬂuence of injection volume with different chemical modiﬁers was evaluated Methods for the direct determination of Cu and Cd in sea water were described using ET-AAS with chemical modiﬁcation. The matrix interference was completely removed so that a simple aqueous calibration curve method could be used for calibration Evaluation of releasing agents (citric acid and EDTA) to lower the atomization temperature of the analyte, permitting temporal separation of the analyte and matrix Direct determination of trace metals in sea water using multielement ET-AAS. Matrix interferences were completely removed, permitting calibration using a simple aqueous curve

232

Th and Pa

SF-ICP-MS

231

Ag, As, Cd, Cr, Hg, Ni and Pb

ET-AAS

Cu, Cd

ET-AAS

Cd, Zn

ET-AAS

Cu, Mn and Mo

ET-AAS

[44]

[45]

[46]

[47]

[48]

[33,34]

[49]

continued

862

The determination of trace elements in water

TABLE 26.1 (continuation) Element

Instrumentation

Comments

Reference

Cu, Mn

ET-AAS

[50]

Cd, Hg and Pb

ETV-ICPMS

Cd

Atom trapping AAS Spectro photometric

Direct and simultaneous determinations of Cu and Mn in sea water using ET-AAS and chemical modiﬁcation. Matrix interferences were removed completely so that a simple calibration curve method could be used Calibration by ID applied to the determination of Cd, Hg and Pb in sea water samples. Various modiﬁers were tested Direct determination of Cd in waters using atom trapping A liquid core waveguide was used to extend the sensitivity of conventional absorbance spectroscopy for Cr and Mo achieving 0.01 and 0.06 ng l21 detection limits, respectively A long liquid waveguide capillary ﬂow cell was ﬁtted to a gas-segmented continuous ﬂow auto-analyzer for trace analysis of Fe in water. LODs of 0.1 ng were achieved using 2 ml sample volumes

Cr, Mo

Fe

Spectro photometric

[39]

[51] [41]

[40]

PROSEP Chelating-1 IDA resin has been utilized on-line in conjunction with ICP-MS for the determination of Co, Ni, Cu, Zn and Cd in sea water [59,64]. Quantitation was performed using a simple calibration curve and the analysis time per run was 5 min. LOD calculated from the calibration curve varied between 0.02 ng ml21 for Co and 0.20 ng ml21 for Zn. Willie et al. [60] reported an automated low-pressure FI method for the determination of Cu, Ni, Zn, Mn, Co, Pb, Cd and V in sea water based on quadrupole ICP-MS detection. A column of Toyopearl AF-Chelate 650M was used to sequester the trace elements from a sample of sea water buffered on-line to a pH of 5.2. Following washing of the column with deionized water, the metals were subsequently eluted into the plasma with 1 M HNO3. LODs were less than 10 pg ml21 with a sample loading time of 60 s. The time required to process one sample was less than 4 min. Quantitation was accomplished by means of an aqueous calibration curve. Several ensuing papers by Warnken [62], Willie [61] and Beck [65] have used this same resin and reported subtle improvements in performance. The later used SF-ICP-MS to identify a number of complications due to interferences. Many subsequent studies have replaced HPLC systems with low-pressure columns,

863

S.N. Willie

simplifying operation as well as decreasing analysis time. Other notable improvements include, rinsing the residual salt from the column with water instead of a buffer and calibration using a simple aqueous standards as opposed to the method of standard additions. Together, these advances decrease analysis time, reduce the blank and simplify the overall procedure. An alternative method for the detection of trace metals in saline samples by ICP-MS utilizes batch preconcentration with 0.2 mm polymeric beads having IDA chelating groups [66]. The beads are subsequently nebulized directly into the plasma. Enrichment factors between 40 and 48 can be obtained due to the concentration of a 120 ml suspension to 2.5 –3.0 ml eluate. The method was applied to the determination of Mn, Fe, Ni, Co, Cu, Zn, Cd, and Pb and several REEs in sea water. A battery powered ﬁeld sampling unit has been described for the in situ preconcentration of trace elements from natural waters by Nickson et al. [67] Water samples were ﬁltered on-line and passed through twin MetPac CC-1 microcolumns. This allowed duplicate samples to be preconcentrated in parallel. The microcolumns were then taken to the laboratory and placed in a FI manifold coupled to an ICP-AES for elution and sample quantitation. The afﬁnity of transition metals for 8-hydroxyquinoline while rejecting the alkaline earths has made this chelating agent a popular choice for the preconcentration of trace elements from sea water. As a result, it has been successfully immobilized onto various polymeric and inorganic supports for use in ion-exchange or chromotographic applications. Rao and Gladis [68] have summarized the use of 8-hyroxyquinoline for this purpose. Initially used in an off-line mode [69], and requiring 500 ml of sample, the method was later adapted to an on-line procedure using ICP-MS coupled to a commercial HPLC system [54]. Procedures describing 8-hydroxyquinoline immobilized onto controlled pore glass [59] and silicone tubing [70] have also been reported for the determination of Mn, Co, Ni, Cu, Zn and Cd in sea water. A disadvantage of coupling the 8-hydroxyquinoline resin systems to ICPMS for detection is the need for both HNO3 and HCl to quantitatively elute the retained trace metals from the column. The resulting chlorine containing polyatomic ions preclude the determination of vanadium, and overlap a number of minor isotopes of other elements. An additional, and perhaps more signiﬁcant disadvantage of the use of 8-hydroxyquinoline resins, is the lack of a commercial supplier, thus requiring in-house synthesis of the material. In an attempt to surmount this difﬁculty, Dierssen et al. [71] have reported several simpliﬁed synthesis procedures along with performance characteristics for 8-hydroxyquinoline covalently bonded to a series of vinyl polymer resins. Points to consider when applying these procedures include the commercial availability of appropriate columns. In particular, the selection of metal-free low-pressure columns and support frits is limited. The availability of reagents to adjust sample pH is less restricted. The operating pH for IDA resins is

864

The determination of trace elements in water

usually chosen to be 5 or 8, enabling the use of ultrapure ammonium hydroxide, acetic acid and hydrochloric acid to prepare appropriate buffers. With the exception of a few elements (Cu, Fe), 8-hydroxyquinoline columns operate most efﬁciently at pH slightly .8 and an ammonium chloride buffer can be used. Three atomic spectrometry techniques (SF-ICP-MS, ET-AAS and HG-AAS) were evaluated for the analysis of water by Townsend et al. [72]. A dithiocarbamate-chelation/back-extraction technique was used to separate and concentrate Co, Ni, Cu, Zn, Cd and Pb from eight water samples prior to analysis by SF-ICP-MS and ET-AAS. A number of other elements in the waters were analyzed directly (Mn, Fe and Zn by ET-AAS; As by HG-AFS), or following sample dilution (V, Mn, Fe, As, Mo, Ba and U by SF-ICP-MS). A combination of all three analytical techniques was necessary for the successful analysis of the elements considered in this study. Table 26.2 summarizes a number of publications that report preconcentration procedures for the multielement determination of trace elements in water.

26.3

PRECONCENTRATION—INDIVIDUAL ELEMENTS

Table 26.3 summarizes methods for the determination of selected individual elements. Several analytes are noteworthy. In January 2001, the Federal Register published EPA’s ﬁnal arsenic regulation, setting a new protective drinking water standard for arsenic of 10 mg l21 with an effective date in early 2001 and a compliance date in 2006 [118]. Arsenic is presented in Table 26.3 using direct ET-AAS and in Table 26.4 with other hydride forming elements. There has been considerable interest in the determination of sub nM (,10 ng l21) Fe by oceanographers studying the biogeochemical dynamics of Fe in sea water [119]. Several speciﬁc reviews are available, D’Ulivo has published a review of Se and Te in environmental samples [120].

26.4

DETERMINATION OF TRACE ELEMENTS AS VOLATILE SPECIES

The production of volatile hydrides by reaction with sodium tetrahydroborate prior to determination by AAS, ICP-OES and ICP-MS is a well-established analytical technique [152,153]. Separation of the analyte from the matrix in this manner can provide beneﬁts such as the removal of the Cl species for As determination by ICP-MS. Although high mass resolution ICP-MS permits the spectral separation of the argon chloride interference, the accompanying reduction in sensitivity at high resolution compromise detection and determination limits [154]. Collection and concentration of the hydride prior to its introduction into the atomization cell allows improved detection limits over direct introduction into

865

S.N. Willie TABLE 26.2 Multielement preconcentration Elements

Matrix

Detection

Comments

Reference

Mn, Co, Cu, Zn and Pb Fe, Mn,Co, Ni, Cu, Zn, Cd and Pb

Sea water

ICP-MS

[53]

Sea water

ICP-MS

Cu, Mo, Ni, U, Zn and In

Brines

ICP-MS

Cd, Co, Cu, Mn, Ni, Pb, U and Zn

Sea water

ICP-MS

Cu, Mo

Sea water

ET-AAS

Multielement

Estuarine and sea water Sea water

ICP-MS

FI method using a column of MetPac CC-1 resin. Three sea water CRM’s were analyzed Comparison of an IDA (MetPac CC-1) and 8-hydroxyquinoline column using a commercial chelation concentration system On-line procedure comparing two types of IDA resin columns. MetPac CC-1 and Xylenol Orange columns were used for the isolation of alkaline earth and transition metals from NaCl An IDA column (Chelite C) in a commercially available low-pressure FI system was used Automatic on-line preconcentration system using a Muromac A-1 IDA column On-line preconcentration using MetPac-CC1 resin

ICP-MS

A comparison of Chelex-100 and Metpac CC-1 IDA resins

[74]

Sea water

ICP-MS

A low-pressure FI procedure using Muromac A-1 IDA resin

[56]

Sea water

ICP-MS

On-line preconcentration using PROSEP IDA, IDA immobilized on controlled pore glass. Calibrations prepared from both pure water and artiﬁcial sea water matrices were found to be comparable

[59]

Al, As, Co, Cu, Mn, Mo, Ni, Pb and V Al, V, Mn, Co, Ni, Cu, Zn, Mo, Cd, Pb and U Mn, Co, Ni, Cu, Zn and Cd

[54]

[55]

[57]

[58]

[73]

continued

866

The determination of trace elements in water

TABLE 26.2 (continuation) Elements

Matrix

Detection

Comments

Reference

Cu, Zn, Cd, Mn and Ni

Sea water

F-AAS

[75]

Co, Cu, Cd, Mn, Zn, Ni and Pb

Sea water

ICP-MS

Cu, Ni, Zn, Mn, Co, Pb, Cd and V

Sea water

ICP-MS

Cu, Ni, Zn, Co, Pb, Cd and Fe

Sea water

ICP-MS

Mn, Cu, Ni, Cd and Pb

Sea water

ICP-MS

18 elements and REEs

Stream and lake water Sea water

ICP-MS

FI procedure comparing controlled pore glass based IDA resin and immobilized 8-hydroxyquinoline A column of 8-hydroxyquinoline chelating reagent immobilized onto a controlled pore glass support was used Calibrations prepared from both pure water and artiﬁcial sea water matrices were found to be comparable Automated on-line preconcentration using Toyopearl AF-Chelate 650M IDA resin. Calibration by aqueous standards On-line preconcentration using Toyopearl AF-Chelate 650M IDA resin. Calibration by ID and method of additions On-line preconcentration using Toyopearl AF-Chelate 650M IDA resin. Elution acid concentration and other conditions optimized. One milliliter of sample required for LOD , 1 pg ml21 A comparison of chelation with MetPac CC-1 IDA resin against direct analysis On-line preconcentration using Toyopearl AF-Chelate 650M IDA resin. Calibration with aqueous standards Batch preconcentration with direct injection of the analyte loaded on IDA chelating resin into the plasma

Cu, Ni, Zn, Co, Pb, Cd and REEs Mn, Fe,Ni, Co,Cu, Zn, Cd and Pb, La, Ce, Eu, Gd, Yb, Lu

Sea water

ICP-TOFMS

ICP-MS

[76]

[60]

[61]

[62]

[77]

[63]

[66]

continued

867

S.N. Willie

TABLE 26.2 (continuation) Elements

Matrix

Detection

Comments

Reference

Cd, Cu, Ni, Zn and Mn

Estuarine water

SF-ICP-MS

[65]

Cu, Ni, Cd, Co, Mn, V and Pb Cu, Ni, Zn, Mn, Co, Pb and Cd Mn, Fe, Zn, Cd, Cu, Ni and Co

Pond and sea water

F-AAS, ICP-OES

Sea water

ICP-MS

Sea water

ET-AAS

V, Ni, Mn, Cu, Zn, Cd, Pb and U

Sea water

Quadrupole and SF-ICPMS

Cd, Cu, Pb, Zn, Sc, La, Ce, Nd and Yb

Sea water

ICP-MS

Cd, Co, Cu, Ni, Pb, U, Y and the 14 REEs Cd, Co, Cu, Fe, Mn, Ni, Pb, Tl, U and Zn Pb, Cu, Cd and Mn

Sea water

ICP-MS

Sea water

ICP-MS

A FI method that uses 3 ml sample and employs an automated on-line preconcentration step using Toyopearl AF Chelate 650M IDA resin On-line procedure using an iminodiacetic acid membrane Immobilization of 8-hydroxyquinoline onto silicone tubing Comparison of ﬁve procedures for covalently bonding 8-hydroxyquinoline to a vinyl polymer resin Comparison of 8-hydroxyquinoline based chelating resin using quadrupole ICP-MS and direct determination using SF-ICP-MS Comparison of the performance of Chelex-100 and Chelamine. Chelexwas preferred for REE’s. Sixty milliliters of sample was required Off-line method using cellulose-immobilized ethylenediaminetriacetic acid Analyte complexation with Na –DDTC on a biphenyl column

Drinking water

IC-spectrophotometry

MetPac CC-1, a sulfonated cation exchanger (TMC-1) and a bifunctional ionexchange column (CS5A) were used

[78]

[70]

[71]

[79]

[80]

[81]

[82]

[83]

continued

868

The determination of trace elements in water

TABLE 26.2 (continuation) Elements

Matrix

Detection

Comments

Reference

River and spring water

Spectrophotometry

[84]

Pb, Cu, Cd, Co, Zn and Ni

Ground water

IC-UV–Vis

Co, Cu, Mo, Mn, Fe, Ti and V

Sea and river water

ICP-OES

Cu, Cd and Pb Cu, As, Se, Cd, In, Hg, Tl, Pb and Bi Cr, Cu, Ni and Pb

Sea water

F-AAS

Fresh water

ICP-MS

Sea water

F-AAS

Sea water

ICP-OES

Two column concentration and separation using an IDA chelating stationary phase followed by a mixedbed ion-exchange column for separation. Pyridine-2,6-dicarboxylic acid was used as the complexing agent in the mobile phase. LODs in the tens of ng l21 range were achieved Three column systems based on ionic separation were evaluated: (1) a cationic column, (2) an anionic column, and (3) a bifunctional ion-exchange column Three phase extraction using a combination of diantipyrylmethane (DAM), SCN2 and HCl. The implementation of the third phase extraction permits a preconcentration of trace elements by a factor ranging from 33 to 45 1,10-Phenanthroline complexes adsorbed on C18 On-line separation by retention of the DDTP complexes on a C18 minicolumn Sorption of metal ions on XAD-2000 as 1-(2-pyridylazo) 2-naphtol (PAN) chelates A battery powered ﬁeld sampling unit for the selective in situ preconcentration of trace elements from natural waters on a IDA resin

Cd, Co, Cu, Mn, Ni, Pb and Zn

[85]

[86]

[87] [88]

[89]

[90]

continued

869

S.N. Willie

TABLE 26.2 (continuation) Elements

Matrix

Detection

Comments

Cd, Co, Cu, Fe, Ni, Pb and Zn

Sea water

ICP-MS

V, Co, Ni, Ga, Y, Mo, Cd, Ce, Pr, Nd, Sm, Eu, Gd, Tb, Dy, Ho, Er, Tm, Yb, Lu, W and U Eu, Tb, Ho, Tm and Lu Mn, Ni, Cu, Zn and Pb

Sea water

SF-ICP-MS

[91] Off-line solvent extraction procedure adapted from a dithiocarbamate–diisobutyl ketone solvent extraction system with Hg back-extraction. The single extraction procedure was quantitative and external standards were used 8-hydroxyquinoline immobilized [92] on ﬂuorinated metal alkoxide glass. Concentrations between 0.01 ng l21 and 10 mg l21. LODs were found to be dependent on the level of contamination

Sea water

ICP-MS

Sea water and natural water

ICP-MS

REEs

Sea water

ICP-MS

Co, Ni, Cu, Zn, Ni, Mn, Cd, Sb, Pb and U Be, Bi, Co, Ga, Ag, Pb, Cd, Cu, Mn and In

Sea water

ICP-MS

Sea water

ICP-MS

Analytes complexed with 8-hydroxyquinoline and adsorbed on XAD-7 column On-line preconcentration using Toyopearl TSK-immobilized 8-hydroxyquinoline. Three milliliters of sample was required. Precision ,5% 8-Hydroxyquinoline immobilized on polyacrylonitrile hollow ﬁber membrane was synthesized for the preconcentration of REEs A nitrilotriacetate (NTA)type chelating resin was applied to the separation and enrichment of the analyte metal ions An ion exchange chelating ﬁber with aminophosphonic and dithiocarbamate groups based on polyacrylonitrile was used for the simultaneous preconcentration of trace elements in sea water

Reference

[93]

[94]

[95]

[96]

[97]

continued

870

The determination of trace elements in water

TABLE 26.2 (continuation) Elements

Matrix

Detection

Comments

Reference

Pb, Cu and Cd

Sea water

ID-ICP-MS

[98]

Co, Ni and Cu

Sea water

Multielement ET-AAS

Sea water

ET-AAS

Pb, Cu

Sea water

ET-AAS

Be, Bi, Co, Ga, Ag, Pb, Cd, Cu, Mn and In

Sea water

ICP-MS

Cu, Mn and Ni

Sea water

ET-AAS

Cd, Pb, Cu, Ni, Co and Fe

Sea water

ET-AAS

Al, Co, Cr, Fe, La, Mn, Ni, Ti, V, Zn, Y and Pb

Sea water

ICP-OES

Cu, Pb and Cd

Sea water

AAS

Mg(OH)2 coprecipitation using 1.3 ml of sample to achieve picomolar detection limits Sample loaded on Muromac A-1 IDA column and eluted directly into furnace APDC chelate coprecipitation with Co FI on-line Co–APDC coprecipitation A new ion exchange chelating ﬁber with aminophosphonic and dithiocarbamate groups based on polyacrylonitrile was used for the simultaneous preconcentration of trace elements in sea water On-line sorption preconcentration with APDC or 8-hydroxyquinoline in a knotted reactor (KR). APDC offered better performance characteristics than HQ for the preconcentration of Cu and Ni but could not be employed for Mn On-line preconcentration using APDC on a miniature C18 column on the tip of the autosampler arm Coprecipitation with gallium hydroxide. Spectral interferences from Ga are minimal. High purity Ga is readily available Coprecipitation using Pd. The palladium is reduced by the introduction of hydrogen gas into the sample solution

[99]

[100] [101] [97]

[102]

[103]

[104]

[105]

continued

871

S.N. Willie

TABLE 26.2 (continuation) Elements

Matrix

Detection

Comments

Reference

As, Cd, Cr, Cu, Mn and Ni

Sea water

ET-AAS

[106]

Mn, As, V, Y, Sb, W and U

Sea water

ICP-MS

REEs

Waters

ICP-TOFMS with USN

Cu, Ni, Sb, Co, Ag, Cd, Mo, In and Pb

Waters

ICP-TOFMS with USN

Cr, Mn, Fe, Co, Ni and Cu Co, Ni, Cu, Cd and Pb

Estuarine water

ICP-MS

Sea and river water

ET-AAS

An on-line preconcentration system using ﬁlterless collection of coprecipitated analytes based on tetrahydroborate reductive precipitation of added Fe and Pd. The precipitate is dissolved in a 20 ml volume of mixed acid and transported direct to the graphite furnace. The sensitivity of the graphite furnace technique can be enhanced over 400-fold (for an 11 ml sample volume) compared to a standard 20 ml injection volume Coprecipitation with lanthanum. Recoveries were greater than 85% for most elements, but less than 30% for Cr(VI), Se(VI) and Mo(VI). The reproducibility was 5% On-line preconcentration and separation in a KR. The REE complexes were sorbed on the inner walls of PTFE KR precoated with the chelating reagent 1-phenyl-3-methyl-4-benzoylpyrazol-5-one (PMBP). Detection limits range from 3 to 40 pg l21 On-line sorption/preconcentration in a KR. The analytes were complexed with APDC from acidic solutions and sorbed onto the KR. The detection limits varied from 0.5 ng l21 for Sb to 26 ng l21 for Pb On-line precipitation using Na –DDTC with collection in a KR-ﬁlter system Coprecipitation with magnesium oxinate. The background absorption due to the collector was negligible

[107]

[108]

[109]

[110]

[111]

continued

872

The determination of trace elements in water

TABLE 26.2 (continuation) Elements

Matrix

Detection

Comments

Reference

Pt, Pd, Au and Rh

Fresh water

ETV-ICPMS, ICP-MS

[112]

Pb, Cd

Water

F-AAS

Cu, Cd

Ultra-high purity water

IC-ET-AAS

U, Th

Sea water ICP-MS and ground water with 8 mg ml21 humic and fulvicacids Sea water Spectrophotometric

Preconcentration on activated charcoal. Comparison of two sample introduction systems: ETV and solution nebulization Electrochemical preconcentration using deposition on a glassy carbon electrode in a ﬂowthrough microcell A Dionex HPIC-CG5 ion exchange column was used Large sample volumes (up to 200 ml) were loaded onto the concentrator and eluted in 0.200 ml volumes. Cu detection limits was 1 pg ml21, Cd was 0.02 pg ml21 An on-line solid-phase extraction method using an actinide-speciﬁc extraction resin (TRU-spec)

Cu, Cd and Pb

Co, Fe, Mn and Ni

Nuclear reactor coolant water

Review

A screening method for the detection of pollution by Cd, Cu, Pb and Zn in the range 2–20 mmol l21 using the non-selective reagent PAR Analytical techniques such as ET-AAS, ICP-MS, ICP-AES, INAA and XRF are discussed with special emphasis on the detection limits

[113]

[114]

[115]

[116]

[117]

an atomizer. Use of the graphite furnace to serve as both the hydride trapping medium and atomization cell is now well established for ultratrace determination of the hydride forming elements. The use of sequestering agents such as Pd, Ir, Zr coated on the graphite tube along with appropriate instrument software has permitted full automation of this approach [155]. Multielement determination using HG techniques are limited as prereduction requirements can impede system optimization for the simultaneous determination of As and Se. Consequently, for the determination of total

873

S.N. Willie TABLE 26.3 Individual element determinations Element

Matrix

Detection

Comments

References

Ag

Sea water

Isotope dilution ICP-MS

[121]

Ag

Fresh and photographic waste water Sea water and fresh water

ICP-OES, DRC-ICPMS

On-line method using a minicolumn packed with Dowex 1-X8 anion exchange resin to separate and concentrate Ag Digestion procedure to improve Ag recovery

[123]

Tap water and fresh zwater Sea water and well water

ET-AAS

Automated on-line preconcentration system coupled to ET-AAS. 8-hydroxyquinoline and XAD-2 columns compared Direct ET-AAS method using up to 100 ml injection volumes Collection of molybdoarsenate on activated carbon. The suspension was directly introduced into an ETV. Both As(III) and (V) collected Octadecyl silica cartridge, modiﬁed with quinalizarine as a chelating agent Octadecyl silica cartridge modiﬁed with cyanex 301 On-line preconcentration using APDC complex adsorbed on C18 column A FI system using C18 as a sorbent with 4-(2-pyridylazo) resorcinol (PAR) or 2-(2-pyridylazo)-5-dimethylaminophenol (PADMAP) as chelating agent with methanol as the eluent Liquid chromatographic separation through on-line complexation with 8-hydroxyquinoline sulphonate, using a C18 column. Cd(II) could be detected at concentrations as low as 2 mg l21

Al

As

As

Be

Bi Cd

Sea water, well and tap water Natural water Sea water

ET-AAS

ET-AAS

F-AAS

ET-AAS ET-AAS

Cd

Saline water

ET-AAS

Cd

Water samples

Fluorescence

[122]

[124]

[125]

[126]

[127] [128,129]

[130]

[131]

continued

874

The determination of trace elements in water

TABLE 26.3 (continuation) Element

Matrix

Detection

Comments

References

Cd

Sea water

ET-AAS

[132]

Co

Sea water and tap water

ET-AAS

Cr

Sea water

ICP-MS

Cr

River and sea water

ET-AAS

Cu

Sea water

F-AAS

Cu

F-AAS

Fe

Potable water and sea water Sea water

Dynamically coated column of quaternary ammonium salt on C18 Automated on-line FI sorption preconcentration of metal chelate complexes on the walls of a PTFE KR precoated with a chelating reagent. Ammonium pyrrolidinedithiocarbamate (APDC), 8-hydroxyquinoline, PMBP and 2-nitroso-1-naphthol-4-sulfonic acid (NNA) were compared as chelating reagents. An enhancement factor of 28 and a detection limit of 8.1 ng l21 were achieved On-line column preconcentration using IDA resin, Muromac A-1 was used to concentrate Cr(III). Total chromium was determined after reduction of Cr(VI) with a hydroxylamine solution Coprecipitation with a combination of 8-hydroxyquinoline and Pd. Solid sampling ET-AAS was used On-line complexation using 1-nitroso-2-naphthol on C18 Cu–APDC complex sorbed on PTFE turnings

Fe

Sea water

ID-SF-ICPMS

Fe

Sea water

Spectrophotometric

Review

A critical review of historical and current analytical methods for the determination of iron in sea water is presented and their capabilities evaluated Mg(OH)2 coprecipitation used to determine Fe with a 3 ng l21 detection limit using 14 ml of sample On-line preconcentration with immobilized 8-hydroxyquinoline

[133]

[134]

[135]

[136] [137]

[138]

[139]

[140,141]

continued

875

S.N. Willie

TABLE 26.3 (continuation) Element

Matrix

Detection

Comments

References

Li

Sea water

TIMS

[142]

Pb

Sea water

SF-ICP-MS

Pb

Waste water

F-AAS

Pu

Sea water

ICP-MS

Sb

Sea and fresh water

ET-AAS

Tc

Sea water

U

Sea water

ETV-ICPMS, UN-SFICP-MS ICP-MS

Two-column ion exchange to separate Li from matrix A low blank preconcentration method using Mg(OH)2coprecipitation Preconcentration by coprecipitation with gallium phosphate. Pb ranging from 0.5 to 50 mg was quantitatively collected FI procedure utilizing USN with membrane desolvation. One liter samples were coprecipitated using NdF3, followed by ion exchange to enrich Pu Sb(III) was preconcentrated on activated carbon as the pyrogallol complex. Sb(V) was reduced to Sb(III) with KI and ascorbic acid Comparison of two instrumental techniques

V

Sea and ground water

Spectrophotometry

Zn

Sea water

ET-AAS

Zr

Sea water

ICP-OES

876

Comparison of coprecipitation with iron hydroxide and direct analysis following dilution FI procedure incorporating preconcentration on a Sephadex column and using the catalytic action of vanadium on the oxidation of chromotropic acid by bromate Dynamically coated column of methyltricaprylylammonium chloride on C18 Preconcentration of Zr from 200 l of sea water using MnO2impregnated ﬁbers

[143]

[144]

[145]

[146]

[147]

[148]

[149]

[150]

[151]

The determination of trace elements in water TABLE 26.4 The determination of elements as volatile species Element

Detection

Comments

Reference

As

OES

[163]

As

AFS

As

ICP-MS

As

SF-ICP-MS, HG-ICP-MS

As

ET-AAS

As

ET-AAS

As

ICP-MS

Ge

LEAFS

Pb

ICP-MS

Pb

ET-AAS

Sb

AAS

Sb

AFS

Sb

ET-AAS

Continuous ﬂow system using L -cysteine to prereduce As(V) to As(III) FI method for the determination of As in sea water. Reaction coil of 200 cm required to ensure reduction of As(V) A tubular silicone rubber membrane was evaluated as a gas– liquid separator for the determination of As. KI plus ascorbic acid treatment produced a similar response for As(III) and As(V) independent of the matrix Comparison of standard liquid sample introduction in the high-resolution mode and HG in the low-resolution mode Batch HG, precollection of the hydride on a Pd coated pyrolytic platform cuvette. Tap, ground and sea water were analyzed Automated FI– HG with trapping of arsine on an Ir coated graphite tube Comparison between chemical (NaBH4) and electrochemical processes for HG Laser-excited AFS using graphite furnace for sample atomization. Two excitation– detection schemes were evaluated Oxalic acid, ammonium cerium(IV) nitrate and sodium tetrahydroborate were used as the reaction matrix. LOD of 7 ng l21 Different permanent coated graphite tubes: Ir, W and Zr were investigated for the in situ preconcentration of lead hydride. The results obtained were compared with conventional HG-AAS Batch system with cryogenic preconcentration and quartz tube atomization Generation at 708C, additional hydrogen ﬂow used to sustain the atomization ﬂame A continuous ﬂow electrochemical HG system was developed for the determination of total Sb in river water and sea water. Both Sb(III) and Sb(V) were equally converted into their hydrides by electrochemical means. The hydride was trapped in a Pd-coated graphite furnace prior to atomization

[164]

[165]

[154]

[166]

[155] [167] [168]

[169]

[170]

[171] [172] [173]

continued

877

S.N. Willie

TABLE 26.4 (continuation) Element

Detection

Comments

Reference

Se

AFS

[174]

Se

AFS, AAS

Tl

ID-ICP-MS

Se(IV), As, Sb and Ge As, Sb and Se

ICP-MS

As, Sb, Bi and Hg

ICP-MS

As, Sb

AFS

As, Se

AFS

As

ICP-MS

FI–HG with in situ collection of SeH2 on a tungsten ﬁlament coated with rhodium. Applied to Se in mineral water. Reduction of Se(VI) by heating with HCl A device for trapping the SeH2 on a heated gold wire was designed. The method was applied to the determination of selenium in mineral water FI procedure for the determination of Tl in sea water. Te was used as a catalyst for Tl hydride generation Automated continuous ﬂow HG system. L -cysteine was used for prereduction of analyte species The method involves an off-line prereduction procedure for the reduction of Se(VI) to Se(IV) by HCl, combined with an on-line reduction of As(V) and Sb(V) to the trivalent state with thiourea and generation of the hydrides FI system incorporating an in situ nebulizer/hydride generator. The elements in the sample were reduced to the lower oxidation states with L -cysteine before being injected into the HG system Continuous HG for As determination. Determination of the total concentration of the analytes was obtained following reduction with KI FI–HG using membrane between hydride generator and detector to selectively remove water vapor. Surface and ground water samples analysed Direct coupling of solid phase microextraction (SPME) with ICP-MS was used for the non-selective determination of arsenic, selenium, antimony and tin species amenable to HG. Results for As were reported

878

ICP-MS

[175]

[176]

[177]

[178]

[179]

[180]

[181]

[182]

The determination of trace elements in water

selenium, Se(VI) and organic Se must be converted to Se(IV) prior to reduction. To overcome these drawbacks, ﬂow systems for the simultaneous HG of Se and As ﬁrst use the addition of NaBH4 followed by merging with a stream of potassium iodide for the reduction of As(V). L -cysteine has found widespread use as a masking agent and to reduce inorganic Sb and As to hydride compatible oxidation states [156,157]. The redox properties of HBr/ HBrO3 have also been used to reduce organoselenium and Se(VI) to Se(IV) [158]. Volatiles species generation is not only limited to the traditional hydride forming elements using sodium tetrahydroborate, but carbon monoxide and sodium tetraethylborate can also be used to form volatile inorganic species. Elemental Ni reacts with CO at room temperature to form the volatile Ni(CO)4. This has been used for the determination of Ni in water by ET-AAS [159] and ICP-MS [160]. The reaction of sodium tetraethylborate with Pb has been used for the determination of Pb in sea water [161]. An expansion in the suite of elements amenable to volatile vapor generation appears possible, but applications remain to be explored [162]. 26.5

MERCURY

The well-known toxic effects of Hg and its compounds justify the concern for the presence of this element in the environment. This section primarily deals with the determination of inorganic mercury; the speciation of Hg is the subject of Chapter 33. A useful paper by Ferrari et al. [183] discusses methods and the laboratory facilities used for the determination of ultra-low levels of Hg. This particular laboratory is pressurized with air ﬁltered through high efﬁciency particle ﬁlters (HEPA) and experiments are conducted on a clean bench especially constructed with both particle and activated charcoal ﬁlters. Ultrapure water with a Hg content of 0.08 ^ 0.02 pg g21 was produced for dilution of standards and cleaning of plastic containers. The determination and control of blanks is also discussed. The EPA has released a performance based method for the determination of low-level Hg, entitled Method 1631, Revision E, Mercury in Water by Oxidation, Purge and Trap, and Cold Vapor Atomic Fluorescence Spectrometry [184]. The determination of total Hg and dissolved Hg is discussed using dual gold traps in either a batch or FI system. This method supports water quality monitoring programs authorized under the Clean Water Act. In some cases, ambient water quality criteria are much lower than levels measurable using previously approved EPA methods. Method 1631E contains practical information for the analyst by addressing many factors necessary to achieve data of a relevant quality, including sample collection, preservation, storage, contamination and measurement interferences. As well, the apparatus and related analytical procedures are

879

S.N. Willie

described in the document to achieve the method detection limit of 0.2 ng l21 and a minimum level of quantitation of 0.5 ng21. There are very few reference materials for Hg in water. Of note is the BCR CRM 579, Hg in sea water with a certiﬁed value of 1.85 ^ 0.20 ng kg21. The preparation of this material and the analytical methods used to accurately achieve this value were described by Kramer et al. [185]. A review of analytical methods for the determination of inorganic Hg and CH3Hg was prepared by Puk and Weber [186]. Topics described for each category include sample treatment, separation, detection and limit of detection. Harrington [187] has prepared a review dealing primarily with the speciation of Hg using HPLC. Table 26.5 summarizes a variety of methods used for the determination of Hg in aqueous samples.

26.6

LUMINESCENCE

Luminescence is the general term used to describe the emission of light which occurs when a molecule in an excited state relaxes to its ground state. The source of energy used to obtain the excited state deﬁnes the type of luminescence. Energy supplied by electromagnetic radiation (ﬂuorescence or phosphorescence) or chemical reaction (chemiluminescence) is commonly used in analytical instrumentation. Luminometers based on light detection by photomultiplier tubes are among the simplest, most robust and least expensive devices available and have found widespread use, not only in the laboratory, but are well suited for use on board ships [223,224]. Chemiluminescence (CL) has a distinct advantage since a chemical reaction produces the excitation, eliminating problems frequently encountered in ﬂuorescence, such as light scattering or source instability. A review of the analytical applications combining FI with CL detection emphasizes the wide range of applications of the technique [225]. However, the elements that can be determined are obviously restricted due to the limited number of reagents that satisfy the chemical requirements. Table 26.6 summarizes a number of sensitive CL methods that have been developed involving reactions with luminol, brilliant sulfoﬂavin, 1,10-phenanthroline and gallic acid. Generally, separation of an interfering species is required prior to the determination of the analyte. This has been accomplished using masking agents or columns of ion exchange/chelating resins. The latter can also be used to preconcentrate the analyte of interest. A summary of the use of immobilized 8-hyroxyquinoline and it derivatives for this purpose has been presented by Rao and Gladis [68].

880

The determination of trace elements in water TABLE 26.5 The determination of mercury Analyte

Detection

Comments

Reference

Hg

CV-AAS

[183]

Hg

CV-AAS

Hg

CV-AAS

Hg

CV-AAS, CV-ICP-OES

Hg

CV-AFS

Hg

FI-AFS

Hg

CV-ICP-MS

Hg

CV-ICP-MS

Hg

CV-AAS

Determination of Hg in ice, snow and ultrapure water Sampling and methodologies are described to produce reliable results at picomolar (sub ppb) levels in natural waters Two-stage amalgamation on Au for the determination of Hg in sea water Use of micelles and vesicles to improve Hg generation kinetics and a membrane drier tube to eliminate water Comparison of single vs. dual stage amalgamation on Au On-line bromide– bromate oxidation to convert organic mercury to inorganic Hg in ﬁltered sea water samples Discussion of the collection, ﬁltration and preservation of surface water samples for the subsequent determination of total Hg by CV-ICP-MS Hg in water samples determined by amalgamation on a gold– platinum gauze prior to ICP-MS determination A sequential injection method using one pump and one valve. A LOD of 0.46 mg l21 was achieved

[188]

[189]

[190]

[191]

[192]

[193]

[194]

[195]

continued

881

S.N. Willie

TABLE 26.5 (continuation) Analyte

Detection

Comments

Reference

Hg

FI-CV-AAS

[196]

Hg

FI-CV-AAS

Hg

CVﬂuorescence

Hg

CV-AAS, ICP-OES

Hg

FI-AAS

Hg

ET-AAS

Optimization of a FI-CV-AAS system coupled with gold amalgam preconcentration and on-line UV digestion. The apparatus is designed for use in the ﬁeld Hg was retained as the Hg(II)– (5-Br-PADAP) complex on a KR. The LOD for the preconcentration of 25 ml was 5 ng l21 in drinking water samples A spectroﬂuorometric method for the determination of total Hg in waste waters based on ﬂuorescence quenching of rhodamine B with Hg(II) in the presence of iodide Comparison of CV-AAS and ICP-AES using a similar amalgamation preconcentration procedure Comparison of three solid chelating reagents for the preconcentration of Pb and Hg from sea water. The evaluated materials were 7-(4-ethyl-1-methyloctyl)8-hydroxiquinoline (Kelex 100) adsorbed on C18, 8-hydroxyquinoline immobilized on vinyl a co-polymer and Chelex-100 A comparative study of different chemical modiﬁers for ET-AAS

[197]

[198]

[199]

[200]

[201]

continued

882

The determination of trace elements in water

TABLE 26.5 (continuation) Analyte

Detection

Comments

Reference

Hg

ET-AAS

[202]

Hg and Bi

ETV-ICP-MS

Hg

ID-CV-ICP-MS

Hg

CV-ICP-OES

Hg

CV-AAS

Hg

CV-AAS

Hg

CV-ICP-MS

CV generation with trapping and atomization in ET-AAS by selective reduction with NaBH4 and SnCl2. Ir, W and Zr coated graphite tubes were investigated for the preconcentration of the mercury vapor Vapor generation ETV-ICPMS for the determination of Hg and Bi in water. Pt coated graphite tubes were used An isotope dilution CV-ICPMS method applied to the certiﬁcation of mercury in various NIST standard reference materials Cloud point extraction was employed for the preconcentration of mercury in tap water samples. The mercury was extracted as Hg(II)– (5-Br-PADAP) complex Pretreatment of water samples using UV irradiation-peroxodisulfate for the determination of total Hg Pretreatment of water samples using ozone for the determination of total Hg in geothermal water samples Hg vapor was collected in a glass chamber prior to injection into the plasma, time resolved acquisition was used for signal measurement

[203]

[204]

[205]

[206]

[207]

[208]

continued

883

S.N. Willie

TABLE 26.5 (continuation) Analyte

Detection

Comments

Reference

Hg

ICP-MS

[209]

Hg

ICP-MS

Hg

Fluorescence

Hg and CH3Hg

LC –ICP-MS

On-line separation and preconcentration by retention of the Hg complex with the ammonium salt of O,Odiethyl dithiophosphoric acid on C18 Electrochemical generation of mercury from a chlor-alkali plant and lagoon waters The determination of Hg(II) in spiked samples of mineral, tap and sea water using ﬂuorescence optosensing Field sampling by passing river water through C18 column modiﬁed with sodium diethyldithiocarbamate 2-Mercaptobenzothiazole resin was used for the separation and preconcentration of inorganic and alkylmercury from natural waters Inorganic Hg was retained as the anionic complex with methylthymol blue on a column of Dowex IX-8 resin. Organomercurial species were oxidized permitted the determination of total mercury. The difference between total and inorganic mercury determined the organomercury content Coupling of pervaporation and AAS applied to Hg speciation in a sample of waste water Large volume injection using a packed GC column serially connected to capillary GC The determination of inorganic and organic mercury species in lake water

Inorganic and CV-AAS alkylmercury species Inorganic Hg and organomercury

FI-CV-AAS

Inorganic Hg and organomercury Hg and CH3Hg

CV-AAS

Hg and CH3Hg

FI-CV-AAS

GC –MIP-OES

[210]

[211]

[212]

[213]

[214]

[215]

[216]

[217]

continued

884

The determination of trace elements in water

TABLE 26.5 (continuation) Analyte

Detection

Comments

Reference

Hg and CH3Hg

CV-AAS

[218]

Hg and CH3Hg Hg and CH3Hg

FI-HPLC-CV-AAS

Hg(II) and CH3Hg adsorbed from river and tap water onto a column packed with dithizone immobilized on sodium dodecyl sulfate coated alumina Hg(II) and CH3Hg were determined in brackish water using ethylation The efﬁciency of tetrabutylammonium bromide reagent and sodium chloride in a methanol– water mixture as mobile phase for HPLC separation of Hg and CH3Hg Collection and on-ship measurement procedures were described The automated speciation analyzer (ASA) for the determination of mercury and methylmercury in environmental samples

HPLC-CV-AAS

CV-AFS Hg, CH3Hg and dimethyl Hg Hg and GC–MIP-OES CH3Hg

26.7

[219] [220]

[221]

[222]

VOLTAMMETRY

Voltammetry is an analytical technique well suited for trace element determination in water samples, especially in sea water. Multielement and speciation analysis is possible, with detection limits in the range of 10210 – 10212 M. As well, the instrumentation is affordable, compact and can also be used on board ship [247–249]. Initially restricted to the few elements that form an amalgam with Hg, the technique has rapidly expanded to include numerous other elements through the use of cathodic stripping voltammetry (CSV) with adsorptive accumulation of a metal complex on the surface of a Hg drop (HMDE). Recently, the technique of catalytic CSV has further improved sensitivity for several elements. Catalytic CSV relies on the reoxidation of the reduction product during the negative potential scan. The reduction current is enhanced in the presence of the catalyst and detection limits in the picomolar range can be achieved. Table 26.7 summarizes voltammetric procedures for the determination of trace elements in water. This list highlights only a few publications, for more complete coverage, several excellent reviews of CSV methods for the

885

S.N. Willie TABLE 26.6 Chemiluminescence Element

Comments

References

Cd

On-line ﬂuorescence method. Cd was separated from sea water on Dowex resin FI-CL was used to determine Co and Fe in estuarine and coastal waters. Co(II) was determined by means of a pyrogallol–hydrogen peroxide– sodium hydroxide reaction in the presence of methanol and the surfactant cetyltrimethylammonium bromide (CTAB). Iron(II þ III) was determined using a luminol reaction with dissolved oxygen as the oxidant FI-CL method for the determination of Co(II) in sea water. Preconcentration using 8-hydroxyquinoline immobilized on silica gel, and CL detection using gallic acid –hydrogen peroxide The oxidation reaction between chromium(VI) and hexacyanoferrate(II) generated hexacyanoferrate(III) which subsequently reacted with luminol in alkaline aqueous solution to produce CL Automated FI-CL determination of Cr(III) in tap water. Cr(III) catalyzed reaction between luminol and hydrogen peroxide FI-CL determination of Cr(III) and Cr(VI) in waste water. Luminol oxidation by hydrogen peroxide. Cr(III) was determined directly, Cr(VI) was reduced to Cr(III) and Cr(VI) is obtained by difference A UV digestion system (batch and on-line) was used to destroy Cu complexing organic ligands in sea water samples prior to FI-CL determination of total Cu. On-line microcolumn preconcentration/matrix removal A ﬂow-through CL method for the determination of Cu in natural water samples. Luminol and cyanide were coimmobilized on an anion-exchange column and Cu was retained temporarily by electrochemical preconcentration on a gold electrode FI-CL determination of copper complexation in sea water. Cu complexation with 1,10-phenanthroline and oxidation with hydrogen peroxide Shipboard determination of dissolved iron by FI-CL. Dissolved Fe(II þ III) levels were determined after Fe(III) reduction using sulphite and on-line matrix elimination/preconcentration on an 8-hydroxyquinoline chelating resin column

[226]

Co, Fe

Co

Cr

Cr

Cr

Cu

Cu

Cu

Fe

[227]

[228]

[229]

[230]

[231]

[232]

[233]

[234]

[235]

continued

886

The determination of trace elements in water

TABLE 26.6 (continuation) Element

Comments

References

Fe

FI-CL determination of total Fe in sea water. On-line matrix elimination/preconcentration on an 8-hydroxyquinoline chelating resin column The determination of Fe(II) and total Fe in natural water samples. Amberlite XAD-4 functionalized by N-hydroxyethylethylenediamine used for preconcentration of iron. CL detection using brilliant sulfoﬂavine and hydrogen peroxide reagent solutions Fe(II) was preconcentrated on a 8-hydroxy quinoline column eluted with HCl and subsequently reacted with luminol and hydrogen peroxide. LOD 0.05 nM FI-CL method for the sequential determination of Fe(II) and Fe(III) in water. Fe(II) was detected by its catalytic effect on the CL reaction between luminol immobilized on an anion exchange resin column and dissolved oxygen; Fe(III) was determined by difference after on-line conversion to Fe(II) in a reducing minicolumn packed with Cu plated Zn granules FI-CL method for the determination of Fe(II) and total Fe in water. Silver reductor column reduced Fe(III) to Fe(II) for total Fe. Luminol and hydrogen peroxide used for CL reaction CL procedure using the reaction between Hg(II) and 6-mercaptopurine. A detection limit of 1.4 ng ml21 of Hg(II) was obtained Room temperature phosphorescence for the determination of Hg in aqueous media. alpha-Bromonaphthalene (BrN) was used as the phosphorescent donor molecule. A LOD of 14 ng ml21 was obtained FI-CL determination of Mn(II) in natural water with solid sodium bismuthate as an oxidant FI-CL method for the determination of Mn in sea water. Interference from Fe was removed by passing the sample through a 8-hydroxyquinoline immobilized chelating resin column prior to luminol–hydrogen peroxide CL detection A selective FI-CL method for the determination of V in environmental water. Based on the CL reaction between V(II) and luminal. V(II) was electrogenerated from vanadate using a ﬂow-through carbon electrolytic cell

[236]

Fe

Fe

Fe

Fe

Hg

Hg

Mn Mn

V

[237]

[238,239]

[240]

[241]

[242]

[243]

[244] [245]

[246]

887

S.N. Willie TABLE 26.7 Voltammetry Element

Comments

References

As

The determination of As(III) by differential pulse ASV using a gold disc electrode. The shift of the potential of the anodic dissolution peak of arsenic caused by adsorption of surface active substances is discussed. The method was applied for analysis of various types of natural waters after elimination of surface active compounds As in river and sea water was determined by square wave CSV in the presence of Cu and HCl. Sub nM LOD was achieved Determination of Al in natural waters by complexation with solochrome violet RS (SVRS) Determination of Al in sea water and fresh water by CSV with complexation with dihydroxyanthraquinone-3-sulphonic acid at pH 7.1 –7.3 Adsorptive accumulation of the Bi(III) alizarine red S complex on a HMDE followed by DPCSV. The procedure was applied to tap water Complexation of Cd in sea water with 5-ﬂuorouracil. Analysis by linear sweep CSV using a HDME. The method was applied to the determination of Cd in sea water Two approaches for modifying glassy carbon electrodes were discussed. The modiﬁed working electrodes were compared for the determination of Pb and Cu in sea water Differential pulse CSV and ASV measurements were achieved using a conventional three-electrode cell and the ammonia–ammonium chloride buffer at pH 9.4 as the supporting electrolyte. The analytical procedure was veriﬁed by the analysis of sea water BCR-CRM 403. Mono and bivariate analysis using standard additions was used A mixture of ligands permitted the simultaneous determination of several elements in sea water. Scans containing six resolved peaks corresponding to the listed analytes were obtained using a combination of dimethylglyoxime and 8-hydroxyquinoline in sea water. In unpolluted waters the sensitivity for Co and Zn was not sufﬁcient Adsorptive voltammetric technique using ammonium 2-amino-cyclopente dithiocarboxylate as a complexing agent. Scans contained three resolved peaks corresponding to the analytes. The method was applied to UV treated water samples

[253]

As

Al Al

Bi

Cd

Cu, Pb

Cu, Cd, As, Se, Zn and Mn

Cu, Pb, Cd, Ni, Co and Zn

Cd, Co and Ni

[254]

[255] [256]

[257]

[258]

[259]

[260]

[261]

[262]

continued

888

The determination of trace elements in water

TABLE 26.7 (continuation) Element

Comments

References

Cd, Pb

Pb and Cd were determined by subtractive ASV in the square wave mode at a silver electrode without removal of oxygen. The silver counter/ quasi-reference electrode generates silver ions that co-deposit with Pb and Cd at the Ag rotating disc electrode. As surfactants distort the signal, the samples were pretreated by wet ashing Simultaneous determination by CSV following absorption of 8-hydroxyquinoline complexes. Results similar to ASV with uncomplexed samples using a rotating disc electrode but superior to ASV using a HMDE DPCSV by adsorptive accumulation of the cobalt– nioxime complex onto a HMDE. The reduction current was catalytically enhanced by the presence of nitrite. A detection limit of 3 pM cobalt (2 ng l21) (at an adsorption period of 60 s) enables the detection of this metal in uncontaminated sea water. UV digestion of sea water was essential to destroy any organo-Co species Complexation with catechol and analysis of sea water by differential pulse CSV. The presence of dissolved oxygen created unstable signals Differential pulse catalytic-CSV procedure using N-(2-hydroxyethyl) ethylenediamine-N,N0 ,N0 -triacetic acid (HEDTA) as a complexing agent. The procedure was applied to the determination of chromium in different water types. Cr(III) was oxidized to Cr(VI) with peroxide for total Cr The determination of total and Cr species in river and lake water based on catalytic CSV with adsorption of Cr(III)– diethylenetriaminepentaacetic acid complexes. Electrochemically distinct behaviors of Cr(III) and Cr(VI) were presented Catalytic CSV based on the addition of 1-nitroso-2-naphthol and bromate. The detection limit is 0.08 nM (5 ng l21)Fe after an adsorption time of 30 s; the detection limit can be lowered by increasing the adsorption time. The method can also be used to determine the redox speciation of iron: Fe-III is determined speciﬁcally by masking Fe-II with 2,2-bipyridyl. The concentration of Fe-II can be calculated by difference from the total Fe concentration

[263]

Cu, Cd and Pb

Co

Cu

Cr

Cr

Fe

[264]

[265]

[266]

[267]

[268]

[269]

continued

889

S.N. Willie

TABLE 26.7 (continuation) Element

Comments

References

Fe

Competitive ligand exchange-adsorptive CSV method for the determination of Fe speciation in sea water using 2-(2-thiazolylazo)-p-cresol. The detection limit is 0.10 nM with an adsorption time of 300 s. The method was applied to Swedish coastal water samples Catalytic CSV based on the adsorptive accumulation of the Fe(III)-2,3-dihydroxynaphthalene complex at pH 8.0 on a HMDE. The reduction current is catalytically enhanced by the presence of bromate. A detection limit of 13 pM (0.7 ng l21) Fe was achieved using a 60 s adsorption period DPCSV following adsorption of the Ni and Co dimethylglyoxime complex at pH 9 The determination of dissolved Mn(II) in coastal and estuarine waters and sea water utilizing CSV on a rotating glassy carbon disk electrode. A sensitivity of 11 nA nM21 min21, and a detection limit of 6 nM (0.3 ng ml21) were achieved Cathodic stripping square wave voltammetry of the Mo– 8-hydroxyquinoline complex on a HMDE. This procedure was found to be superior to DPCSV due to the faster scan rate and improved calibration. The applicability of this method to analysis of sea water was assessed by the determination of Mo in two CRMs An adsorptive CSV method based on the catalytic effect of the Pt– formazone complex with Pt on the development of hydrogen at a mercury electrode A method for the determination of Sb(III) and total Sb in sea water samples by DPASV. The method is based on the distinct electrochemical behavior of Sb(III) and Sb(V) with different acid strengths. Sb(III) was determined in 0.5 mol l21 HCl and total antimony in 5 mol l21 HCl. No previous elimination of organic matter was needed A ﬂow-through cell with the mercury ﬁlm electrode based on epoxy resin impregnated graphite. The detection limit of the method was 2 ng l21 using 90 min preconcentration in a circulating ﬂow system. The determination of Tl in tap water and river water was reported A CSV method for the direct determination of U in sea water based on the adsorption of the U catechol complex

[270]

Fe

Ni, Co Mn

Mo

Pt

Sb

Tl

U

[271]

[272] [273]

[274]

[275]

[276]

[277]

[278]

continued

890

The determination of trace elements in water

TABLE 26.7 (continuation) Element

Comments

References

V

A method based on the preconcentration of the V(V)–2-(5-bromo-2-pyridylazo)-5-diethylaminophenol (5-Br-PADAP) complex. The catalytic action of bromate ions on the reduction of the V(V)– 5-Br-PADAP complex enhanced the sensitivity. The method of standard additions was employed in evaluating the concentration of vanadium in water samples CSV procedure using the catalytic effect of V on the bromate oxidation of gallocyanine. A detection limit of 0.05 ng ml21 and calibration graph from 0.30 to 200 ng ml21 vanadium was obtained. Vanadium in river water was determined by this method The interference by Cu in the determination of Zn by ASV was removed by the addition of sulphide. The proposed method was found to be applicable to a number of environmental samples. The calibration graph was linear from 0.3 to 11 mg of zinc. The recovery of zinc in three real samples and two synthetic sea water samples spiked with 10 trace metals using the proposed method ranged from 98 to 102%, and the precision of the method was 1.6% CSV procedure based on the complexation of Zn with APDC. A LOD of 0.03 nM was obtained

[279]

V

Zn

Zn

[280]

[281]

[282]

determination of trace elements in sea water have been published [250–252]. 26.8

TOTAL-REFLECTION X-RAY FLUORESCENCE SPECTROMETRY

Total-reﬂection X-Ray ﬂuorescence spectrometry (TXRF) is a competitive technique for the determination of trace elements in water due to its multielement capability, simple quantitation and small sample mass required. Typically, sample preparation involves ﬁltration and evaporation of an aliquot on a silica sample carrier. For saline matrices, analyte separation from the matrix is required. Instrumental detection limits are a few picogram for most of elements ðZ . 13Þ: However, some form of analyte concentration is usually required. Klockenkamper and vonBohlen [283] have prepared a review of TXRF that illustrates its applicability to environmental analyses using some typical examples (Table 26.8).

891

S.N. Willie TABLE 26.8 X-ray Elements

Comments

References

Na, Mg, K, Ca, Ni, Cu, Zn and Sr

An intercomparison evaluating the performance of a TXRF and grazing emission XRF (GEXRF) for the analysis of mineral water samples. It was concluded that the TXRF technique is suitable for the direct determination of heavy elements in drinking water (above Z ¼ 19) while GEXRF spectrometry can be successfully applied to the determination of low-Z elements A poly(acrylamidoxime) cloth incorporated into a continuous ﬂow chamber was used to collect trace metals from water. The metals were analyzed using wavelength dispersive XRF Non-saline water sample aliquots were dried on quartz glass. Brackish water required separation of the analytes by complexation with sodium dibenzyldithiocarbamate, adsorption on a column and elution with methanol. High organics required sample digestion Preconcentration by complexation using a mixture of carbamates followed by solvent extraction with methyl isobutyl ketone. The minimum detection limits for the determination of these elements in mineral and tap water samples were found to be approximately 60 ng l21 Sample aliquots (20 ml) of drinking water were dried on a quartz substrate. The detection limits were in the range 0.5 –1.7 mg l21 for Cr, Mn, Fe, Co, Ni, Cu, Zn, As, Se, Rb, Sr, Hg and Pb and 4.9– 11 mg l21 for K, Ca, V, Cd, Sb and Ba. Ga used as internal standard Trace metal concentrations of mineral spring waters determined by TXRF using an internal Ga standard. The element concentrations obtained by TXRF are comparable to results obtained by ET-AAS after separation–preconcentration on an Amberlite XAD-4 column and ICP-MS The simultaneous determination of hydride forming elements in sea water by TXRF was investigated using the following procedures: (1) preconcentration of hydrides by absorption in solvents, (2) decomposition of hydrides in a heated silica tube, (3) decomposition of hydrides directly on the surface of a heated silica sample carrier as a thin amorphous ﬁlm, and (4) combustion of hydrides in the hydrogen ﬂame and deposition of an elemental ﬁlm on the sample carrier

[285]

Pb, Cu, Zn, Cd, Mn, Fe and Mg

Multielement

Cu, Hg and Pb

Multielement

Multielement

Ge, As, Sn and Sb

[286]

[287,288]

[289]

[290]

[291]

[292]

continued

892

The determination of trace elements in water

TABLE 26.8 (continuation) Elements

Comments

References

Fe, Ni, Cu, Zn and Pb

The determination of trace elements in mineral water by coprecipitation with APDC. The sample was ﬁltered and a small amount placed on a ﬁlter and analyzed by TXRF An evaluation of metal contamination in estuarine waters was conducted using two analytical processes: soluble metals were examined by radioisotope XRF (RIXRF) and soluble and insoluble metals by TRXRF. The soluble heavy metals determined were Fe, Ni, Cu, Zn and Pb and the soluble and insoluble metals were Ti, Mn, Fe, Co, Ni, Cu, Zn, As and Pb Multielement determination in pore water and river water was carried out using TXRF. Aliquots were dried on sample carrier following ﬁltration. Yttrium was used as internal standard Preconcentration by complexation with dibenzyldithiocarbamate and collection of the complexes on a silica gel column. The eluted concentrate was analyzed by TXRF The energy-dispersive miniprobe multielement XRF was used to measure trace elements in small samples. Liquid samples (100–150 ml) were evaporated onto a Teﬂon ﬁlm in increments of 10 ml. Detection limits ranged from 20 pg for Sr to 700 pg for Pb. NIST 1643c, Trace Elements in Water was analyzed Coastal sea water samples were concentrated by precipitation as dithiocarbamates, ﬁltered and analyzed by TXRF Estuarine water samples were prepared for analysis by TXRF using electrodeposition on a rotating polished glassy carbon disc working electrode. Typical detection limits range from 5 to 20 ng l21. Results were obtained for the certiﬁed reference material CRM 505 For direct analysis of waste water samples the preparation of quasi-solid specimens based on gelatin (agar) was evaluated. Such specimens may also be prepared following extraction of diethyldithiocarbamate complexes and their back-extraction into an aqueous phase. An alternate method using lowtemperature crystallization of water creating a glassy saccharose-based specimen from aqueous concentrates was also proposed

[293]

Fe, Ni, Cu, Zn and Pb

Cu, Pb and Zn

V, Cr, Mn, Fe, Ni, Cu, Zn, As, Se, Pb and U As, Cr, Cu, Fe, Ga, Ge, Hf, Mn, Ni, Pb, Rb, Se, Sr, Th, Y, U and Zn V, Fe, Co, Ni, Cu, Zn, As, Cd and Pb Cr, Mn, Fe, Co, Ni, Cu, Zn, Ag, Cd, Pb, As and U

Ag, As, Bi, Cd, Co, Cu, Fe, Ga, Ni, Pb, Se and Zn

[294]

[295]

[296]

[297]

[298]

[299]

[300]

893

S.N. Willie

For recent developments in TXRF, a good source of information are the special issues of Spectrochimica Acta B published in alternate years subsequent to an international conference devoted to the topic [284].

26.9

CONCLUSIONS

There is a wide choice of analytical techniques available for the determination of trace elements in water. For single element analysis, many cost-effective analytical procedures that utilize voltammetry or CL are available. In most cases, these procedures possess sufﬁcient detection power to rival more expensive instrumental techniques. HG and cold vapor procedures satisfactorily ﬁll certain voids left by the other techniques. The advent of ICP-MS has had a profound consequence on the multielement determination of trace elements in water. In particular, procedures that automate many repetitive chemical processing steps through the utilization of on-line sample pretreatment have had a signiﬁcant effect. Beneﬁts that include: enhanced throughput, reduced contamination, increased detection power and the removal of interferences, all result in the improved reliability of inorganic analytical measurements. More recently, the availability of reaction cell technology and SF-ICP-MS instrumentation will further enhance the analysts’ capability.

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13

894

S.J. Hill, T.A. Arowolo, O.T. Butler, S.R.N. Chenery, J.M. Cook, M.S. Cresser and D.L. Miles, J. Anal. At. Spectrom., 17 (2002) 284. M.R. Cave, O. Butler, J.M. Cook, M.S. Cresser, L.M. Garden, A.J. Holden and D.L. Miles, J. Anal. At. Spectrom., 15 (2000) 489. M.R. Cave, O. Butler, S.R.N. Chenery, J.M. Cook, M.S. Cresser and D.L. Miles, J. Anal. At. Spectrom., 16 (2001) 194. S.J. Hill, S. Chenery, J.B. Dawson, E.H. Evans, A. Fisher, W.J. Price, C.M.M. Smith, K.L. Sutton and J.F. Tyson, J. Anal. At. Spectrom., 15 (2000) 763. E.H. Evans, J.B. Dawson, A. Fisher, S.J. Hill, W.J. Price, C.M.M. Smith, K.L. Sutton and J.F. Tyson, J. Anal. At. Spectrom., 16 (2001) 672. S.D. Richardson, Anal. Chem., 71 (1999) 181R. S.D. Richardson, Anal. Chem., 73 (2001) 2719. R.E. Clement and P.W. Yang, Anal. Chem., 69 (1997) 251R. R.E. Clement and P.W. Yang, Anal. Chem., 71 (1999) 257R. R.E. Clement and P.W. Yang, Anal. Chem., 73 (2001) 2761. W.A. Telliard, Crit. Rev. Anal. Chem., 29 (1999) 249. Technical Notes on Drinking Water Analysis, EPA/600/R-94/173, USEPA, Washington, DC, 1998. Annual Book of ASTM Standards, 1994 and 1996, Vols. 11.01 and 11.02, American Society for Testing and Materials, West Conshohocken, PA, 19428-2959 USA.

The determination of trace elements in water 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50

Ofﬁcial Methods of Analysis of AOAC International, Arlington, VA. Vol. 17th Edition, 2000. B. Sansoni, W. Brunner, G. Wolff, H. Ruppert and R. Dittrich, Fresenius J. Anal. Chem., 331 (1988) 154. N.J. Miller-Ihli and S.A. Baker, J. Food Compos. Anal., 14 (2001) 619. L. Halicz, I. Segal and O. Yoffe, J. Anal. At. Spectrom., 14 (1999) 1579. E. Rosland and W. Lund, J. Anal. At. Spectrom., 13 (1998) 1239. M.P. Field, J.T. Cullen and R.M. Sherrell, J. Anal. At. Spectrom., 14 (1999) 1425. M.M. Bayon, J.I.G. Alonso and A. SanzMedel, J. Anal. At. Spectrom., 13 (1998) 1027. H. Louie, M. Wu, P. Di, P. Snitch and G. Chapple, J. Anal. At. Spectrom., 17 (2002) 587. P. Bermejo-Barrera, J. Moreda-Pin˜eiro, A. Moreda-Pin˜eiro and A. BermejoBarrera, Talanta, 45 (1998) 807. P. Bermejo-Barrera, J. Moreda-Pin˜eiro, A. Moreda-Pin˜eiro and A. BermejoBarrera, Talanta, 43 (1996) 35. P. Bermejo-Barrera, J. Moreda-Pin˜eiro, A. Moreda-Pin˜eiro and A. BermejoBarrera, Fresenius J. Anal. Chem., 355 (1996) 174. J.Y. Cabon, Fresenius J. Anal. Chem., 367 (2000) 714. O. Acar, A.R. Turker and Z. Kilic, Talanta, 49 (1999) 135. S.-D. Huang and K.-Y. Shih, Spectrochim. Acta B, 50 (1995) 837. S. Salomon, P. Giamarchi, A. Le Bihan, H. Becker-Ross and U. Heitmann, Spectrochim. Acta B, 55 (2000) 1337. L. Yan-zhong, L. Mei and R. Zhu, Fresenius J. Anal. Chem., 357 (1997) 112. J.Y. Cabon, Spectrochim. Acta B, 57 (2002) 513. J.Y. Cabon, Spectrochim. Acta B, 57 (2002) 939. J.Y. Cabon and A. Le Bihan, Spectrochim. Acta B, 50 (1995) 1703. R. Guevremont, Anal. Chem., 52 (1980) 1574. R. Guevremont, Anal. Chem., 53 (1981) 911. D.C. Gre´goire and M.L. Ballinas, Spectrochim. Acta B, 52 (1997) 75. G. Chapple and J.P. Byrne, J. Anal. At. Spectrom., 11 (1996) 549. C.-C. Chang and S.-J. Jiang, Anal. Chim. Acta, 353 (1997) 173. J.P. Byrne and G. Chapple, At. Spectrosc., 19 (1998) 116. H.-W. Liu, S.-J. Jiang and S.-H. Liu, Spectrochim. Acta B, 54 (1999) 1367. J.Z. Zhang, C. Kelble and F.J. Millero, Anal. Chim. Acta, 438 (2001) 49. W.S. Yao and R.H. Byrne, Talanta, 48 (1999) 277. I. Rodushkin and T. Ruth, J. Anal. At. Spectrom., 12 (1997) 1181. R. Hoelzl, L. Fabry, L. Kotz and S. Pahlke, Fresenius J. Anal. Chem., 366 (2000) 64. J.A. McLean, J.S. Becker, S.F. Boulyga, H.J. Dietze and A. Montaser, Int. J. Mass Spectrom., 208 (2001) 193. C. Barbante, G. Cozzi, G. Capodaglio, K. van de Velde, C. Ferrari, C. Boutron and P. Cescon, J. Anal. At. Spectrom., 14 (1999) 1433. M.S. Choi, R. Francois, K. Sims, M.P. Bacon, S. Brown-Leger, A.P. Fleer, L. Ball, D. Schneider and S. Pichat, Mar. Chem., 76 (2001) 99. P. Bermejo-Barrera, J. Moreda-Pin˜eiro, A. Moreda-Pin˜eiro and A. BermejoBarrera, J. Anal. At. Spectrom., 13 (1998) 777. M.S. Chan and S.D. Huang, Talanta, 51 (2000) 373. C.Y. Chen, K.S.K. Danadurai and S.D. Huang, J. Anal. At. Spectrom., 16 (2001) 404. P.G. Su and S.D. Huang, Spectrochim. Acta B, 53 (1998) 699.

895

S.N. Willie 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89

896

H. Sun, L. Yang, D. Zhang and J. Sun, Talanta, 44 (1997) 1979. J.P. Riley and D. Taylor, Anal. Chem. Acta, 40 (1968) 479. M.J. Bloxham, S.J. Hill and P.J. Worsfold, J. Anal. At. Spectrom., 9 (1994) 935. J.W. McLaren, J.W.H. Lam, S.S. Berman, K. Akatsuka and M.A. Azeredo, J. Anal. At. Spectrom., 8 (1993) 279. L. Ebdon, A. Fisher, H. Handley and P. Jones, J. Anal. At. Spectrom., 8 (1993) 979. S. Hirata, Y. Ishida, M. Aihara, K. Honda and O. Shikino, Anal. Chim. Acta, 438 (2001) 205. D.B. Taylor, H.M. Kingston, D.J. Nogay, D. Koller and R. Hutton, J. Anal. At. Spectrom., 11 (1996) 187. Y.-H. Sung, Z.-S. Liu and S.-D. Huang, Spectrochim. Acta B, 52 (1997) 755. S.M. Nelms, G.M. Greenway and D. Koller, J. Anal. At. Spectrom., 11 (1996) 907. S.N. Willie, Y. Iida and J.W. McLaren, At. Spectrosc., 19 (1998) 67. S.N. Willie, J.W.H. Lam, L. Yang and G. Tao, Anal. Chim. Acta, 447 (2001) 143. K.W. Warnken, D.G. Tang, G.A. Gill and P.H. Santschi, Anal. Chim. Acta, 423 (2000) 265. S.N. Willie and R.E. Sturgeon, Spectrochim. Acta B, 56 (2001) 1707. G.M. Greenway, S.M. Nelms and D. Koller, Anal. Commun., 33 (1996) 57. N.G. Beck, R.P. Franks and K.W. Bruland, Anal. Chim. Acta, 455 (2002) 11. M. Kuhn and M. Kriews, Fresenius J. Anal. Chem., 367 (2000) 440. R.A. Nickson, S.J. Hill and P.J. Worsfold, Int. J. Environ. Anal. Chem., 75 (1999) 57. T.P. Rao and J.M. Gladis, Anal. Sci., 18 (2002) 517. R.E. Sturgeon, S.S. Berman, S.N. Willie and J.A.H. Desaulniers, Anal. Chem., 53 (1981) 2337. S.N. Willie, H. Tekgul and R.E. Sturgeon, Talanta, 47 (1999) 439. H. Dierssen, W. Balzer and W.M. Landing, Mar. Chem., 73 (2001) 173. A.T. Townsend, J. O’Sullivan, A.M. Featherstone, E.C.V. Butler and D.J. Mackey, J. Environ. Monit., 3 (2001) 113. M. Nicolaı¨, C. Rosin, N. Tousset and Y. Nicolai, Talanta, 50 (2000) 433. M.S. Jime´nez, R. Velarte and J.R. Castillo, Spectrochim. Acta B, 57 (2002) 391. G.M. Greenway, S.M. Nelms, I. Skhosana and S.J.L. Dolman, Spectrochim. Acta B, 51 (1996) 1909. S.M. Nelms, G.M. Greenway and R.C. Hutton, J. Anal. At. Spectrom., 10 (1995) 929. G.E.M. Hall, J.E. Vaive and J.C. Pelchat, J. Anal. At. Spectrom., 11 (1996) 779. X. Wang, Z. Zuang, C. Yang and F. Zhyu, Spectrochim. Acta B, 53 (1998) 1437. C.N. Ferrarello, M.M. Bayo´n, J.I.G. Alonso and A. Sanz-Medel, Anal. Chim. Acta, 429 (2001) 227. C. Gue´guen, J. Dominik and D. Perret, Fresenius J. Anal. Chem., 370 (2001) 909. K.E. Jarvis, J.G. Williams, E. Alcantara and J.D. Wills, J. Anal. At. Spectrom., 11 (1996) 917. W. Klemm, G. Bombach and K.P. Becker, Fresenius J. Anal. Chem., 364 (1999) 429. H.T. Lu, S.F. Mou, Y. Yan, S.Y. Tong and J.M. Riviello, J. Chromatogr. A, 800 (1998) 247. S. Motellier and H. Pitsch, J. Chromatogr. A, 739 (1996) 119. E. Santoyo, S. Santoyo Gutie´rrez and S.P. Verma, J. Chromatogr. A, 884 (2000) 229. X.J. Li, P. Schramel, H.Z. Wang, P. Grill and A. Kettrup, Fresenius J. Anal. Chem., 356 (1996) 52. A. Ali, X. Yin, H. Shen, Y. Ye and X. Gu, Anal. Chim. Acta, 392 (1999) 283. V.L. Dressler, D. Pozebon and A.J. Curtius, Spectrochim. Acta B, 53 (1998) 1527. I. Narin, M. Saylak, I. Elci and M. Dogan, Anal. Lett., 34(11) (2001) 1935.

The determination of trace elements in water 90 91 92

93 94 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109 110 111 112 113 114 115 116 117 118 119 120 121 122 123

R.A. Nickson, S.J. Hill and P.J. Worsfold, Int. J. Environ. Anal. Chem., 75 (1999) 57. G.J. Batterham, N.C. Munksgaard and D.L. Parry, J. Anal. At. Spectrom., 12 (1997) 1277. Y. Sohrin, S.-i. Iwamoto, S. Akiyama, T. Fujita, T. Kugii, H. Obata, E. Nakayama, S. Goda, Y. Fujishima, H. Hasegawa, K. Ueda and M. Matsui, Anal. Chim. Acta, 363 (1998) 11. O. Vicente, A. Padro´, L. Martinez, R. Olsina and E. Marchevsky, Spectrochim. Acta B, 53 (1998) 1281. K.W. Warnken, G.A. Gill, L.S. Wen, L.L. Grifﬁn and P.H. Santschi, J. Anal. At. Spectrom., 14 (1999) 247. B. Wen, X.-Q. Shan and S.-G. Xu, Analyst, 124 (1999) 621. H. Kumagai, M. Yamanaka, T. Sakai, T. Yokoyama, T.M. Suzuki and T. Suzuki, J. Anal. At. Spectrom., 13 (1998) 579. B. Wen, X.Q. Shan, R.X. Liu and H.X. Tang, Fresenius J. Anal. Chem., 363 (1999) 251. J. Wu and E.A. Boyle, Anal. Chem., 69 (1997) 2464. P.-H. Lin, K.S.K. Danadurai and S.-D. Huang, J. Anal. At. Spectrom., 16 (2001) 409. E.A. Boyle and J.M. Edmond, Anal. Chim. Acta, 91 (1977) 189. S. Zhuang, X. Wang, P. Yang, C. Yang and B. Huang, Can. J. Appl. Spectrosc., 39 (1994) 101. E. Ivanova, K. Benkhedda and F. Adams, J. Anal. At. Spectrom., 13 (1998) 527. V. Porta, O. Abollino, E. Mentasti and C. Sarzanini, J. Anal. At. Spectrom., 6 (1991) 119. T. Akagi, K. Fuwa and H. Haraguchi, Anal. Chim. Acta, 177 (1985) 139. Z.X. Zhuang, C.L. Yang, X.R. Wang, P.Y. Yang and B.L. Huang, Fresenius J. Anal. Chem., 355 (1996) 277. S. Sella, R.E. Sturgeon, S.N. Willie and R.C. Campos, J. Anal. At. Spectrom., 12 (1997) 1281. T. Yabutani, S. Ji, F. Mouri, A. Itoh, K. Chiba and H. Haraguchi, Bull. Chem. Soc. Jpn., 73 (2000) 895. K. Benkhedda, H.G. Infante, E. Ivanova and F.C. Adams, J. Anal. At. Spectrom., 16 (2001) 995. K. Benkhedda, H.G. Infante, E. Ivanova and F.C. Adams, J. Anal. At. Spectrom., 15 (2000) 1349. H.H. Chen and D. Beauchemin, J. Anal. At. Spectrom., 16 (2001) 1356. Z.S. Chen, M. Hiraide and H. Kawaguchi, Mikrochim. Acta, 124 (1996) 27. G.E.M. Hall and J.C. Pelchat, J. Anal. At. Spectrom., 8 (1993) 1059. E. Bulska, M. Walcerz, W. Jedral and A. Hulanicki, Anal. Chim. Acta, 357 (1997) 133. S. Scaccia, G. Zappa and N. Basili, J. Chromatogr. A, 915 (2001) 167. E.R. Unsworth, J.M. Cook and S.J. Hill, Anal. Chim. Acta, 442 (2001) 141. E. Engstro¨m, I. Jo¨nebring and B. Karlberg, Anal. Chim. Acta, 371 (1998) 227. R.J. Rosenberg, J. Trace Microprobe Tech., 14 (1996) 325. USEPA, http://www.epa.gov/safewater/ndwacsum.html(ndwac, August 14, 2001. K.S. Johnson, R.M. Gordon and K.H. Coale, Mar. Chem., 57 (1997) 137. A. D’Ulivo, Analyst, 122 (1997) 117R. L. Yang and R.E. Sturgeon, J. Anal. At. Spectrom., 17 (2002) 88. X.J. Yang, R. Foley and G.K.-C. Low, Analyst, 127 (2002) 315. D. Yuan and I.L. Shuttler, Anal. Chim. Acta, 316 (1995) 313.

897

S.N. Willie 124 125 126 127 128 129 130 131 132 133 134 135 136 137 138 139 140 141 142 143 144 145 146 147 148 149 150 151 152 153 154 155 156 157 158 159 160 161

898

S. Imai, K. Fujikawa, A. Yonetani, N. Ogawa and Y. Kikuchi, Anal. Sci., 16 (2000) 163. T. Kubota, T. Yamaguchi and T. Okutani, Talanta, 46 (1998) 1311. Y. Yamini, J. Hassan, R. Mohandesi and N. Bahramifar, Talanta, 56 (2002) 375. Y. Yamini, M. Chaloosi and H. Ebrahimzadeh, Talanta, 56 (2002) 797. D. Colbert, K.S. Johnson and K.H. Coale, Anal. Chim. Acta, 377 (1998) 255. Z.-R. Xu, H.-Y. Pan, S.-K. Xu and Z.-L. Fang, Spectrochim. Acta B, 55 (2000) 213. P.-G. Su and S.-D. Huang, Anal. Chim. Acta, 376 (1998) 305. B. Paull, E. Twohill and W. Bashir, J. Chromatogr. A, 877 (2000) 123. K. Akatsuka, Y. Yoshida, N. Nobuyama, S. Hoshi, S. Nakamura and T. Kato, Anal. Sci., 14 (1998) 529. K. Benkhedda, H.G. Infante, E. Ivanova and F. Adams, J. Anal. At. Spectrom., 15 (2000) 429. S. Hirata, K. Honda, O. Shikino, N. Maekawa and M. Aihara, Spectrochim. Acta B, 55 (2000) 1089. Q. Zhang, H. Minami, S. Inoue and I. Atsuya, Anal. Chim. Acta, 401 (2000) 277. A. Ali, Y. Ye, G. Xu and X. Yin, Fresenius J. Anal. Chem., 365 (1999) 642. A.N. Anthemidis, G.A. Zachariadis and J.A. Stratis, Talanta, 54 (2001) 935. E.P. Achterberg, T.W. Holland, A.R. Bowie, R.F.C. Mantoura and P.J. Worsfold, Anal. Chim. Acta, 442 (2001) 1. J. Wu and E.A. Boyle, Anal. Chim. Acta, 367 (1998) 183. C.I. Measures, J. Yuan and J.A. Resing, Mar. Chem., 50 (1995) 3. D.A. Weeks and K.W. Bruland, Anal. Chim. Acta, 453 (2002) 21. S.K. Sahoo and A. Masuda, Anal. Chim. Acta, 370 (1998) 215. D. Weiss, E.A. Boyle, V. Chavagnac, M. Herwegh and J. Wu, Spectrochim. Acta B, 55 (2000) 363. S. Kagaya, Y. Araki and K. Hasegawa, Fresenius J. Anal. Chem., 366 (2000) 842. A.E. Eroglu, C.W. McLeod, K.S. Leonard and D. McCubbin, Anal. Chim. Acta, 53 (1998) 1221. T. Kubota, A. Kawakami, T. Sagara, N. Ookubo and T. Okutani, Talanta, 53 (2001) 1117. M.J. Keith-Roach, S. Stu¨rup, D.H. Oughton and H. Dahlgaard, Analyst, 127 (2002) 70. C.L. Chou and J.D. Moffatt, Fresenius J. Anal. Chem., 368 (2000) 59. T. Yamane, Y. Osada and M. Suzuki, Talanta, 45 (1998) 583. K. Akatsuka, T. Katoh, N. Nobuyama, T. Okanaka, M. Okumura and S. Hoshi, Anal. Sci., 12 (1996) 209. K. Norisuye, H. Hasegawa, S. Mito, Y. Sohrin and M. Matsui, Talanta, 53 (2000) 639. J. Dedina and D.L. Tsalev, Hydride Generation Atomic Absorption Spectrometry. Wiley, New York, 1995. R.K. Anderson, M. Thompson and E. Culbard, Analyst, 111 (1986) 1153. B. Klaue and J.D. Blum, Anal. Chem., 71 (1999) 1408. S. Willie, Spectrochim. Acta B, 51 (1996) 1781. I.D. Brindle, X.C. Le and X.F. Li, Analyst, 113 (1988) 1377. I.D. Brindle, X.C. Le and X.F. Li, J. Anal. At. Spectrom., 4 (1989) 227. J.M.G. LaFuente, M.L.F. Sanchez, J.M. Marchante-Gayon, J.E.S. Uria and A. Sanz-Medel, Spectrochim. Acta B, 51 (1996) 1849. R.E. Sturgeon, S.N. Willie and S.S. Berman, J. Anal. At. Spectrom., 4 (1989) 443. C.J. Park and S.A. Yim, J. Anal. At. Spectrom., 14 (1999) 1061. R.E. Sturgeon, S.N. Willie and S.S. Berman, Anal. Chem., 61 (1989) 1867.

The determination of trace elements in water 162 163 164 165 166 167 168 169 170 171 172 173 174 175 176 177 178 179 180 181 182 183 184 185 186 187 188 189 190 191 192 193 194 195 196 197 198 199

R.E. Sturgeon and Z.Z. Mester, Appl. Spectrosc., 56 (2002) 202A– 213A. I.D. Brindle, H. Alarabi, X.C. Le, S. Zheng and H. Chen, Analyst, 117 (1992) 407. J. Moreda-Pin˜eiro, M.L. Cervera and M. de la Guardia, J. Anal. At. Spectrom., 12 (1997) 1377. J.T. Creed, M.L. Magnuson, C.A. Brockhoff, I. Chamberlain and M. Sivaganesan, J. Anal. At. Spectrom., 11 (1996) 505. L.A. Liang, S. Lazoff, C. Chan, M. Horvat and J.S. Woods, Talanta, 47 (1998) 569. L.F.R. Machado, A.O. Jacintho, A.A. Menegario, E.A.G. Zagatto and M.F. Gine, J. Anal. At. Spectrom., 13 (1998) 1343. R.Q. Aucelio, V.N. Rubin, E. Becerra, B.W. Smith and J.D. Winefordner, Anal. Chim. Acta, 350 (1997) 231. J.X. Li, F. Lu, T. Umemura and K. Tsunoda, Anal. Chim. Acta, 419 (2000) 65. P. Bermejo-Barrera, J. Moreda-Pin˜eiro, A. Moreda-Pin˜eiro and A. BermejoBarrera, Anal. Chim. Acta, 368 (1998) 281. S.C. Apte and A.G. Howard, J. Anal. Spectrom., 1 (1986) 221. M.E. Moreno, C. Perez-Conde and C. Camara, J. Anal. At. Spectrom., 13 (1998) 1181. W.W. Ding and R.E. Sturgeon, J. Anal. At. Spectrom., 11 (1996) 225. F. Barbosa, S.S. de Souza and F.J. Krug, J. Anal. At. Spectrom., 17 (2002) 382. X.M. Guo and X.W. Guo, J. Anal. At. Spectrom., 16 (2001) 1414. M.T. Wei and S.J. Jiang, J. Anal. At. Spectrom., 14 (1999) 1177. S.J. Santosa, H. Mokudai and S. Tanaka, J. Anal. At. Spectrom., 12 (1997) 409. J. Bowman, B. Fairman and T. Catterick, J. Anal. At. Spectrom., 12 (1997) 313. C.S. Chen and S.J. Jiang, Spectrochim. Acta B, 51 (1996) 1813. F. El-Hadri, A. Morales-Rubio and M. de la Guardia, Talanta, 52 (2000) 653. L. Wei, P. Gupta, R. Hernandez and F. Farhat, Microchem. J., 62 (1999) 83. Z. Mester, R.E. Sturgeon and J.W. Lam, J. Anal. At. Spectrom., 15 (2000) 1461. C.P. Ferrari, A.L. Moreau and C.F. Boutron, Fresenius J. Anal. Chem., 366 (2000) 433. Ofﬁce of Water, EPA-821-R-02-019, USEPA, Washington, DC, August 2002. K.J.M. Kramer, P. Quevauviller, W. Dorten, E.M. van der Vlies and H.P.M. de Haan, Analyst, 123 (1998) 950. R. Puk and J.H. Weber, Appl. Organomet. Chem., 8 (1994) 293. C.F. Harrington, TRAC Trends Anal. Chem., 19 (2000) 167. G.A. Gill and W. Fitzgerald, Mar. Chem., 20 (1987) 227. N.S. Bloom and E.A. Crecelius, Mar. Chem., 14 (1983) 49. B.A. Ferna´ndez, M.R. Ferna´ndez de la Campa and A. Sanz-Medel, J. Anal. At. Spectrom., 8 (1993) 1097. L. Liang and N.S. Bloom, J. Anal. At. Spectrom., 8 (1993) 591. M.J. Bloxham, S.J. Hill and P.J. Worsfold, J. Anal. At. Spectrom., 11 (1996) 511. G.E.M. Hall, J.C. Pelchat, P. Pelchat and J.E. Vaive, Analyst, 127 (2002) 674. E. Debrah, J.F. Tyson and E.R. Denoyer, J. Anal. At. Spectrom., 11 (1996) 127. W.E. Doering, R.R. James and R.T. Echols, Fresenius J. Anal. Chem., 368 (2000) 475. O. Wurl, O. Elsholz and R. Ebinghaus, Talanta, 52 (2000) 51. J.C.A. de Wuilloud, R.G. Wuilloud, J.A. Gasquez, R.A. Olsina and L.D. Martinez, At. Spectrosc., 22 (2001) 398. L. Balint, I. Vedrina-Dragojevic, B. Sebecic, J. Momirovic-Culjat and M. Horvatic, Mikrochim. Acta, 127 (1997) 61. R.C. Campos, C.L.P. daSilveira and R. Lima, At. Spectrosc., 18 (1997) 55.

899

S.N. Willie 200 201 202 203 204 205 206 207 208 209 210 211 212 213 214 215 216 217 218 219 220 221 222 223

224 225 226 227 228 229

900

L.R. Bravo-Sa´nchez, B. San Vicente de la Riva, J.M. Costa-Ferna´ndez, R. Pereiro and A. Sanz-Medel, Talanta, 55 (2001) 1071. P. Bermejo-Barrera, J. Moreda-Pin˜eiro, A. Moreda-Pin˜eiro and A. BermejoBarrera, Mikrochim. Acta, 124 (1996) 111. P. Bermejo-Barrera, J. Moreda-Pin˜eiro, A. Moreda-Pin˜eiro and A. BermejoBarrera, J. Anal. At. Spectrom., 12 (1997) 317. C.C. Chang and S.J. Jiang, Anal. Chim. Acta, 353 (1997) 173. S.J. Christopher, S.E. Long, M.S. Rearick and J.D. Fassett, Anal. Chem., 73 (2001) 2190. J.C.A. de Wuilloud, R.G. Wuilloud, M.F. Silva, R.A. Olsina and L.D. Martinez, Spectrochim. Acta B, 57 (2002) 365. K. Nagashima, T. Murata and K. Kurihara, Anal. Chim. Acta, 454 (2002) 271. H. Sakamoto, J. Taniyama and N. Yonehara, Anal. Sci., 13 (1997) 771. D. Karunasagar, J. Arunachalam and S. Gangadharan, J. Anal. At. Spectrom., 13 (1998) 679. E.L. Seibert, V.L. Dressler, D. Pozebon and A.J. Curtius, Spectrochim. Acta B, 56 (2001) 1963. P. Ugo, S. Zampieri, L.M. Moretto and D. Paolucci, Anal. Chim. Acta, 434 (2001) 291. A. Segura-Carretero, J.M. Costa-Ferna´ndez, R. Pereiro and A. Sanz-Medel, Talanta, 49 (1999) 907. R.M. Blanco, M.T. Villanueva, J.E.S. Uria and A. Sanz-Medel, Anal. Chim. Acta, 419 (2000) 137. J. Chwastowska, A. Rogowska, E. Sterlinska and J. Dudek, Talanta, 49 (1999) 837. J.C.A. de Wuilloud, R.G. Wuilloud, R.A. Olsina and L.D. Martinez, J. Anal. At. Spectrom., 17 (2002) 389. C. Fernandez-Rivas, R. Mun˜oz-Olivas and C. Camara, Fresenius J. Anal. Chem., 371 (2001) 1124. S. Ha¨nstrom, C. Briche, H. Emteborg and D.C. Baxter, Analyst, 121 (1996) 1657. E. Kopysc, K. Pyrzynska, S. Garbos and E. Bulska, Anal. Sci., 16 (2000) 1309. J.L. Manzoori, M.H. Sorouraddin and A.M.H. Shabani, J. Anal. At. Spectrom., 13 (1998) 305. J. Qvarnstrom, Q. Tu, W. Frech and C. Lu¨dke, Analyst, 125 (2000) 1193. S. Rio Segade and C. Bendicho, Talanta, 48 (1999) 477. R.P. Mason, K.R. Rolfhus and W.F. Fitzgerald, Mar. Chem., 61 (1998) 37. S. Slaets and F.C. Adams, Anal. Chim. Acta, 414 (2000) 141. P.J. Worsfold, E.P. Achterberg, A.R. Bowie, V. Cannizzaro, S. Charles, J.M. Costa, F. Dubois, R. Pereiro, B. San Vicente, A. Sanz-Medel, R. Vandeloise, E. Vander Donckt, P. Wollast and S. Yunus, J. Autom. Methods Manage. Chem., 24 (2002) 41. P.J. Worsfold, E.P. Achterberg, A.R. Bowie, R.C. Sandford and R.F.C. Mantoura, Chem. Sensors Oceanogr., 1 (2000) 71. P. Fletcher, K.N. Andrew, A.C. Calokerinos, S. Forbes and P.J. Worsfold, Luminescence, 16 (2001) 1. S. Charles, S. Yunus, F. Dubois and E. Vander Donckt, Anal. Chim. Acta, 440 (2001) 37. V. Cannizzaro, A.R. Bowie, A. Sax, E.P. Achterberg and P.J. Worsfold, Analyst, 125 (1999) 51. S. Hirata, M. Aihara, Y. Hashimoto and G.V. Mallika, Fresenius J. Anal. Chem., 355 (1996) 676. Z. Zang, W. Qin and S. Liu, Anal. Chim. Acta, 318 (1995) 71.

The determination of trace elements in water 230 231 232 233 234 235 236 237 238 239 240 241 242 243 244 245 246 247 248 249 250 251 252 253 254 255 256 257 258 259 260 261 262 263 264 265 266 267

A. Economou, A.K. Clark and P.R. Fielden, Anal. Commun., 35 (1998) 389. R. Escobar, Q. Lin, A. Guiraum and F.F. DeLaRosa, Int. J. Environ. Anal. Chem., 61 (1995) 169. E.P. Achterberg, C.B. Braungardt, R.C. Sandford and P.J. Worsfold, Anal. Chim. Acta, 440 (2001) 27. W. Qin, Z.J. Zhang and H.J. Liu, Anal. Chem., 70 (1998) 3579. H. Zamzow, K.H. Coale, K.S. Johnson and C.M. Sakamoto, Anal. Chim. Acta, 377 (1998) 133. A.R. Bowie, E.P. Achterberg, R.F.C. Mantoura and P.J. Worsfold, Anal. Chim. Acta, 361 (1998) 189. J.T.M. de Jong, J. den Das, U. Bathmann, M.H.C. Stoll, G. Kattner, R.F. Nolting and H.J.W. de Baar, Anal. Chim. Acta, 377 (1998) 113. S. Hirata, H. Yoshihara and M. Aihara, Talanta, 49 (1999) 1059. H. Obata, H. Karatani, M. Matsui and E. Nakayama, Mar. Chem., 56 (1997) 97. H. Obata, H. Karatani and E. Nakayama, Anal. Chem., 65 (1993) 1524. W. Qin, Z.J. Zhang and F.C. Wang, Fresenius J. Anal. Chem., 360 (1998) 130. K. Saitoh, T. Hasebe, N. Teshima, M. Kurihara and T. Kawashima, Anal. Chim. Acta, 376 (1998) 247. B. San Vicente de la Riva, J.M. Costa-Ferna´ndez, R. Pereiro and A. Sanz-Medel, Anal. Chim. Acta, 419 (2000) 33. B. San Vicente de la Riva, J.M. Costa-Ferna´ndez, W.J. Jin, R. Pereiro and A. SanzMedel, Anal. Chim. Acta, 455 (2002) 179. B.X. Li, Z.J. Zhang and W. Liu, Anal. Sci., 17 (2001) 1347. K. Okamura, T. Gamo, H. Obata, E. Nakayama, H. Karatani and Y. Nozaki, Anal. Chim. Acta, 377 (1998) 125. J.J. Li, D.X. Du and J.R. Lu, Talanta, 57 (2002) 53. E.P. Achterberg and C.M.G. van den Berg, Mar. Pollut. Bull., 32 (1996) 471. C. Colombo, C.M.G. van den Berg and A. Daniel, Anal. Chim. Acta, 346 (1997) 101. A. Daniel, A.R. Baker and C.M.G. van den Berg, Fresenius J. Anal. Chem., 358 (1997) 703. C.M.G. van den Berg, Sci. Total Environ., 49 (1986) 89. C.M.G. van den Berg, Analyst, 114 (1989) 1527. E.P. Achterberg and C. Braungardt, Anal. Chim. Acta, 400(Special Iss. SI) (1999) 381. M. Kopanica and L. Novotny´, Anal. Chim. Acta, 368 (1998) 211. H. Li and R.B. Smart, Anal. Chim. Acta, 325 (1996) 25. X.L. Wang, J.P. Lei, S.P. Bi, N. Gan and Z.B. Wei, Anal. Chim. Acta, 449 (2001) 35. C.M.G. van den Berg, K. Murphy and J.P. Riley, Anal. Chim. Acta, 186 (1986) 177. E. Shams, Electroanalysis, 13 (2001) 1140. M. Khodari, Electroanalysis, 10 (1998) 1061. A. Merkoci, M. Vasjari, E. Fabregas and S. Alegret, Mikrochim. Acta, 135 (2000) 29. C. Locatelli and G. Torsi, Microchem. J., 65 (2000) 293. C. Colombo and C.M.G. van den Berg, Anal. Chim. Acta, 337 (1997) 29. A.A. Ensaﬁ and K. Zarei, Talanta, 52 (2000) 435. Y. Bonﬁl and E. Kirowa-Eisner, Anal. Chim. Acta, 457 (2002) 285. C.M.G. van den Berg, J. Electroanal. Chem., 215 (1986) 111. M. Vega and C.M.G. van den Berg, Anal. Chem., 69 (1997) 874. C.M.G. van den Berg, Anal. Chim. Acta, 164 (1984) 195. O. Dominguez, S. Sanllorente, M.A. Alonso and M.J. Arcos, Electroanalysis, 13 (2001) 1505.

901

S.N. Willie 268 269 270 271 272 273 274 275 276 277 278 279 280 281 282 283 284 285

286 287 288 289 290 291 292 293 294 295 296 297 298 299 300

902

Y.J. Li and H.B. Xue, Anal. Chim. Acta, 448 (2001) 121. A.P. Aldrich and C.M.G. van den Berg, Electroanalysis, 10 (1998) 369. P.L. Croot and M. Johansson, Electroanalysis, 12 (2000) 565. H. Obata and C.M.G. van den Berg, Anal. Chem., 73 (2001) 2522. B. Pihlar, P. Valenta and H.W. Nurnberg, Fresenius J. Anal. Chem., 307 (1981) 337. J.S. Roitz and K.W. Bruland, Anal. Chim. Acta, 344 (1997) 175. Y.C. Sun, J. Mierzwa and C.R. Lan, Talanta, 52 (2000) 417. C.M.G. van den Berg and G.S. Jacinto, Anal. Chim. Acta, 211 (1988) 129. F. Quentel and M. Filella, Anal. Chim. Acta, 452 (2002) 237. Z. Lukaszewski, W. Zembrzuski and A. Piela, Anal. Chim. Acta, 318 (1996) 159. C.M.G. van den Berg and Z.Q. Huang, Anal. Chim. Acta, 164 (1984) 209. K.S. Rao, M.M. Palrecha and R.G. Dhaneshwar, Electroanalysis, 9 (1997) 804. A.A. Ensaﬁ and B. Naderi, Fresenius J. Anal. Chem., 358 (1997) 480. O.W. Lau and O.M. Cheng, Anal. Chim. Acta, 376 (1998) 197. C.M.G. van den Berg, Talanta, 31 (1984) 1069. R. Klockenkamper and A. vonBohlen, X-Ray Spectrom., 25 (1996) 156. C.L. Babiarz, J.P. Hurley, S.R. Hoffmann, A.W. Andren, M.M. Shafer and D.E. Armstrong, Environ. Sci. Technol., 35 (2001) 4773. B. Holynska, M. Olko, B. Ostachowicz, J. Ostachowicz, D. Wegrzynek, M. Claes, R. Van Grieken, P. de Bokx, P. Kump and M. Necemer, Fresenius J. Anal. Chem., 362 (1998) 294. M.E. McComb and H.D. Gesser, Talanta, 49 (1999) 869. A. Prange, H. Boddeker and K. Kramer, Spectrochim. Acta 48B, (1993) 207. A. Prange and A. Knochel, Anal. Chim. Acta, 172 (1985) 79. B. Holynska, B. Ostachowicz and D. Wegrzynek, Spectrochim. Acta B, 51 (1996) 769. M.A. Barreiros, M.L. Carvalho, M.M. Costa, M.I. Marques and M.T. Ramos, X-Ray Spectrom., 26 (1997) 165. M. Dogan and M. Soylak, J. Trace Microprobe Tech., 20 (2002) 261. E. Haffer, D. Schmidt, P. Freimann and W. Gerwinski, Spectrochim. Acta B, 52 (1997) 935. P. Kump, M. Necemer and M. Veber, X-Ray Spectrom., 26 (1997) 232. V. Mazo-Gray, L. Sbriz and M. Alvarez, X-Ray Spectrom., 26 (1997) 57. H. Miesbauer, Spectrochim. Acta B, 52 (1997) 1003. W. Gerwinski and D. Schmidt, Spectrochim. Acta B, 53 (1998) 1355. A.K. Cheburkin and W. Shotyk, X-Ray Spectrom., 28 (1999) 379. M.M. Costa, M.A. Barreiros, M.L. Carvalho and I. Queralt, X-Ray Spectrom., 28 (1999) 410. S. Griesel, U. Reus and A. Prange, Spectrochim. Acta B, 56 (2001) 2107. L.P. Eksperiandova, A.B. Blank and Y.N. Makarovskaya, X-Ray Spectrom., 31 (2002) 259.

Chapter 27

Aerosol sampling and sample preparation for elemental analysis Jo´zsef Hlavay

27.1

INTRODUCTION

Increasingly strict environmental regulations require the development of new methods for analysis. We also need simple and meaningful tools to obtain information on toxic fractions of different mobility and bioavailability in aerosol samples. Gathering of information on the existence and concentration of substances in the environment, either naturally occurring or of anthropogenic origin, is achieved by measurement of a component of interest. Usually, a single measurement is virtually worthless since temporal and spatial variations cannot be deduced. For a realistic assessment of variations and trends, it is necessary to monitor the concentration of analytes by repeated measurements with sufﬁcient sampling density, both temporally and spatially. An aerosol is a dispersed system consisting of solid and liquid particles suspended in a gas. The gas is the air containing particles produced by different formation processes, such as vapor condensation, combustion or mechanical disintegration of the Earth’s surface. Primary particles are emitted into the atmosphere from sources on the surface, while secondary particles enter the air by gas-to-particle conversion. Aerosol particles formed by gas-to-particle conversion, either in the gas or liquid phase, generally have sizes , 1 mm and are called ﬁne particles. Coarse particles, with a diameter of .1 mm, are produced by surface dispersion. Particles of different origin possess different chemical composition and are of different forms and physical states. Particles can be generated from natural and anthropogenic sources. In general, it is estimated that the annual total amount of particles arising from these sources is about 3000 and 400 million tons, respectively [1]. Particles from natural sources are overwhelmingly coarse particles, arising from wind erosion, sea-spray formation and other similar processes. Anthropogenic emissions, on the other hand, contribute about 60% to the total ﬁne particle mass in the atmosphere. A rough estimate of their contribution to particulate aerosol mass on a global scale is given in Table 27.1. Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

903

J. Hlavay TABLE 27.1 Estimated contributions to the global atmospheric particulate mass (Tg ¼ terra gram) [1] Annual emission or production (Tg/a)

Natural Wind erosion Sea salt Volcanoes Biological primary particles Forest ﬁres Inorganic secondary particlesa Organic secondary particlesb Anthropogenic Direct emissions Inorganic secondary particlesa Organic secondary particlesc

Range

Best estimate

1000–3000 1000–10,000 4 –10,000 26–80 3 –150 100–260 40–200

1500 1300 30 50 20 180 60

50–160 260–460 5 –25

120 330 10

a

Oxidation of sulphur dioxide, reduced sulphur compounds and nitrogen dioxide, uptake of ammonia. b Mainly photochemical formation of PM from isoprene and monoterpenes. c Photochemical formation of PM from anthropogenic emissions of VOCs.

The frequently used method of relating an element in atmospheric aerosols to its source is to calculate enrichment factors (EFs) by employing an indicator element. For crustal aerosols, Al is generally used as the indicator element. The EF can be calculated using the formula: EFcrust ¼

ðCðiÞ =CðAlÞ Þaerosol ðCðiÞ =CðAlÞ Þsoil

ð27:1Þ

Those elements that have EF values between 1 and 10 are usually termed crustal, while elements having EF values in the range of 10 –5000 are generally emitted by anthropogenic sources [2]. 27.1.1 Objectives of monitoring The objectives of monitoring are to assess pollution effects on man and his environment, to identify possible sources and establish relationships between pollutant concentrations [3]. Thus, it is necessary to investigate and understand the mechanisms of transport of trace elements and their complexes to understand their chemical cycles in nature. Concerning natural systems, the mobility, transport and partitioning of trace elements are dependent on the

904

Aerosol sampling and sample preparation for elemental analysis

chemical form of the elements. Major variations of these characteristics are found in time and space due to the dissipation and ﬂux of energy and materials involved in the biogeochemical processes, which determine the speciation of the elements. Chemical species present in a given sample are often not stable enough to be determined individually. Sampling usually consists of two phases: design and implementation. In the design phase, the analyst applies statistical principles to generate a sampling plan. The sampling plan results in a speciﬁcation of the size, number and location of portions to be removed from the lot. The implementation phase involves the physical realization of the design. This phase also requires consideration of the tools of sampling, preservation of portions removed and the precautions necessary to keep the characteristics of the sample unchanged until analysis [4]. The establishment of a nomenclature for sampling is essential for reliable science and simpliﬁes communication. General terms used in sampling are summarized in the Appendix. In many cases, analysts have no responsibility for either the design or the implementation phase of sampling. Knowledge of the sampling error is important because it constitutes a large portion of the total error [5]. Regardless of whether sampling is done by an accredited laboratory, by the customer himself or by a third party, the proﬁt from the analytical results is inherently connected with sampling [6]. Sampling is an error-generating step, often contributing a considerable fraction of the total measurement uncertainty. If one takes the wrong sample, or takes the right sample incorrectly, it is obvious that all data generated are worthless. If the test portion is not representative of the original material, it is not possible to relate the analytical results obtained to the original material, no matter how good the analytical method is, or how carefully the analysis is performed. The result may be dependent on the analytical method, but it always depends on the sampling process. As analytical methodology improves and methods require the use of much smaller test portions, the sampling errors become even more important. Sampling errors cannot be controlled by the use of standards or reference materials. These aspects have been treated in detail by Kratochvil in Chapter 1. In many areas of aerosol analysis, the problems associated with sampling have been addressed, and methods have been validated. Ideally, sampling should be accomplished by, or under the direction of an experienced person, with an understanding of the overall context of the analysis. Whether designing, adapting or following a sampling strategy, there are some important rules to be followed: – –

the analytical work has to be understood and the sampling procedure designed accordingly; equipment used for sampling, subsampling, sample preparation, sample handling and sample extraction has to be selected in order to

905

J. Hlavay

–

avoid unintentional changes in the nature of the sample which may inﬂuence the ﬁnal results. Any critical equipment has to be calibrated; whatever strategy is used, it is of critical importance that the person collecting the samples keeps a clear record of the procedures followed in order that the sampling process can be repeated exactly. Routine sampling procedures must be fully documented.

Once the sample is taken, other interferences can be introduced during transport or preservation. Sometimes, the analytes of interest are unstable and the sampling is often considered to be the weakest link in the chain of planning – sampling – sample preparation –analysis – reporting activities. The purpose and expectation of a sampling program must be realistic and can never surpass the measurement and sample limitations. Costs and beneﬁts must be considered in the design of every measurement program. Sampling contains a degree of uncertainty, and this uncertainty must be considered whenever the data are used [7]. Measurement uncertainty can be controlled and evaluated by an appropriate QA program [7,8]. Sampling devices should be calibrated when factors such as ﬂow rates and size discrimination are important because errors in these factors could cause bias in sample data. Environmental sampling and analysis present numerous opportunities for sample contamination from a wide variety of different sources. Because environmental analysis often deals with very low concentrations of analytes, contamination is an important source of potential error. To minimize error due to contamination, the potential source of contamination must be identiﬁed and eliminated whenever possible. Once the measurement system is established, appropriate types of blanks have to be used to deﬁne background levels of contamination for the different parts of the sampling and analytical processes. By recognizing potential sources of contamination and using blanks to detect changes in background levels, reducing or correcting for contamination is generally possible, and the associated measurement biases can thus be reduced to acceptable levels.

27.2

SAMPLING OF AEROSOLS

27.2.1 General considerations Before sampling, several questions have to be answered in the sampling plan. This is a necessary step for the quality assurance of the environmental analysis. Factors to be taken into account are as follows: Sampler location: It is a key action to place the sampling equipment appropriately to fulﬁll the monitoring requirements. Recommended criteria for the setting of monitoring stations have been developed to cover all situations where human exposures to aerosols could arise (US EPA [9]). A number of

906

Aerosol sampling and sample preparation for elemental analysis

considerations have to be addressed which will affect the aerosol measurement when selecting the location of outdoor monitoring sites: (a) the proximity of point sources, which could result in highly variable concentration gradients; (b) obstructions or changes to air ﬂow caused by tall buildings, trees, etc., and abrupt changes in terrain, which could introduce localized separations; (c) the height of the sampler entry above the ground, which is a compromise between setting the entry of the head height and ensuring that it does not get damaged or receive any extraneous, non-sampled material. Sampling frequency and duration: Ideally to ensure accurate estimates of the exposures to aerosols in the ambient atmosphere, measurements should be continuously taken for the whole period of the study. However, this would be prohibitively expensive, so statistically based sampling programs have to be developed. For long-term network studies, the high-volume gravimetric samplers are usually used that provide a mean concentration over an integrated period of e.g., 24 h. The application of gravimetric samplers in the monitoring system requires the ﬁlters to be changed and weighed for each sample, which is very costly. As such, the development of continuous samplers is necessary. In the shorter term, more speciﬁc studies to determine the exposures from particular operation should be undertaken for long enough to relate any changes in emissions to changes in weather conditions. These effects are usually seasonal in nature, so the program should cover at least a year. Aerosol ﬁlters: Aerosol sampling by ﬁltration is based on the passage of air by a pump through a ﬁlter substrate placed in a suitable ﬁlter holder. The choice of suitable ﬁlters depends upon the sampling and analytical requirements. Fibrous ﬁlters consist of mats of ﬁbers generally made of glass, quartz or cellulose. The most commonly used ﬁlters for sampling aerosols in the ambient atmosphere are glass-ﬁber ﬁlters. These ﬁlters have low moisture retention and high collection efﬁciencies at relatively low-pressure drops. As particles are collected in the depth of the ﬁber bed, glass-ﬁber ﬁlters also have the ability to collect and retain large sample masses with a low rate of back pressure build up. However, if determination of elements or microscopic investigation of collected particles is required, membrane ﬁlters are more suitable. Membrane ﬁlters contain small pores of controlled size, ranging from 0.03 to 8 mm, and they are usually composed of thin ﬁlms of different polymeric materials [10]. For analysis, cellulose nitrate or cellulose triacetate membrane ﬁlters are most widely used as they can be dissolved easily in organic solvents, leaving the collected sample portion for chemical analysis. However, they change their weights due to moisture adsorption, and more suitable membrane ﬁlters are made from PVC, which are widely used for quartz-containing aerosols that can be analyzed by direct on-ﬁlter infrared absorption techniques.

907

J. Hlavay

Application of membrane ﬁlters shows a great advantage in the analysis of samples by microscopy. For optical microscopy, the ﬁlters are made transparent to light by the application of a liquid of the same refractive index as the ﬁlter. Polycarbonate pore ﬁlms (e.g., Nuclepore) are ideal for collecting particles for analysis by electron microscopy. The main problem with all membrane ﬁlters is that, unlike glass-ﬁber ﬁlters, particle collection takes place at the surface of the membrane ﬁlter. This severely limits the amount of sample that can be collected because when more than a single layer of particles is deposited on the surface, the resistance to air ﬂow increases rapidly and there is a tendency for the deposit to be dislodged from the ﬁlter, especially during transport back to the laboratory. By using any type of ﬁlter, in the absence of electric forces, larger and smaller particles are captured from the air and pumped through the ﬁlter material by impaction and diffusion, respectively. The characteristics of different types of ﬁbrous and membrane ﬁlters widely used for sampling aerosol particles for subsequent chemical analysis are summarized in Table 27.2. Filter handling and conditioning are important aspects that are often overlooked in the methodology of sampling, as are the handling and conditioning of the ﬁlters. When taking new ﬁlters from their boxes, it is essential to inspect the state of the surfaces. The surface reactivity is important for chemical analysis, since some ﬁlters react with atmospheric trace gases, resulting in sampling artifacts. The ﬁlters require pretreatment before sampling, which is usually done by acid washing to remove alkaline sites. Teﬂon, quartz and Nuclepore ﬁlters have been found to be the best substrates for chemical analysis. Teﬂon and Nuclepore ﬁlters are used for inorganic substances, while quartz ﬁlters are commonly applied for sampling of organic species. However, in the latter case it should be taken into account that quartz ﬁlters can adsorb organic vapors. This can be checked by using quartz back-up ﬁlter in the air stream to correct the concentration in the aerosol phase determined on the ﬁrst ﬁlter. It is recommended that a second ﬁlter be used after the Teﬂon ﬁlter which does not adsorb organic vapors [11]. Consequently, TABLE 27.2 Properties of ﬁlters used for particulate sampling with a face velocity of 10 cm/s [10]. Note that the efﬁciencies refer to particles having diameters above 0.03 mm Filter

Composition

Density (mg/cm2)

Surface reactivity

Efﬁciency (%)

Teﬂon Whatman 41 Whatman GF/C Gelman Quartz Nuclepore Millipore

Polytetraﬂuoroethene Cellulose ﬁber Glass ﬁber Quartz ﬁber Polycarbonate Cellulose acetate/nitrate

0.5 8.7 5.2 6.5 0.8 5.0

Neutral Neutral Basic (pH 9) pH 7 Neutral Neutral

99 58 99 98 93 99

908

Aerosol sampling and sample preparation for elemental analysis

before sampling, it is necessary to desorb volatile organics from the quartz ﬁlters at high temperature. Aerosol sampling is usually accomplished using ﬁlters having a diameter of several centimeters. The sampling rate is thus typically some m3/d, which gives an amount of aerosol sufﬁcient for the majority of chemical analysis techniques. However, in some measurement programs, large masses of aerosol are needed and a high volume of aerosol is sampled. The ﬂow rate in high volume sampling is about 60 m3/h with a ﬁlter having a diameter around 25 cm or larger. Glass-ﬁber ﬁlters can fulﬁll this requirement. A set of control ﬁlters to take account of changing conditions is highly recommended; one blank for every ﬁve sample ﬁlters is reasonable. 27.2.2 Sampling of aerosol by impactors Aerosol sampling is usually accomplished either by collection devices, like impactors, or by ﬁbrous or membrane ﬁlters. The operational principle of impactors is based on the fact that particles have much larger inertia than gas molecules, which makes their separation in a ﬂuid in motion possible. Since aerosol particles may have different forms and density, impactor data are generally given for aerodynamic particle diameter. Usually, cascade impactors (CIs) are used that collect the particles in different size ranges. An important characteristic of a given impactor stage is the particle diameter (cut-off diameter) where the collection efﬁciency is equal to 50%. The main characteristics of a Berner impactor [12], widely used in recent aerosol studies in Europe and in the United States, are summarized in Table 27.3. The ﬂow rate is 1.9 m3/h at 208C with an exhaust pressure of 150 hPa. The impactor consists of eight stages, while a pre-stage (No. 9) excludes the sampling of particles with diameters .16 mm. Another type of impactor is the Dekati cascade impactor [13]. The Dekati low pressure impactor (DLPI) has 13 successive impactor stages: 10, 6.8, 4.4, TABLE 27.3 Principal characteristics of the Berner-type cascade impactor [12] Stage No.

Cut-off diameter (mm)

Slit diameter (mm)

Number of slits

9 8 7 6 5 4 3 2 1

16 8.0 4.0 2.0 1.0 0.50 0.25 0.125 0.0625

15.9 5.0 2.7 1.2 0.70 0.60 0.42 0.30 0.25

1 8 13 36 53 30 31 63 128

909

J. Hlavay

Fig. 27.1. Dekati DLPI impactor with 13 impactor stages.

2.5, 1.6, 1.0, 0.65, 0.4, 0.26, 0.17, 0.108, 0.060 and 0.030 mm. A picture of the impactor can be seen in Fig. 27.1 [13]. Particle size distribution is deﬁned by measuring the number of particles impacted on the stages of the CI. A DLPI can be used in many applications, including stack measurements, ambient air measurements, automotive applications, educational purposes, etc. The aerosol is collected on 25 mm substrates, which enables chemical analysis to be done on the sample. The particle size range can be extended to , 30 nm with the ﬁlter stage. The impactor is a simple device having two co-linear plates of which one has a small nozzle in it. An aerosol passes through this nozzle at high speed and makes a sharp turn with the ﬂow between the plates. Particles with sufﬁcient inertia cannot follow the ﬂow and impact on the second plate, but particles with small enough inertia remain in the ﬂow. The operational principle of the Dekati DLPI impactor is shown in Fig. 27.2 [13]. CIs can be used for different measuring tasks in chemical engineering as well as for measurements of emissions and environmental pollution. In operation, they often have the advantage of measuring the particle size directly in the gas loaded state, in contrast to other particle measuring techniques. Because of their compact construction, CIs can be directly placed inside the ﬂow channel, avoiding error sources resulting from a partial removal of the gas stream. Generally, the measuring range of a CI lies approximately between 0.5 and 15 mm. As such, it mostly meets the requirements for measurement of emissions in industrial plants. The particle size range of ,1 mm is important for some special measuring tasks and also for the measurement of pollution. For such cases, low-pressure impactors can be used, making a classiﬁcation down to approximately 0.03 mm possible. If the particles

910

Aerosol sampling and sample preparation for elemental analysis

Fig. 27.2. Operational principle of the Dekati DLPI impactor.

are subjected to a chemical analysis after classiﬁcation, high volume CIs with volume ﬂow rates up to 1000 l/min could be used. The Dekati PM-10 impactor is a CI with cut-off points of PM10, PM2.5 and PM1.0 and the ﬁlter stage, so the impactor has three successive stages: 10, 2.5 and 1.0 mm. The PM-10 impactor is available with three different ﬂow rates, i.e., 10, 20 and 30 l/min, which enables the impactor to be used in a wide range of applications. The Dekati gravimetric impactor (DGI) is a CI with cut-off points of 2.5, 1.0, 0.5 and 0.2 mm designed to increase the standard ﬁlter measurements. A back-up ﬁlter after the impaction stages collects all particles ,0.2 mm. The ﬂow rate is 70 l/min. This gravimetric CI can be operated in two modes. As a two-stage sampler, the two lowest impaction stages are replaced with a spacer. In this mode, the ﬁlter stage collects all particles ,1 mm in diameter. Particles are collected on 47 mm diameter substrates, allowing an easy and reliable size distribution and total mass concentration measurement and chemical/elemental analysis for each size fraction. The DGI is designed to ﬁt a standard mass ﬁlter holder so it is easy to add to an existing measurement system. It can also be used in other applications, such as stack sampling and ambient air sampling. The small deposit area impactor (SDI) is a CI in the product group of Aerosol Instruments (Dekati Ltd.). Originally, the SDI was developed in the Finnish Meteorological Institute (FMI) to collect size-fractionated aerosol samples in remote locations for subsequent chemical analysis by proton induced X-ray emission (PIXE) [14]. The SDI-10 impactor classiﬁes particles according to their aerodynamic diameter, from 8.5 mm to 45 nm, with 12 evenly

911

J. Hlavay

distributed impaction stages, and it is available with a ﬂow rate of 11 l/min. The sampling times remain short, even in background areas with low aerosol mass concentrations, because of the relatively high-ﬂow rate. The SDI design enables increased sensitivity for impactor sampling. This has been achieved by optimizing the number of nozzles and the volume ﬂow rate, while keeping the sample deposit diameter small and the size range sufﬁcient for typical aerosol mass size distributions. The small sample size is advantageous if analytical methods, which are sensitive to the deposit area, will be used (PIXE, XRF). The substrate holder is specially designed to ﬁt the PIXE to make the analysis easier. The substrate holder permits contamination free and sensitive chemical analysis also by using other methods. Analysis of inorganic ions or organic acids can be performed by ion chromatography with minimum risk of contamination. An electrical low-pressure impactor (ELPI) [15] is an impactor that provides real-time size and concentration measurements for particles from 0.03 to 10 mm. This unique instrument combines the principles of electrical detection with size classiﬁcation by impaction. The ELPI sensor measures electrical current carried by charged particles at 12 impactor stages using a highly sensitive, multichannel electrometer. Particles can be collected on substrates for microscopic analysis or additional measurements of mass or composition. ELPI systems are robust and, therefore, suited for use in rough environments. They are designed for applications where wide sizerange coverage and fast response are required. This includes combustion aerosol studies, engine emission measurements, ﬁlter testing, indoor/outdoor air quality studies and pharmaceutical research. The micro oriﬁce uniform deposit impactor (MOUDI) [16] impactor samples air at 30 l/min, collecting particles on nine stages. The 50% size cuts for the stages range from 18 down to 0.18 mm aerodynamic particle diameter. Size cuts are 18, 10, 5.6, 3.2, 1.8, 1.0, 0.56, 0.32 and 0.18 mm. The impactor operates for 24–48 h. Particles are impacted onto aluminum foil substrates coated with silicone spray to minimize particle bounce. After operation, the impactor can be disassembled and the aluminum substrates are extracted in water. Ambient atmospheric particles were studied at an ecosystem research site in the Fichtelgebirge Mountains in Central Europe by single-particle analysis and bulk impactor measurements [17]. A ﬁve-stage Berner impactor with polyvinylﬂuoride (Tedlar, DuPont) impaction surfaces was used as a reference sampler for concentration of ammonium, nitrate and sulfate in aerosol particles ranging from 0.05 to 10 mm. Ambient airborne particles were sampled in a 100 m £ 200 m forest clearing surrounded mainly by Norway Spruce. The mobile time-of-ﬂight mass spectrometer (TOF-MS) for on-line analysis of the chemical composition of individual aerosol particles was used. Ambient particles were introduced into the measuring system through a PM10 particle separator with an air ﬂow rate of 16.7 l/min (STP). The sample was introduced

912

Aerosol sampling and sample preparation for elemental analysis

into the TOF-MS through a differentially pumped three-stage inlet module. Particles were analyzed in six different size classes, ranging from 0.2 to 1.5 mm aerodynamic particle diameter. Fuzzy clustering analysis of mass spectra of individual aerosol particles allowed chemical classiﬁcation of the atmospheric aerosol. The chemical composition of aerosols has been determined in 30 sizeresolved samples collected using a Berner low-pressure impactor (BLPI) during two campaigns conducted at a coastal site in the Eastern Mediterranean [18]. The impactor had an inlet providing approximately 15 mm upper cut-off particle size and operated at a ﬂow rate of 25 l/min. The impactor separated the particles into the following aerodynamic equivalent diameter (Dp) ranges: 6.8, 3.39, 1.72, 0.85, 0.44, 0.25, 0.17, 0.11, 0.06 and 0.024 mm. To reduce particle bounce, Nuclepore polycarbonate foils coated with Apiezon L vacuum grease were used. Sampling was accomplished in approximately 24 h intervals. Sulfate (SO4 22) and ammonium (NH4 þ) have been identiﬁed as the main ionic components of the sub-mm aerosol fraction, with SO4 22 accounting for up to 38% of the total ﬁne mass and up to 65% of the total ionic mass during both seasons. On the other hand, NO3 2, Cl2, Naþ and Ca2þ were identiﬁed as the main components of the super-mm mode. The ionic organic compounds (including carboxylic, dicarboxylic and ketoacids) were distributed between both sub-mm and super-mm mode, indicating origin from both gas-to-particle conversion and heterogeneous reactions on pre-existing particles. A ﬁve-stage CI (K-JIST cascade impactor), illustrated in Fig. 27.3, has been designed and tested [19]. The gravimetric method was used to determine the collection efﬁciency by weighing the mass of particles collected on the substrate. In terms of the design of a ﬁve-stage CI with multinozzles, the number and the size of nozzles for each stage were determined using the Reynolds number. The cut-point diameters of stages 1–5 were chosen to be 10, 5, 2.5, 1.0 and 0.7 mm, respectively, with the nominal ﬂow rate of 30 l/min at atmospheric pressure and air temperature of 208C. Experimental data were ﬁtted by non-linear regression using a newly suggested sigmoidal function, and stage response functions were obtained from the collection efﬁciency curves. Calibrated cut-point diameters were well matched with designed values with sufﬁcient stiffness. Results of a source apportionment for the urban aerosol in Erfurt, Germany were presented for the period 1995–1998 [20]. The analysis was based on the data of particle number concentrations, the concentration of the ambient gases SO2, NO, NO2 and CO, particle mass ,2.5 mm (PM2.5) and , 10 mm (PM10) (Harvard impactor sampling, mean PM2.5 26.3 mg/m3, mean PM10 38.2 mg/m3) and the size-fractionated concentrations of 19 elements (impactor sampling 0.05–1.62 mm, PIXE analysis). Crustal EFs for the PIXE elements using Si as reference element, and the diurnal pattern of the measured pollutants on weekdays and on weekends were calculated. The highly correlated PIXE elements Si, Al, Ti and Ca having low EFs were identiﬁed as soil elements.

913

J. Hlavay

Fig. 27.3. Cross-section view of the K-JIST cascade impactor.

Besides NO and particle number concentrations, other pollutants such as NO2, CO, as well as the elements Zn and Cu, were strongly correlated and appear to reﬂect motor vehicle trafﬁc. Highly correlated elements V and Ni had similar EFs and were considered as tracers for oil combustion. The Spearman correlation coefﬁcient, r, was calculated to describe associations between the elemental concentrations. In the study, Zn, Pb and Cu were found to be highly correlated (all the elements had high EFs), an observation that suggested that these elements originated from a common source. Wear of tires and brakes was reported to contribute to the Zn and Cu load in street dust. Factor analysis results of other studies suggested that Ni, V, Zn, Cr and Se originated from oil. A study has been conducted of the concentrations of a range of inorganic elements and polycyclic aromatic hydrocarbons during a winter period at a

914

Aerosol sampling and sample preparation for elemental analysis

roadside location in Birmingham, UK [21]. Samples for trace metal and ion analysis were collected using a MOUDI (MSP Corp., USA) giving cut-offs at approximately 10, 2, 1 and 0.2 mm. Collection substrates were polypropylenebacked 0.5 mm PTFE ﬁlters. The instrument was run at ﬂow rate of 30 l/min. Following sampling, the ﬁlters were cut into two equal portions in a clean air hood; one segment was used for ion chromatographic and ammonium analyses whilst the other was used for the trace metal analyses. Further segments of the PTFE impaction substrates were digested in aqua regia. The digestion process involved addition of the acid mix to samples before heating and ultrasonication. Metals were analyzed by the ICP-MS. The use of sizediscriminated metal concentrations in regression analyses showed a weakening of correlations with NOx and particle counts for Cu and Mo as the size fraction becomes smaller, which is indicative of the trafﬁc-generated component being in the larger re-suspended particles. Exposure to ambient particulate matter (PM) has recently received considerable attention as the result of epidemiological ﬁndings, which showed associations between ambient particulate concentrations and mortality [22]. These associations have been initially demonstrated for TSP and PM10. However, results from other studies suggest that ﬁne particles (PM2.5) and þ particle components, such as sulfate (SO22 4 ) and aerosol strong acidity (H ), may also be associated with increased mortality and other adverse health impacts. The personal cascade impactor sampler (PCIS) is a miniaturized CI, consisting of four impaction stages, followed by an after-ﬁlter (Fig. 27.4). Particles are separated in the following aerodynamic particle diameter ranges: ,0.25, 0.25–0.5, 0.5 –1.0, 1.0 –2.5 and 2.5 –10 mm. The PCIS operates at a ﬂow rate of 9 l/min using a very high efﬁciency, battery-operated light pump at a pressure drop of 2.7 kPa. PTFE (Teﬂon), quartz and aluminum substrates were chosen for characterization of the PCIS. The sampler operates with a state-ofthe-art battery-operated, light-weight, high efﬁciency personal air pump. The entire sampler is enclosed in a cassette holder, 4 cm in diameter and 6 cm high, made of soft aluminum in order to avoid particle losses due to electrostatic deposition. The total weight of the sampler is approximately 150 g; the pump weighs about 450 g (including the battery). Aerosol samples were collected by stacked ﬁlter units and CIs at an urban background site, two downtown sites, and within a road tunnel in ﬁeld campaigns conducted in 1996, 1998 and 1999 in Budapest [23]. Stacked ﬁlter units (Gent-type PM10 SFUs) and CIs were employed for collecting sizefractionated aerosol samples. Two 47-mm-diameter Nuclepore polycarbonate ﬁlters with pore sizes of 8 mm (Apiezon-coated) and 0.4 mm were placed in a NILU-type open face stacked ﬁlter cassette in series. The sampler was operated at an air ﬂow rate of 15 –16 l/min. The aerosol particles were separated into a coarse (about 10–2 mm equivalent aerodynamic diameter, EAD) and a ﬁne (,2 mm EAD) size fraction. The other sampling device was a Battelle-type single-oriﬁce PIXE International cascade impactor operating at a nominal air

915

J. Hlavay

Fig. 27.4. (a– d) The personal cascade impactor sampler (PCIS).

ﬂow rate of 1 l/min. The cut-off diameters for 50% collection efﬁciency of the impaction stages were 16, 8, 4, 2, 1, 0.5 and 0.25 mm EAD. The coarse and ﬁne Nuclepore ﬁlters were weighed before and after the sampling using a microbalance to obtain the particulate mass. Approximately a half section of each Nuclepore ﬁlter was analyzed by INAA. The method was validated using NIST Standard Reference Material 1648: urban PM. A quarter section of each Nuclepore ﬁlter was also analyzed by PIXE for up to 29 elements. The analytical results were used for characterization of the concentration levels, elemental composition, time trends, enrichment of and relationships among the aerosol species in coarse and ﬁne size fractions, for studying their ﬁne-tocoarse concentration ratios, spatial and temporal variability, for determining detailed elemental mass size distributions and for examining the extent of chemical mass closure. 27.2.3 Ambient sampling for the respirable fraction In contrast to sampling methodology in the workplace, very little attention has been paid to sampling the respirable aerosol fraction in the ambient atmosphere. This is probably because the respirable fraction is mainly relevant

916

Aerosol sampling and sample preparation for elemental analysis

to diseases of the deep lung, such as pneumoconiosis, silicosis, asbestosis, etc., which are normally derived from occupational exposures. When formulating a strategy for the health-related sampling of aerosol in ambient atmosphere, the exact application needs to be considered. There are long-term studies at a number of ﬁxed sites to monitor pollution trends, and shorter-term studies designed to address a speciﬁc pollution problem. In these later studies, a large number of samplers are concentrated in a small area around a particular aerosol source, and measurement is normally taken before and after a speciﬁc operation has taken place. Nowadays, many different sampling devices can be found in the market. For example, the DUSTTRAK monitor is factory-calibrated for the respirable fraction of standard (ISO 12103-1, A1) test dust that typiﬁes common ambient aerosols [16]. For ultimate precision, it can be conveniently calibrated in the ﬁeld for the speciﬁc airborne contaminant in a workplace, even liquid aerosol mists. The 10 mm cyclone in each instrument permits isolation of respirable aerosol fractions in accordance with the internationally accepted 4 mm (50%) cut-off criterion. Impactors for 2.5 and 1.0 mm are also included with every instrument. Consistent performance in the dirtiest environments is assured, due to the monitor’s unique sheath air system. Critical lenses are isolated from particle-laden ﬂows by surrounding the sample with ﬁltered air. The new weatherproof environmental enclosure protects the sampler from harsh environmental extremes, indoors or outdoors as well. The effect of inhaled particles depends on where they deposit in the respiratory tract. Exposure sampling must reﬂect the physiology of the human body to be useful. This can only be accomplished through the use of a particle size-selective sampling method that models the human respiratory tract. The RESPICON particle sampler [16] is a multistage virtual impactor (VI) that traps airborne dust onto three individual collection ﬁlters. The mass of deposited dust can be determined by comparing before and after ﬁlter weights. The sampler is a lightweight, compact device for measuring human exposure to airborne particle concentrations in the workplace. It uses a single sampling head to model the human respiratory tract and simultaneously determine the three most critical particle fractions: inhalable, thoracic and respirable. These three size fractions represent the size of particles that can penetrate progressively deeper into the respiratory system. Airborne particles corresponding to the inhalable fraction are drawn into the sampler through a ringgap sampling inlet via a conventional personal sampling pump. Coarse particles pass straight through to the lower collector while other particles are aerodynamically separated onto the appropriate ﬁlter. The ﬁrst VI stage separates out and collects the particles smaller than 4 mm. The second stage collects particles below 10 mm, while the third stage collects the remaining particles, as illustrated in Fig. 27.5. The inherent design advantages of virtual impaction allow the RESPICON to avoid many of the sampling problems

917

J. Hlavay

Fig. 27.5. Conventional curves as agreed upon by ISO, ACGIH, (American Conference of Governmental Industrial Hygienists, threshold limit values for chemical substances and physical agents and biological exposure indices, 1999) and CEN (Comite´ Europe´en de Normalization) for the inhalable, thoracic and respirable fractions (1993/94). Respirable fraction: 50% cut-off at 4 mm (Stage 1) Thoracic fraction: 50% cut-off at 10 mm (Stage 1 þ 2) Inhalable fraction: 50% efﬁciency at 100 mm (Stage 1 þ 2 þ 3) [16].

common to conventional impactors. Particle bounce losses are eliminated and there is no need for messy grease traps. 27.2.4 High-volume aerosol samplers The DIGITEL AUTOMATIC high-volume aerosol sampler (Da-80 H) [24] can be operated fully automatically, permitting proportioned air samples to be taken. The work area of the air volume amounts to 100 –1000 l/min for the standard design (according to 6–60 m3/h). The ﬂoating dust portions of aspirated air are separated with measuring ﬁlters (standard diameter 150/ 140 mm). The choice of the ﬁlter material and the ﬁlter structure (e.g., cellulose, quartz and glass-ﬁber ﬁlter) depends on the goal of the investigation. Schulze [25] developed a fully automated high-volume aerosol sampling device, the ISAPw 2000. The microprocessor controlled aerosol sampling device is designed for the analysis of suspended particles in ambient air, meeting DIN EN 12341/DIN ISO 7708/DIN ISO 8756/EN 481/VDI 2463 and VDI 3786. Deﬁned air volumes are sucked through ﬁlters and the particles are deposited on the ﬁlter material. By means of differential weighing and chemical analysis of the coated ﬁlters, the mass concentrations of the aerosols and their components, such as trace metals, or heavy volatile organic compounds may be determined. The device works with a high precision volumetric ﬂow rate controller with linear ﬂow sensor, scaled by sensing the temperature and the pressure. At size-selective samplings (PM) the temperature, the pressure and the humidity of the ambient air are measured by sensors and sequentially

918

Aerosol sampling and sample preparation for elemental analysis

taken into consideration into the ﬂow rate regulation in order to always ensure the correct cut-point of the impactor system. A comparability of the mass concentration/analyses of different sampling sites is secured by the sequential calculation and accumulation of the dry standard volume. Flow rates up to 230 m3/h may be chosen. The maximum differential pressure of 2230 hPa allows the application of ﬁlter materials with minimized porosity. In order to attain signiﬁcant information regarding the sources of the aerosol, the transport mechanisms in the air and the input mechanisms from the air to bordering media, the device allows an event-induced change of ﬁlters. The designated ﬁlters are changed, depending on a number of relevant criteria, such as wind direction and wind velocity, deposit, humidity, temperature, air pressure and radioactivity, or combinations of these parameters. One ﬁlter depot contains 30 ﬁlters, which may be applied in any required order. The multiple use of ﬁlters with identical sets of sampling parameters ensures that the optimal deposition capacity is reached. Thus, the operating time between the changing of the boxes is maximized, while blank concentration problems during analysis are minimized. As closed units, the ﬁlter magazines may be changed quickly, easily and contamination-free. The aerosol sampling device is designed as a stand-alone system to be applied in rough environments, such as off-shore sampling sites on platforms or research vessels. The modular design of the system allows a broad spectrum of applications and fairly uncomplicated service. After the system is placed and connected to power, the device executes the sampling program automatically. Through the controlling interface, an unnoticed manipulation of the sampling is executed. After a power failure, this is registered by the microprocessor, the device works automatically. The date, time and all data are saved. All device and event parameters are recorded with date and time every 5 min to enable a complete evaluation of the sampling period by read out through the interface, ﬁle saving, report printing function and transfer function for table calculation programs. The optional teleprocessing allows the on-line control of the saved and updated device and event parameters on the host PC at the supervising institute via modem or satellite. Here, the device parameters, such as the ﬂow rate, the observation time, the starting and sampling times of the individual ﬁlters, etc. as well, the combination of event parameters may be altered. For a better understanding of the atmospheric transport of aerosol bound heavy metals to the Arctic and their deposition processes to the snow surface, wet and total deposition sampling, as well as aerosol sampling were performed ˚ lesund [26]. An aerosol sampler was installed on top of the roof (8 m at Ny-A above ground level) of the Japanese research station, which is located 3 km ˚ lesund. This sampler was specially designed for northwest of the center of Ny-A low temperature applications and was equipped with a high-volume pump and an automatic ﬁlter changer for 30 ﬁlters or with 15 single-stage impactors for size-separated aerosol sampling. Multielement analyses of these samples were performed in the home laboratory. Aerosol sampling was controlled by a

919

J. Hlavay

computer program that permitted repeated selection of each ﬁlter. The ﬂow rate was kept constant, although the ﬁlter resistance increased due to the aerosol load. The aerosol sampler was connected to a server PC combined with a ˚ lesund. It was possible to perform on-line control of the modem located in Ny-A device and meteorological parameters; such as ﬂow rate, starting and sampling time of the individual ﬁlters and wind direction, wind speed, air temperature and air pressure, as well as the combination of event parameters via the host PC at the home laboratory. In addition to on-line control, all relevant sampling parameters could be stored on the server PC and transferred by telecommunication to the host PC for further evaluation. The main purpose of the sampling was to obtain speciﬁc information about the origin and spatial distribution patterns of trace elements polluting the atmosphere in the Arctic by using a sampling system that took direction and speed of the local horizontal wind into account. Up to eight sectors could be chosen as sampling sectors on eight ﬁlters. The system assured that each ﬁlter was charged only with aerosol from a selected sampling sector. Situations with low wind speeds and variable wind directions were sampled on two separate ﬁlters. During strong wind situations, the sampler was shut off in order to protect the ﬁlters from drifting snow. It was found that the main fraction of Pb was associated with particles , 2 mm. Glacial ice cores reveal that pollution of the Arctic can be observed to begin in 1912 and has increased signiﬁcantly since 1956 [27]. However, there are relatively few long-term studies of the composition of the Arctic aerosol. The chemical composition of weekly aerosol samples collected between 1964 and 1978 from Finnish Arctic was measured to provide a data set spanning more than 13 years. From October 1964 to February 1978, 685 samples were collected weekly. Rectangular 12 cm £ 14 cm sized Whatman paper ﬁlters (Grade 42) were used and the collection ﬂow rate was about 7 m3/h, giving a total sample volume of 1200 m3. The ﬁlters have been stored at room temperature in an envelope with other ﬁlters from the same year. The weekly 12 cm £ 14 cm ﬁlters were cut into two equal pieces within a laminar ﬂow clean bench. Half of each ﬁlter was retained in the archives, while the other half was cut into two further pieces. One quarter of each ﬁlter was used for black carbon (BC) and IC analysis. NIST Standard Reference Material (SRM) coals 1632a, 1632b, and 1632c, citrus leaves 1572, oyster tissue 1566a and San Joaquin Soil 2709 were analyzed for quality control on the process. Signiﬁcant changes can be seen in the year-to-year variation, resulting from the variation in emission strength and transportation efﬁciency. Recent epidemiological studies and health effect assessments have indicated that the most severe adverse health effects of ambient air pollution (increased daily deaths, hospital admissions of cardiorespiratory patients, shortened life expectancy) are consistently associated with thoracic particles (PM10), and possibly even more strongly with ﬁne particles (PM2.5). Moreover, there have been independent associations of adverse outcomes with ultraﬁne

920

Aerosol sampling and sample preparation for elemental analysis

(PM0.1) particles. A high-volume cascade impactor (HVCI) has been developed at the Harvard School of Public Health (HSPH) for collection of ambient aerosol particles for physicochemical and toxicological characterization studies [28]. The main beneﬁt of the HVCI is its ability to collect large quantities of sizefractionated particle masses in a short time onto a small area of high-capacity collection substrate, i.e., polyurethane foam (PUF). The collection substrate is adhesive free and, therefore, the deposited particle mass can be successfully used in various chemical and biological tests. The performance of three different conﬁgurations of a recently developed HVCI system was tested in both ﬁeld and laboratory experiments. In the ﬁeld, the HVCI was run simultaneously with a virtual impactor and a low-pressure impactor using a 3- or 4day sampling duration. One high volume and two low volume samplers were used for simultaneous collection of particulate mass in different size fractions: three conﬁgurations of the HVCI at 850 l/min, a VI at 16.7 l/min and a BLPI at 25 l/min. PUF was used as the collection substrate in stages 2–4. PM that was not captured by the impaction stages was collected on a backup ﬁlter made of glass ﬁber (Munktell MGA, Grycksbo, Sweden). The laboratory calibration indicated that using stages with 10.5, 2.4, 0.9 and 0.2 mm the ambient air PM10, PM2.5, PM1 and PM0.2 mass fractions could be collected with the HVCI for chemical and toxicological characterization studies. Ambient PM10 were sampled at three sites in an industrialized urban area of Northern Greece and analyzed for 17 chemical elements, 5 water-soluble ions and 13 PAHs [29]. Samples were collected for 24 h every 6th day using high volume PM10 samplers (Graseby-Andersen Ltd., equipped with rectangular ﬁlters 20 £ 25 cm2). Two samplers were used in parallel collecting airborne particles on cellulose-ﬁber ﬁlters for subsequent elemental/ionic component analysis and on glass-ﬁber ﬁlter for PAH analysis. Particulate emissions were collected on glass-ﬁber ﬁlters (5 cm diameter), using low-volume air samplers (IND, Berlin) equipped with PM10 inlets. Two identical air samplers were used in parallel operation for particle collection on cellulose-ﬁber ﬁlters for elemental/ionic component analysis and on glass-ﬁber ﬁlters for PAH analysis. An INAA technique was employed for the determination of As, Br, Co, Fe, Cr, Zn, Sb, La, Sc, Na and K in cellulose-ﬁber ﬁlter and dust samples. Watersoluble ions in source samples were determined by ion chromatography (SO4 22, NO3 2, Cl2, NH4 þ) and ET-AAS (Ca2þ) after sample extraction with deionized distilled water. Blank (including ﬁlters) and duplicate sample analyses were performed for about 10% of all samples according to standard operating procedures. Certiﬁed reference materials (CRM) were used to ensure good accuracy and precision. SOIL-7 (soil dust) and SL-1 (lake sediment), both obtained from the IAEA, were used for elemental analysis by INAA and ETAAS. The chemical mass balance (CMB) source apportionment indicated diesel vehicle exhaust as major source of ambient PM10 at all sites. Signiﬁcant contribution from industrial oil burning was also evidenced at the site located closest to the industrial area.

921

J. Hlavay

PM10 and PM2.5 samples were collected by two high-volume samplers (Graseby-Andersen) at three different sampling locations in Hong Kong and the concentrations of major elements, ions, organic and elemental carbons were quantiﬁed [30]. All ﬁlters were maintained in a condition of 50% RH and 258C for over 48 h and then weighted before sampling. The high-volume samplers were operated at ﬂow rates of 1.13–1.41 m3/min. Both PM10 and PM2.5 were collected on 20.3 cm £ 25.4 cm Whatman quartz microﬁber ﬁlters. Partisol 2000 low-volume sampler with 10 or 2.5 mm inlet was used to collect PM10 and PM2.5 samples, respectively, for data comparison (mass concentrations). The ﬂow rate of Partisol 2000 was 16.7 l/min and samples were collected on a 47 mm quartz ﬁlter. All the quartz ﬁlters were preheated at 9008C for 24 h before being used. After sampling, the loaded high-volume ﬁlters were conditioned and weighed again to determine the mass concentration of the ﬁltered particles. The ﬁlters were digested using a laboratory microwave extraction system to extract metals with an HCl/HNO3 solution (US EPA Method IO 3.1). The solution was ﬁltered and then diluted to 25 ml with distilled –deionized water. Major heavy metals were analyzed by ICP-OES, and trace metals were analyzed by ICP-MS. The deviations of high-volume samplers and Partisol samplers were within 13% for PM10 and within 15% for PM2.5. The PM2.5 levels in urban areas were very high in Hong Kong when compared with other Asian cities. Moreover, the mean PM2.5/PM10 mass ratio was also high due to the relatively higher trafﬁc ﬂow combined with industrial emission. 27.2.5 Speciation aerosol sampling system The aerosol speciation sampler is a portable integrated ambient particulate sampling system [31]. The key element is an integrated sampling cassette with PM2.5 pre-separator, denuder, and ﬁlter holder; mounted in an aspirated solar radiation shield. A sharp cut cyclone (SCC), with a ﬂow of 6.7 l/min, is integrated in every sampling canister to remove particles .2.5 mm aerodynamic diameter. The sample cassette contains the SCC and all the necessary components for excluding particles above 2.5 mm aerodynamic diameters, for removing interfering gases, and for collecting particles including semi-volatile analytes. This cassette is the size of a typical 47 mm ﬁlter holder. It is transported to the laboratory for changing of sampling substrates. With the integrated cassette, every element of the sampler that is contacted by the sampled air stream ahead of the ﬁlter is cleaned with each sample change. The sampling cassettes are designed to accommodate denuders and tandem ﬁlters for sampling of semi-volatile species, and for collection of gases such as nitric acid, ammonia and formic acid. The aerosol speciation sampler provides for the collection of as many as ﬁve samples in parallel. These can be used as follows: (1) Teﬂon ﬁlter for mass and trace metals, (2) Teﬂon or quartz for inorganic ions by ion chromatography, (3) denuded Nylon or impregnated ﬁlter for nitrate, (4) tandem quartz for organic and elemental carbon, with backup

922

Aerosol sampling and sample preparation for elemental analysis

ﬁlter for artifact correction, and (5) denuded carbon-impregnated ﬁlter for semi-volatile organic compounds. All of the cassettes can be operated with either one or two 47 mm ﬁlters, and with or without denuder. For example, a Teﬂon–Nylon ﬁlter pair can be operated behind a nitric acid denuder to give inorganic ions and nitrate in the same cassette. Weekly PM2.5 samples were collected in Shanghai, China at two sites for 1 year [32]. PM2.5 sampling was performed with low-ﬂow rate samplers (LFS, Aerosol Dynamics Inc., Berkeley, CA). The LFS has three parallel PM2.5 sampling inlets. One inlet uses a 47 mm, Gelman PTFE Teﬂon-membrane ﬁlter with a 2 mm pore size for PM2.5 mass concentrations and for elemental analysis. The second inlet uses a denuder to remove acidic gases followed by another Teﬂon membrane ﬁlter and a Nylon ﬁlter (Gelman 47 mm diameter, 1 mm pore size). The third inlet was followed by two quartz ﬁlters (47 mm diameter Gelman quartz-ﬁber ﬁlters) that were used for subsequent determination of organic and elemental carbon. The LFS was well suited to collect integrated weekly samples of PM2.5 at a ﬂow rate of 0.4 l/min through each sampling inlet. The Teﬂon-membrane ﬁlter of the single-ﬁlter PM2.5 cassette was analyzed for mass gravimetrically and for 40 elements (from aluminum to uranium) by XRF. The front quartz ﬁlter of the tandem-ﬁlter PM2.5 cassette was analyzed for organic and elemental carbon (OC and EC) by the thermal/ optical reﬂectance (TOR) method. Weekly PM2.5 mass concentrations ranged from 21 to 147 mg/m3, with annual average concentrations of 57.9 and 61.4 mg/m3 at the two sites, respectively. Ammonium sulfate and nitrate accounted for 41.6% of the PM2.5 mass with sulfate alone accounting for 23.4% of the PM2.5 mass. Carbonaceous material accounted for 41.4% of the PM2.5 mass, with 73% of that mass being organic. Crustal components averaged 9.6% of the PM2.5 mass. Potassium, which was 95% water soluble, accounted for 2.7% of the PM2.5 mass. Twenty-four hour average atmospheric concentrations of PM10 and PM2.5 were measured at a site near downtown Buenos Aires [33]. A MiniVol (Airmetrics, Springﬁeld, OR, USA) portable air sampler equipment was used to collect total suspended particulate (TSP), PM10 or PM2.5. The ﬂow rate was 5 l/min for up to 24 h. Glass-ﬁber ﬁlters were used for the gravimetric and IC determinations, Nucleopore ﬁlters were used for the PIXE, while both types of ﬁlters, glass-ﬁber and Nucleopore, were used for the SEM measurements. Before the exposure, the glass-ﬁber ﬁlters (Whatman, 47 mm diameter) were stored into a desiccator at room temperature for at least 24 h. SEM images were recorded with energy dispersive X-ray analysis (EDX) facility. Nucleopore ﬁlters (polycarbonate membrane kit, of 47 mm diameter) coated with gold and glass-ﬁber ﬁlters were used. The values of PM2.5 concentrations correlated well with the concentrations of CO during the winter period, indicating that direct trafﬁc emissions had an important contribution to PM2.5. The data were less correlated for the case of PM10, indicating that the sources of the coarse fraction were not only trafﬁc emissions, with an important contribution of other

923

J. Hlavay

sources, for example re-suspended material. The PM10 and PM2.5 levels were high compared to typical North American and West European concentrations. The concentration levels of PM2.5 were near or larger than the US EPA alarm value. 27.2.6 Passive samplers A miniature, passive aerosol sampler to estimate long-term average concentrations and size distributions was developed by Wagner and Leith [34– 36]. Particles were collected by gravity, convective diffusion and inertia in a 1.5 cmdiameter passive sampler. Scanning electron microscopy and automated image analysis were then used to count and size all collected particles . 0.1 mm. The measured particle ﬂux and a particle-size dependent deposition velocity model were used to calculate the average concentration and size distribution over the sampling period. A wind tunnel was developed, characterized and used to test the passive sampler. The empirical component of the deposition velocity model was determined as a function of particle size, and precision was assessed using three collocated passive samplers. Field tests were conducted in a wellventilated occupational environment. Measured friction velocities were ,0.4 m/s, a range in which passive sampler performance did not depend on turbulence. Passive sampler results correlated well with those of eight-stage impactors. Discrepancies between the passive samplers and impactors were attributed to the small amount of ﬁne particles present, hygroscopic particles and particle bounce in the impactors. Epidemiological studies have shown a relationship between particulate exposure and community health effects. The causal mechanisms in this relationship are not yet clear, partly due to uncertainties in exposure assessment. The longer sampling times of the passive sampler should improve assessments of long-term mean exposures. The sampler is cheaper and easier to operate than conventional samplers, and thus a larger number of passive samplers can be used. Because the passive sampler is much lighter, smaller and quieter than pump-operated personal samplers, it may yield more representative measurements. The size distributions provided by the passive sampler can be used to determine PM2.5, PM10 or other size-dependent concentrations. 27.3

SEQUENTIAL EXTRACTION SCHEMES FOR AEROSOL SAMPLES

Atmospheric speciation of metals has, until recently, received rather little attention because of the real difﬁculties in measuring even the total concentration of most elements in aerosol samples. Fractionation of trace metals in the atmosphere is somewhat different from those applied to speciation in aqueous media. This arises mainly from two considerations: (i) the mechanism of interaction of the biosphere and the atmosphere, and (ii) the mechanism of

924

Aerosol sampling and sample preparation for elemental analysis

transport in the atmosphere. Several atmospheric parameters are dictated by aerosol particles and human health as well. The life of aquatic and terrestrial ecosystems is also affected by the toxic metal content of particles. Total concentrations of the elements in the atmospheric aerosol particles can indicate the sources of the pollutant, while the chemical fractionation can be used to assess the different deﬁned species present in the aerosol sample. Fractionation is usually performed by a sequence of selective chemical extraction techniques, including the successive removal, or dissolution of these phases and their associated metals. Fractionation has been deﬁned as the process of classiﬁcation of an analyte or a group of analytes from a certain sample according to physical (e.g., size, solubility) or chemical (e.g., bonding, reactivity) properties [37]. Fractionation by size is accomplished by the separation of samples into different particle size fractions, usually during sampling. The concept of chemical leaching is based on the idea that a particular chemical solvent is either selective for a particular phase or selective in its dissolution process. Although a differentiated analysis is advantageous over investigations of bulk chemistry of solids, veriﬁcation studies indicate that there are many problems associated with operational fractionation by partial dissolution techniques. Selectivity for a speciﬁc phase or binding form cannot be expected for most of these procedures. There is no general agreement on the solutions preferred for the various components in the aerosol to be extracted, mostly due to the matrix effect involved in the heterogeneous chemical processes [38]. All factors have to be critically considered when an extracting agent for a speciﬁc investigation is chosen. Important factors include the aim of the study, the type of solid material and the elements of interest. Partial dissolution techniques should include reagents that are selective to only one of the various components signiﬁcant in trace metal binding. Whatever extraction procedure is selected, the validity of selective extraction results primarily depends on the sample collection and preservation prior to analysis. In many cases, it is impossible to determine the large numbers of individual species [39]. Fractionation may simply involve an actual physical separation, e.g., ﬁltration. Although a direct determination of the speciation of an element is often not possible, the available methods can still be applied to get valuable information. An evaluation of the environmental impact of an element can sometimes be done without the determination of the exact species and, notwithstanding, fractions that are only operationally deﬁned. It is also desirable to measure the total concentration of the element in order to verify the mass balance. Sequential extractions have been applied using extractants having increasing extraction capacity, and several schemes have been developed to determine species present in the solid samples. Although initially thought to distinguish some well-deﬁned chemical forms of trace metals [40], they rather address operationally deﬁned fractions [41]. The selectivity of many extractants is inadequate or not sufﬁciently understood, and it is questionable as to whether

925

J. Hlavay

speciﬁc trace metal compounds actually exist and can be selectively removed from multicomponent systems. Due to varying extraction conditions, similar procedures may extract a signiﬁcantly different amount of metals. Concentration, operational pH, liquid/solid ratio and duration of the extraction process considerably affect the selectivity of extractants. The conventional approach of equilibration during a single extraction step is the shaking or stirring of the solid phase/extracting agent mixture. There is no general agreement on the solutions preferred for the extraction of various components in aerosol samples, mostly due to the matrix effects involved in the heterogeneous chemical processes. The aim of the study, the type of solid materials and the elements of interest determine the most appropriate leaching solutions. Partial dissolution techniques should include reagents that are sensitive to only one of the various components signiﬁcant in trace metal binding. In sequential multiple extraction techniques, leaching solutions of various types are successively applied to the sample, each follow-up treatment being more drastic in chemical action or different in nature from the previous one. Selectivity for a speciﬁc phase or binding form cannot be expected for most of these procedures. In practice, some factors may inﬂuence the success in selective leaching of components, including: (i) the chemical properties of the leaching solutions chosen, (ii) experimental parameters, (iii) the sequence of the individual steps, (iv) speciﬁc matrix effects, such as cross-contamination and readsorption, and (v) heterogeneity, as well as physical associations (e.g., coatings) of the various solid fractions. Aerosol fractions arising from sequential extraction schemes can be: 1. Mobile, exchangeable elements (environmentally mobile fraction): this fraction includes the water-soluble and easily exchangeable (non-speciﬁc adsorbed) metals and easily soluble metallo-organic complexes. Most of the recommended protocols seek to ﬁrst displace the exchangeable portion of metals as a separate entity. Chemicals used for this fractionation fall commonly into one of the following groups [42]: * water or highly diluted salt solutions (ionic strength , 0.01 M, e.g., MgCl2); * neutral salt solutions without pH buffer capacity (e.g., CaCl2, NaNO3); * salt solutions with pronounced pH buffer capacity (e.g., NH4OAc at pH 7). 2. Elements bound to carbonates and oxide fractions: to dissolve trace elements bound on carbonates and Fe/Mn oxides, commonly buffer solutions (e.g., NH2OH p HCl/acetic acid) are used. 3. Elements bound to silicates and organic fractions (residual fraction): this fraction mainly contains crystalline bound trace metals and is most commonly dissolved with concentrated acids and special digestion procedures, i.e., strong acid mixtures are applied (HF/HClO4/HNO3) to leach all remaining metals.

926

Aerosol sampling and sample preparation for elemental analysis

Finally, the whole procedure has to be optimized with regard to selectivity, simplicity and reproducibility. Information on the chemical speciation of aerosols indicates the mobility of the elements once the aerosol is mixed directly with natural waters or during scavenging of the aerosol by wet deposition. Atmospheric aerosols have important roles in the biogeochemistry and transportation of trace elements in the atmosphere. The aerosol particles inﬂuence solar radiation transfer, cloud –aerosol interactions, and control the optical, electrical and radioactive properties of the atmosphere. Aerosols sampled within the urban environment exhibit a greater solubility than aerosols having a crustal origin and this should be kept in mind when one interprets the results of sequential leaching. Atmospheric removal occurs by dry deposition of aerosol particles to water, soil, buildings or plants, or by wet deposition of aerosol particles and gases in rain, fog, hail and snow. Wet deposition is a very important removal process for those elements associated with small particles and is predominantly anthropogenic in origin. Approximately 80% of the atmospheric removal of elements such as Pb, Cd, Cu, Ni and Zn, to the ocean takes place by wet deposition, while 40% of that process for Fe and Al is related to dry deposition [43]. This is mostly size-dependent, so size fractionation is an important control on removal processes. Wet deposition provides a mechanism by which the metals in aerosol particles can be solubilized and the pH of rainwater is a major control on metal solubility in precipitation. Rainwater pH is governed by a balance between the concentration of acid and neutralizing species present in solution. Usually, metal solubility increases as pH decreases. Aerosol particles are cycled within the atmosphere through clouds and subjected to repeated wetting and drying cycles before they are removed in precipitation. The atmospheric speciation of metals suffers from several difﬁculties. Measurement of the total concentration of most metals in atmospheric samples is even hampered by the lack of proper quality assurance programs. Speciation techniques for trace metals in the atmosphere are different from those applied to the hydrosphere. Therefore, forms of an element are determined by the mechanism of interaction of the biosphere and the atmosphere, and the mechanism of transport in the atmosphere. For estimation of the elemental budget in the atmosphere, dry and wet deposition rates have to be calculated. The dry deposition rate can be calculated from the results of elemental contents of atmospheric aerosols. Chester et al. [44] suggested a sequential leaching scheme for the characterization of the sources and environmental mobility of trace metals in the marine aerosols. The distribution of elements can be reliably determined in three fractions as environmentally mobile, bound to carbonates and oxides (Fe-, Mn-oxides), and bound to organic matter and silicates (environmentally immobile). Aerosols sampled within the urban environment usually exhibit a greater solubility than aerosols having a crustal origin. Particular attention has to be paid in distinguishing between environmentally mobile and

927

J. Hlavay

environmentally immobile fractions because these represent the two extreme modes by which the metals are bound to the solid matrices. The interaction of trace metals in the aerosols with the other receiving spheres (hydrosphere, lithosphere and biosphere) depends greatly on the solubility of metals under environmental conditions. Chester et al. [45] have demonstrated that aerosol speciation data can be related to the extent of the solubility of an element in an aqueous medium, with metals in the environmentally mobile form being most soluble, and can provide a framework for assessing the reactivity of the elements once they have been deposited at the surface. Lum et al. [46] provided data on the chemical speciation of a number of elements in a pollutant dominated, mainly high temperature generated aerosol, by applying a sequential leaching scheme to samples of an Urban Particulate Matter Standard Reference Material (SRM 1648). The crustal aerosol samples consisted of a higher proportion of the stable fractions for all elements studied. The environmentally mobile material bound to silicate fractions can be interpreted as particle surface/particle matrix associations formed as a result of high temperature anthropogenic processes and low temperature crustal weathering processes. Recently, fractionation by size and chemical bonding on aerosols was studied for the ﬁrst time [47]. The sequential leaching technique has been applied to ﬁlter-collected aerosols in eight particle size ranges for determination of the distribution patterns of several elements. Particular attention was paid to distinguishing between the ﬁne and coarse particle size fractions, and the environmentally mobile and environmentally immobile portions. Cadmium and Cr were determined in small concentration both in ﬁne and coarse fractions, 1.1 and 0.6 ng/m3, respectively. Along with the average concentration of samples collected in 13 months, the ranges proved to be narrow (0.3–8.9 ng/m3 for Cd and 0.5 –4.4 ng/m3 for Cr), which means that the pollution originating from these two compounds was nearly constant. The concentration of Cd in aerosols depends considerably on the location, pollution sources, time, as well as meteorological conditions, and ranges from 1 to 300 ng/m3 in major cities [48]. The chemical bonding of cadmium resulted in its association with the environmentally mobile fractions; (55% in ﬁne and 68% in coarse fractions), and less was found in bound to carbonate/oxides (30 and 20%, respectively) and bound to silicate/organic matter (15 and 12%, respectively). In an earlier study, it was reported that Cd in urban aerosol was found to be almost completely in an exchangeable form [46]. Chromium compounds are generally emitted into the atmosphere from waste incineration, coal combustion, smelting furnaces, kilns and the metal industry. Chromium partitioned evenly into the three fractions in both particle size ranges. High proportions of Cr were identiﬁed in stable fractions in urban PM analyzed by sequential extraction procedures [46]. Different samples of aerosol particles over the North Atlantic, South Atlantic, an area near the equator inﬂuenced by Saharan mineral dust, and the Antarctic Ocean were collected by a six-stage CI and analyzed for Cd, Pb, Tl, Ni,

928

Aerosol sampling and sample preparation for elemental analysis

Cr and Fe [49]. In all the samples, heavy metal content was mostly ,1 ng/m3. The more anthropogenically inﬂuenced aerosol from an industrial area showed higher mean metal concentrations, except for iron and chromium. There are several projects in which the fractionation by particle size has been applied and only the total concentration of elements was determined. Recently, three size fractions of PM, ﬁne particulates (PM2.5), coarse particulates (PM2.5 – 10) and PM10 were measured at eight sampling sites in four large Chinese cities during 1995 and 1996 [50]. Annual means of PM10 concentrations, in which 52–75% were PM2.5 ranged from 68 to 273 mg/m3. Within each city, the urban site had higher annual means of all measured PM size fractions. It was clearly demonstrated that the elements were enriched more in ﬁne particles than in coarse ones. An air quality monitoring program in the Czech Republic provided data for the concentrations of aerosol and gasphase pollutants [51]. Fine particulate matter (PM2.5) was composed mainly of organic carbon and sulfate, with smaller amounts of trace metals. Coarse particle mass concentrations were typically between 10 and 30% of PM2.5 concentrations. The ambient monitoring and the source characterization data were used in receptor modeling calculations. For evaluation of the atmospheric budget and the environmental effects of trace metals on the biosphere, the calculation of the dry and wet depositions is of vast importance. The relative signiﬁcance of the two depositional processes varies between locations and is primarily a function of the rainfall intensity in that area. Wet deposition is a very important removal procedure for those elements associated with small particles and is predominantly anthropogenic in origin. Wet deposition rates are based upon the concentration of trace metals in precipitation samples collected by a wet-only sampler. Using the methods developed for fractionation by chemical bonding, i.e., the three-stage sequential leaching protocols, the dry deposition of aerosols can further be divided and more reliable estimation can be performed. In a recent survey an atmospheric budget was calculated for Lake Balaton, Hungary using a monitoring system that had been in place for the past 3 years [52]. The ratio of Ddry/Dwet was signiﬁcant, in particular, for Pb and Zn and usually higher for the others (V, Cr, Ni, Cu and As). Later, it was determined that, for elements such as Fe, Al and Cu, the ratio of the two depositions gives a contrasting effect; namely, dry deposition plays a more important role in the pollution of the environment [53]. On the other hand, ratios of Ddry/Dwet indicated that elements like Mn, As, Cd, Pb and mainly Zn, were entirely deposited in much higher amount by wet deposition than dry one. The environmentally mobile fractions of total dry deposition of elements were also calculated and compared to the total wet deposition. It was obvious that the contribution of mobile fractions of the dry deposition to pollution was, except for Cu, minor. Copper compounds were mainly removed from the atmosphere by dry deposition and only one-third of the amount of Cu was environmentally mobile. In the case of elements such as Al, Fe, Cu, dry deposition has been

929

J. Hlavay

found to be the major source of pollution, but the mobile fraction of dry deposition plays a minor role compared to the total soluble deposition. Further, the soluble fractions of depositions (Ddry mobile þ Dwet) were compared to the total depositions (Ddry þ Dwet). The water quality of the lake has been inﬂuenced by the soluble part of atmospheric depositions, and it has been found that 85 –94% of toxic elements (Pb, Cd, Ni, Zn and As) were dissolved in the water. The other portions of elements were stable compounds formed under natural environmental conditions and, after precipitation, they settle to the bottom of the lake. Thus, the metal compounds, sooner or later, become part of the bottom sediment, since the fate of dissolved metals greatly depends on the physical and chemical conditions of the bulk water (pH, complex forming capacity, adsorption on clays, quartz, organic matter, biological activities, etc.). There are huge amounts of literature on speciﬁc research areas in which appropriate leaching protocols can be found for a given problem. Reviews exist on trace metal speciation in aerosols [43] in the book by Ure and Davidson [54]. 27.4

DISCUSSION

Despite all limitations, sequential extraction schemes can provide a valuable tool to distinguish among trace metal fractions of different particle size and solubility. These fractions are empirically related to mobility classes in aerosol matrices. The speciation of metals governs their availability to the biota and their potential to contaminate the environment. Available forms of metals are not necessarily associated with one particular chemical species or a speciﬁc aerosol sample component. Soluble and exchangeable forms of metals will decrease with time if there are other solid components present that can adsorb the metal more strongly and have free sites that are accessible (e.g., hydrous oxide, organic matter). The present state of knowledge on aerosol fractionation of trace elements is still somewhat unsatisfactory because the appropriate techniques are only operationally deﬁned and associated with practical problems. With respect to estimating the bioavailability of element concentrations, one problem is the effect of competition between binding sites on the solid substrate and selective mechanisms of metal translocation by the different organisms involved. This situation cannot yet be improved by more sophisticated analytical approaches to fractionation. On the other hand, the usefulness of a differentiated approach to the interactive processes between spheres, even if only operationally deﬁned, has been clearly demonstrated. The method of sequential chemical extraction is the least sophisticated and most convenient technique available for fractionation assessment. However, we must be certain that we fully understand what is happening during extraction to minimize the risk of producing artifacts and choose standard procedures to ensure that results are comparable. The primary importance of proper sampling protocols has been emphasized,

930

Aerosol sampling and sample preparation for elemental analysis

since the sampling error can cause erroneous results even using highly sophisticated analytical methods and instruments. The number of fractionation steps required depends on the purposes of the study. Geoscientists and environmental engineers extensively use results of chemical fractionation analysis and scientists have the responsibility to show the pitfalls and limitations of sequential extraction procedures developed. Declaration of the uncertainty of results is a must and greatly improves the quality of these activities.

Acknowledgements The ﬁnancial support of the Hungarian Science Foundation (OTKA TS 40903 and T 043220) is greatly acknowledged. ´ DY, T. LENGYEL AND A.M. URE, COMPENDIUM OF APPENDIX. J. INCZE ANALYTICAL NOMENCLATURE, DEFINITIVE RULES 1997, IUPAC, BLACKWELL SCIENCE GENERAL TERMS USED IN SAMPLING Sample: A portion of material selected from a larger quantity of material. Sample handling: This refers to the manipulation to which samples are exposed during the sampling process, from the selection from the original material through to the disposal of all samples and test portions. Sampling plan: A predetermined procedure for the selection, withdrawal, preservation, transportation and preparation of the portions to be removed from a population as samples. Homogeneity, heterogeneity: The degree to which a property or a constituent is uniformly distributed throughout a quantity of material. (a) a material may be homogeneous with respect to one analyte or property but heterogeneous with respect to another. (b) the degree of heterogeneity is the determining factor of sampling error. Sampling error: That part of the total error (the estimate from a sample minus the population value) associated with using only a fraction of the population and extrapolating to the whole, as distinct from analytical or test error. It arises from a lack of homogeneity in the parent population. (a) in chemical analysis, the ﬁnal test results reﬂect the value only as it exists in the test portion. It is usually assumed that no sampling error is introduced in preparing the test sample from the laboratory sample. Therefore, the sampling error is usually associated exclusively with the variability of the laboratory sample.

931

J. Hlavay

(b) sampling error is determined by replication of the laboratory samples and their multiple analyses. Since sampling error is always associated with analytical error, it must be isolated by the statistical procedure of analysis of variance. Lot: A quantity of material that is assumed to be a single population for sampling purposes. Subsample: This refers to a portion of the sample obtained by selection or division; an individual unit of the lot taken as part of the sample or; the ﬁnal unit of multistage sampling. Sample preparation: This describes the procedures followed to select the test portion from the sample (or subsample) and includes: in-laboratory processing, mixing, reducing, coning and quartering, rifﬂing, and milling and grinding. Laboratory sample: The sample or subsample(s) sent to or received by the laboratory. Test sample/analytical sample: The sample, prepared from the laboratory sample, from which test portions are removed for testing or for analysis. Test portion/analytical portion: The quantity of material, of proper size for measurement of the concentration or other property of interest, removed from the test sample. (a) the test portion may be taken from the primary sample or from the laboratory sample directly if no preparation of sample is required, but usually it is taken from the prepared test sample. (b) a unit or increment of proper homogeneity, size and ﬁneness, needing no further preparation, may be a test portion. Test solution/analytical solution: The solution prepared by dissolving, with or without reaction, the test portion in a liquid. Treated solution: The test solution that has been subjected to reaction or separation procedures prior to measurement of some property. Aliquot: A known amount of a homogeneous material, assumed to be taken with negligible sampling error. The term is usually applied to ﬂuids. When a laboratory sample or a test sample is “aliquoted” or otherwise subdivided, the portions have been called split samples.

REFERENCES 1 2 3 4

932

W.C. Hinds, Aerosol Technology. Properties, Behavior, and Measurement of Airborne Particles. Wiley, New York, 1999. R. Chester, F.J. Lin and K.J.T. Murphy, Environ. Technol. Lett., 10 (1989) 887. C.N. Hewitt and R.M. Harrison, Monitoring. In: R.E. Hester (Ed.), Understanding our Environment. The Royal Society of Chemistry, London, 1986, pp. 1– 10, Ch. 1. W. Horwitz, Pure Appl. Chem., 62(6) (1990) 1193–1208.

Aerosol sampling and sample preparation for elemental analysis 5 6 7

8 9

10

11 12 13 14

15 16 17 18

19 20 21 22 23 24 25 26

27 28 29

R.E. Majors, LC–GC, 10(7) (1992) 500– 506. W. Wegscheider, In: H. Gu¨nzler (Ed.), Accreditation and Quality Assurance in Analytical Chemistry. Springer, Berlin, 1996, pp. 95–103. J.K. Taylor, Deﬁning the accuracy, precision, and conﬁdence limits of sample data. In: L.H. Keith (Ed.), Principles of Environmental Sampling, ACS Professional Reference Book, American Chemical Society, Washington, DC, 1988. J.K. Taylor, Anal. Chem., 53 (1981) 1588A– 1596A. US Environmental Protection Agency (US EPA), Site selection for the monitoring of petrochemical air pollutants, EPA-450/3-78-013. OAQPS, Research Triangle Park, NC. J.M. Waldman, J.W. Munger and D.J. Jacob, In: M. Radojevic and R.M. Hamson (Eds.), Measurement Methods for Atmospheric Acidity and Acid Deposition, Atmospheric Acidity, Sources, Consequences and Abatement. Elsevier Appl. Sci., London and New York, 1992, pp. 205 –243. S.R. McDow and J.J. Huntzicker, Atmos. Environ., 24A (1990) 2563. A. Berner, Design principles of the AERAS low pressure impactor. In: B.Y.H. Liu, D.Y.H. Piu and H.J. Fissan (Eds.), Aerosols. Elsevier, Amsterdam, 1984. DEKATI Measurements Oy, Osuusmyllynkatu 1333700 Tampere, Finland. W. Maenhaut, R. Hillamo and T. Ma¨kela¨, A new cascade impactor for aerosol sampling with subsequent PIXE analysis, Nuclear Instrum. Methods Phys. Res. B, 109/110 (1996) 482 –487. TSI Incorporated http://www.tsi.com ELPI. MSP Corporation, 5910 Rice Creek Parkway, Ste 300 Shoreview, MN, USA. A. Held, K.P. Hinz, A. Trimborn, B. Spengler and O. Klemm, J. Aerosol Sci., 33(4) (2002) 581–594. H. Bardouki, H. Liakakou, C. Economou, J. Sciare, J. Smolı´k, V. Zdı´mal, K. Eleftheriadis, M. Lazaridis, C. Dye and N. Mihalopoulos, Atmos. Environ., 37(2) (2003) 195–208. S.B. Kwon, K.S. Lim, J.S. Jung, G.N. Bae and K.W. Lee, J. Aerosol Sci., 34(3) (2003) 289 –300. J. Cyrys, M. Sto¨lzel, J. Heinrich, W.G. Kreyling, N. Menzel, K. Wittmaack, T. Tuch and H.E. Wichmann, Sci. Total Environ., 305(1– 3) (2003) 143 –156. R.M. Harrison, R. Tilling, M.S.C. Romero, S. Harrad and K. Jarvis, Atmos. Environ., 37(17) (2003) 2391– 2402. C. Misra, M. Singh, S. Shen, C. Sioutas and P.M. Hall, J. Aerosol Sci., 33(7) (2002) 1027– 1047. ´ . Zemple´n-Papp and G. Za´ray, Atmos. Environ., 35(25) I. Salma, W. Maenhaut, E (2001) 4367– 4378. http://[email protected] Schulze Automation Engineering Ingenieurbu¨ro Schulze, Im Heidewinkel 64-66, D21271 Asendorf, Germany, 2003. M. Kriews and O. Schrems, Untersuchungen zur Schwermetalldeposition im Schnee und Eis der Arktis Beitrag zur 18. Internationalen Polartagung, Potsdam, 1996, pp. 18.3–22.3. T. Yli-Tuomi, L. Venditte, P.K. Hopke, M.S. Basunia, S. Landsberger, Y. Viisanen and J. Paatero, Atmos. Environ., 37(17) (2003) 2355–2364. M. Sillanpa¨a¨, R. Hillamo, T. Ma¨kela¨, A.S. Pennanen and R.O. Salonen, J. Aerosol Sci., 34(4) (2003) 485 –500. C. Samara, Th. Kouimtzis, R. Tsitouridou, G. Kanias and V. Simeonov, Atmos. Environ., 37(1) (2003) 41–54.

933

J. Hlavay 30 31 32 33 34 35 36 37 38 39

40 41 42 43

44 45 46 47 48 49 50 51 52 53 54

934

K.F. Ho, S.C. Lee, C.K. Chan, J.C. Yu, J.C. Chow and X.H. Yao, Atmos. Environ., 37(1) (2003) 31– 39. MetOneInstruments, Inc. Corporate: 1600 Washington Blvd., Grants Pass, OR 97526. B. Ye, X. Ji, H. Yang, X. Yao, C.K. Chan, S.H. Cadle, T. Chan and P.A. Mulawa, Atmos. Environ., 37(4) (2003) 499–510. H. Bogo, M. Otero, P. Castro, M.J. Ozafra´n, A. Kreiner, E.J. Calvo and R.M. Negri, Atmos. Environ., 37(8) (2003) 1135–1147. J. Wagner and D. Leith, Passive aerosol sampler. I: Principle of operation, Aerosol Sci. Technol., 34(2) (2001) 186 –192. J. Wagner and D. Leith, Passive aerosol sampler. II: Wind tunnel experiments, Aerosol Sci. Technol., 34(2) (2001) 193– 201. J. Wagner and D. Leith, Passive Aerosol Sampler and Methods, US Patent 6,321, 608, granted 27 November 2001. D. Templeton, F. Ariese, R. Cornelis, H. Muntau, H.P. van Leeuven, L.-G. Danielsson and R. Lobinski, Pure Appl. Chem., 72 (2000) 1453. J.M. Martin, P. Nirel and A.J. Thomas, Mar. Chem., 22 (1987) 313. J. Bufﬂe, K.J. Wikinson, M.L. Tercier and N. Parthasarathy, in: F. Palmisano, L. Sabbatini and P.G. Zambonin (Eds.), Reviews on Analytical Chemistry, Euroanalysis IX. Societa´ Chimica Italiana, 1997, pp. 67–82. G. Sposito, L.J. Lund and A.C. Chang, Soil Sci. Soc. Am. J., 46 (1982) 260. H. Zeien and G.W. Bru¨mmer, Mitt. Dtsch. Bodenkundl. Ges., 59 (1989) 505. P.H.T. Beckett, Adv. Soil Sci., 9 (1989) 143. L.J. Spokes and T.D. Jickells, Speciation of metals in the atmosphere. In: A.M. Ure and C.M. Davidson (Eds.), Chemical Speciation in the Environment. Blackie Academic & Professional, London, 1995, pp. 137 –168, Ch. 6. R. Chester, F.J. Lin and K.J.T. Murphy, Environ. Technol. Lett., 10 (1989) 887. R. Chester, K.J.T. Murphy, J. Towner and A. Thomas, Chem. Geol., 54 (1986) 1. K.R. Lum, J.S. Betteridge and R.R. Macdonald, Environ. Technol. Lett., 3 (1982) 57. ´ . Molna´r and E. Me´sza´ros, Analyst, 123 (1998) 859. J. Hlavay, K. Polya´k, A J.E. Ferguson and D.E. Ryan, Sci. Total Environ., 34 (1984) 101 –116. N. Radlein and K.G. Heumann, Fresenius J. Anal. Chem., 352 (1995) 748–755. F. Wei, E. Teng, G. Wu, W. Hu, W.E. Wilson, R.S. Chapman, J.C. Pau and J. Zhang, Environ. Sci. Technol., 33 (1999) 4188. J.P. Pinto, R.K. Stevens, R.D. Willis, R. Kellogg, Y. Mamane, J. Novak, J. Santroch, I. Benes, J. Lenicek and V. Bures, Environ. Sci. Technol., 32 (1998) 843. M. Bikkes, K. Polya´k and J. Hlavay, J. Anal. At. Spectrom., 16(1) (2001) 74– 81. J. Hlavay, K. Polya´k and M. Weisz, J. Environ. Monit., 3 (2001) 74. A.M. Ure and C.M. Davidson, Chemical Speciation in the Environment. Blackie Academic & Professional, London, 1995.

Chapter 28

Sample preparation for industrial waste analysis Peter Drouin and Ray E. Clement

28.1

TYPES OF INDUSTRIAL WASTE

The major types of industrial waste products generated, by total mass, fall under the general categories of sludge, general construction waste, slag, plastics and glass scraps. Wastes that may result in high concentrations of metals may be expected from speciﬁc industries, such as metal plating, battery manufacture, mining and mineral, and municipal waste incineration, among others. Whether any of the waste products from industrial activity may be deemed hazardous depends not only on the speciﬁc chemicals present, but also on whether these chemicals can be leached from the waste products into the environment. The distinction of an industrial waste as being hazardous or not is an important consideration for its treatment and disposal, and will be speciﬁcally discussed here with respect to sampling and analytical approaches. Although there is no universal deﬁnition of exactly what a hazardous waste is, there has certainly been considerable regulatory action on this topic worldwide. Additionally, waste products can be deemed hazardous because of the presence of metals or other inorganic chemicals, organic chemicals, or a mixture of both organics and inorganics. By any deﬁnition of hazardous, a waste product would have to exhibit one or more of the following properties: ignitability, corrosivity, reactivity, or toxicity. As the topic of this book does not include organic materials, this chapter will only consider sample preparation for determination of metals in waste products. Therefore, only hazardous wastes that may contain potentially toxic high concentrations of metals are considered. Almost all such wastes are the result of anthropogenic activities, either the direct result of industrial processes or waste products deposited into landﬁlls or illegally dumped into the environment. From the above, one may expect that industrial wastes, whether hazardous or not, are expected to have generally greater concentrations of metals than other typical environmental samples, and therefore trace metal analysis is not of concern for this application. This is not true, because industrial processes Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

935

P. Drouin and R. Clement

often require large quantities of water, which may become contaminated at trace levels by leaching metals from the industrial waste originally generated. As sample preparation for the analysis of water has been covered by Willie in Chapter 26 of this volume, it is only brieﬂy discussed here, with one exception: the toxicity characteristic leaching procedure (TCLP). TCLP has become an important test in the regulatory determination of whether a waste is classiﬁed as hazardous. Basically, if metals or other toxic components of a waste are able to leach into the environment in sufﬁcient concentrations to potentially cause toxic effects, then the original waste product is deemed hazardous, and must be treated accordingly.

28.2

SAFETY CONSIDERATIONS FOR INDUSTRIAL WASTE ANALYSIS

Because of the uncertainty associated with the composition of any waste product, sampling personnel and analysts who handle such samples should consider the adoption of universal safety precautions (i.e. samples are assumed to be hazardous). Even if such samples are eventually shown to be safe, in terms of the concentrations of a few selected analytes tested, hazardous concentrations of sample components not tested for may still be present. For example, an industrial waste sample shown to contain only traces of toxic metals may still contain high concentrations of PCBs, chlorinated dioxins, or other regulated organics.

28.3

SAMPLE CHARACTERISTICS AND INDUSTRIAL WASTE SAMPLING

In Chapter 1, Kratochvil describes the principles required to ensure that samples brought back to a laboratory for analysis are representative. The concept of representativeness must be modiﬁed to include the practical consideration of ﬁt-for-purpose. In the case of industrial waste samples, the original source material composition is often inhomogeneous, and may consist of mixed solids/liquids/sludges, and the bulk material may be highly stratiﬁed. Therefore, to obtain a truly representative sample may require an inordinate amount of work. However, it may be possible to obtain samples that are sufﬁciently representative to achieve the objectives of a speciﬁc study, even if the ideal is not reached. These objectives should be clearly understood before any industrial waste sampling is undertaken. The variety of industrial waste samples possible is huge, and may consist of any of the following: liquid waste, soil or non-liquid waste, containerized solids, truckloads of waste, containerized liquids, various types of sludge, waste piles, landﬁll sites, open ﬁelds, sediments, buried drums, and others. Depending on the speciﬁc application, any of the following considerations may be important:

936

Sample preparation for industrial waste analysis

1. Mixed solid/liquid wastes. A decision must be made as to whether it is necessary for the sample to proportionally represent the solid/liquid ratio of the original source material. Liquid and solid components can be collected separately, but it may be difﬁcult to accurately relate results to the industrial waste under study. 2. Stratiﬁed waste piles. Because of the inherent inhomogeneity of such sample types, sampling of the upper, middle, and lower portions of the pile must be attempted. If good precision and accuracy of analytical results are critical, several replicate transects of the waste pile must be taken and analyzed. Specialized equipment may be needed to bore into the waste pile, and special care must be taken to protect individual sample components from gross contamination or cross-contamination from other sample components. 3. Sampler bias and record keeping. There can be considerable sampler bias involved in industrial waste sampling, because more judgment than usual may be required in deciding where to take a limited number of samples. For example, in sampling a ﬁeld where industrial waste had been historically dumped, isolated areas where dark patches are observed may be preferentially selected because of their “contaminated look”. Without prior knowledge of the situation, however, such assumptions may lead to results that signiﬁcantly under- or overestimate the degree of contamination. Because of these considerations, it is especially important for sampling personnel to keep accurate records of ﬁeld observations and reasons why speciﬁc sampling locations were chosen, especially in cases where personal judgment was allowed. With the widespread availability of high quality, relatively inexpensive digital cameras, it is now advisable for personnel to include digital photos of sampling areas as part of the ﬁeld record. 4. Composite sampling/composite analysis. Most of the time, it is advisable to collect more, rather than fewer, samples. Not all samples collected necessarily have to be analyzed, but it is not always possible to obtain additional samples at a later time. Also, to obtain multiple samples at various time intervals introduces additional variables that may complicate data interpretation. To estimate average contamination over a large area with a minimum number of analyses, many samples can be collected, and a few composite samples prepared by mixing aliquots of the original samples collected. Of course, the representativeness of the individual aliquots may be questionable, and any additional sample handling may introduce a greater chance of either sample contamination or gross errors, such as mislabeling samples. Nevertheless, analyses of a relatively few composited samples may be sufﬁcient to satisfy the objectives of a study, depending on the original objectives. If the composited sample analyses do not provide sufﬁcient information, there is still a good chance that the results will at least narrow the selection of a second round of samples to a speciﬁc sub-group or area for more detailed study.

937

P. Drouin and R. Clement

5. Cost. If cost were not an issue, accurate results would be possible for almost any industrial waste application (within any inherent bias of the measurement system), by simply analyzing as many samples as required. Cost is not only an important consideration of an overall sampling and analysis plan, it sometimes is, unfortunately, the overriding consideration. This is why the objectives of any study or investigation must be clear before sampling is initiated. Then a sampling and analysis plan that is ﬁt-for-purpose can be developed, and its associated costs calculated. 28.4

DIGESTIONS

A number of different digestion procedures have been developed to prepare samples for analysis by ﬂame atomic absorption spectroscopy (F-AAS), graphite furnace AAS (GF-AAS), hydride generation AAS (HG-AAS), cold vapor AAS (CV-AAS), inductively coupled plasma-optical emission spectrometry (ICP-OES), and inductively coupled plasma-mass spectrometry (ICP-MS). Digestions typically involve the use of concentrated acids (nitric acid, HCl, perchloric acid, and hydrogen peroxide) and elevated temperature (and pressure for methods utilizing microwaves) to extract metals from the sample. Alkaline digestions have been developed to selectively dissolve hexavalent chromium from solid sample types. One of the most common digestions employed uses aqua regia, which is a mixture of hydrochloric and nitric acids in a 3:1 ratio. The combination of the two acids results in the release of chlorine gas, a powerful oxidizing agent. 3HCl þ HNO3 ! 2H2 O þ NOCl þ Cl2

ð28:1Þ

Not only does this acid mixture have strong oxidizing properties, but also the chloride ions help to stabilize certain elements in solution (Fe, Sb, and Sn). Silver will precipitate from solutions containing low concentrations of chloride ions as AgCl, which will cause poor recoveries. However, in the presence of high level of chloride ions, an anionic complex is formed, resulting in acceptable recoveries [1 –4]. Agþ þ Cl2 ! AgClðsÞ þ Cl2 ! AgCl2 2 ðaq

28:2Þ

Unfortunately, the presence of high concentrations of chloride ions can pose difﬁculties for certain instrumental methods of analysis. The presence of high chloride levels can affect the determination of certain elements when employing ICP-MS due to isobaric interferences, such as the ArClþ ðm=z ¼ 75Þ interference on arsenic ðm=z ¼ 75Þ: Additionally, when using GF-AAS methods, high chloride concentrations can result in analyte loss during the pyrolysis stage, gas phase interference, and signiﬁcant reductions in the lifetime of the graphite tube due to corrosion [1,2,5,6]. Although work has shown that the use of ruthenium as a permanent modiﬁer can reduce these problems in GF-AAS

938

Sample preparation for industrial waste analysis

[5], nitric acid or nitric acid/hydrogen peroxide mixtures are often used to digest samples for GF-AAS or ICP-MS determinations. Although these digestions are aggressive, the sample is typically not completely dissolved. Often the results from these digestions are erroneously reported as “total” metal concentrations. The concentrations of certain elements may approach the true total metal concentrations, but the recovery efﬁciency of target metals from an industrial waste sample depends on the acids employed in the digestion, and the mineralogical composition of each speciﬁc sample. To maximize metal recoveries, the acid mixture used for sample digestion must include hydroﬂuoric acid (HF), to liberate metals bound in silicate matrix. Even the use of digestions with HF may not completely recover all minerals present in a sample [6– 9]. Alternatively, a fusion procedure (i.e. lithium metaborate/tetraborate or sodium peroxide) or the use of alternative instrumental techniques (i.e. neutron activation or X-ray ﬂuorescence) may be employed if total metal concentrations are required. Generally, the analysis of industrial wastes does not require determination of total metal content because metals bound in the silicate matrix are not considered to be environmentally available. The US EPA Ofﬁce of Solid Waste publication SW-846: Test Methods for Evaluating Solid Waste, Physical/ Chemical Methods, describes analytical methods for both inorganic and organic parameters [10– 18]. A summary of reported digestion methods for inorganic parameters is given in Table 28.1. Methods for the sample digestion of industrial wastes for determination of metals are generally grouped into two main categories, open vessel hot plate/block digestions (convection heating) and closed vessel microwave digestions (microwave heating). Closed vessel microwave digestions are conducted at higher pressures and temperatures, which result in shorter digestion times (minutes versus hours). In addition, the use of closed vessels minimizes the loss of volatile elements and reduces the potential for sample contamination [1 –4,19–21]. Feedback loops designed as part of microwave systems allow for accurate and repeatable temperature control, resulting in improved analytical precision [1,2,22]. Microwave digestion vessels constructed of ﬂuorocarbon polymers such as perﬂuoralkoxy (PFA) Teﬂon and teraﬂuoroethylene/perﬂuoro(propyl vinyl ether) copolymer (TFM) are semipermeable to water, carbon dioxide, and acid gases (HCl and NOCl) that can be released with subsequent heating [23]. This can have signiﬁcant analytical implications on the determination of Ag. For example, a nitric acid-only digestion performed after an aqua regia digestion would result in low Ag recovery due to the precipitation of AgCl. 28.4.1 Aqueous sample types—US EPA methods Although many methods have been reported worldwide for preparing aqueous samples for determination of metals, the inherent differences between many of

939

940

TABLE 28.1 Summary of US EPA digestion methods for metals in various sample types [10–18] Matrix

Digestion equipment

Acids

Analytical instrumentation

Elements

3005

Surface and ground water

Hot plate

HNO3/HCl

F-AAS, GF-AAS, ICP-OES

3010A

Extracts, wastes

Hot plate

HNO3/HCl

F-AAS, ICP-OES

3015A

Extracts, surface and ground water

Microwave

HNO3

F-AAS, GF-AAS, ICP-OES

3020A 3031

Extracts, water Oils

Microwave Hot plate

HNO3 HNO3/H2SO4/KMnO4

3050B

Soils, sediments, sludges

Hot plate

HNO3/HCl/H2O2

3051A

Soils, sediments, sludges, oils

Microwave

HNO3/HCl

3052

Siliceous, organic

Microwave

HNO3/HCl, H2O2/HF

3060A

Soils, sediments, sludges

Hot plate

NaOH/Na2CO3

GF-AAS F-AAS, GF-AAS, ICP-OES F-AAS, GF-AAS, ICP-OES, ICP-MS F-AAS, GF-AAS, ICP-OES, ICP-MS F-AAS, GF-AAS, ICP-OES, ICP-MS UV– VIS, IC

Ag, Al, As, Ba, Be, Ca, Cd, Co, Cr, Cu, Fe, K, Na, Pb, Mg, Mn, Mo, Ni, Sb, Se, Tl, V, Zn Al, As, Ba, Be, Ca, Cd, Co, Cr, Cu, Fe, K, Na, Pb, Mg, Mn, Mo, Ni, Se, Tl, V, Zn Ag, Al, As, Ba, Be, Ca, Cd, Co, Cr, Cu, Fe, K, Na, Pb, Mg, Mn, Mo, Ni, Sb, Se, Tl, V, Zn Be, Cd, Co, Cr, Mo, Pb, Tl, V Ag, As, Ba, Be, Cd, Co, Cr, Cu, Mo, Ni, Pb, Sb, Se, Tl, V, Zn Al, As, Ba, Be, Ca, Cd, Co, Cr, Cu, Fe, K, Na, Pb, Mg, Mn, Mo, Ni, Se, Tl, V, Zn Ag, Al, As, B, Ba, Be, Ca, Cd, Co, Cr, Cu, Fe, Hg, K, Na, Pb, Mg, Mn, Mo, Ni, Sb, Se, Sr, Tl, V, Zn Ag, Al, As, Ba, Be, Ca, Cd, Co, Cr, Cu, Fe, K, Na, Pb, Mg, Mn, Mo, Ni, Se, Tl, V, Zn Cr(IV)

F-AAS, ﬂame atomic absorption spectroscopy; GF-AAS, graphite furnace atomic absorption spectroscopy; ICP-OES, inductively coupled plasma-atomic emission spectrometry; ICP-MS, inductively coupled plasma-mass spectrometry; UV– VIS, ultraviolet– visible spectrometry; IC, ion chromatography.

P. Drouin and R. Clement

Method

Sample preparation for industrial waste analysis

these methods is small. By describing the methods currently used by the United States Environmental Protection Agency (US EPA), the general approaches to industrial waste sample preparation can be understood. Four methods have been developed by the US EPA for the digestion of aqueous sample types [10–13]. The applicability of these methods depends on the type of sample being analyzed, digestion equipment available (hot plate/block or microwave), and analytical instrumentation being employed. Methods 3005A and 3010A are hot plate digestions, and method 3015A is a microwave digestion method employing nitric and HCl [10–12]. Method 3020 is also a hot plate digestion procedure but is designed for samples to be analyzed by GF-AAS and employs nitric acid only [13]. Method 3005A is designed for surface and ground water analysis. As deﬁned by the method, analytical results from samples that have ﬁrst been passed through a 0.45 mm ﬁlter are designated as “dissolved” metals, and results from unﬁltered samples are designated as “total recoverable” metals. In this procedure, nitric acid (2 ml of HNO3) and HCl (5 ml of HCl) are added to a 100 ml sample, and the sample is heated (90 –958C) until its volume has been reduced to about 15–20 ml. It is important that the samples not be boiled because this may result in the loss of antimony. After adjusting the volume to 100 ml with reagent, and mixing, samples are ready for analysis by F-AAS, GF-AAS, or ICP-OES [10]. Method 3010A is similar to 3005A but is designed for samples that contain suspended solids (i.e. TCLP extracts and waste water). In this method, 100 ml samples are acidiﬁed with nitric acid (3 ml of HNO3) and the samples heated (90–958C) on a hot plate until the volume is reduced to about 5 ml. During heating, additional nitric acid is added until the digestion is complete. This is indicated by a clear digestate and the absence of brown fumes when additional acid is added. After reducing the sample volume to about 5 ml, and adding 10 ml of HCl (1:1), samples are reﬂuxed for 15 min. The sample is brought to volume (100 ml) and analyzed by F-AAS or ICP-OES [11]. Method 3020 is similar to 3010A except no HCl is employed, and 3020 was developed for GF-AAS [13]. The Ontario Ministry of the Environment (MOE) method for TCLP extracts is similar to US EPA 3010A [24]. An aliquot of sample (25 ml) is transferred to a digestion tube (Folin-Wu tube). Samples are placed in an oven and heated until the volume is reduced (, 5 ml). Nitric acid (1 ml of HNO3) and 3 ml of HCl are added, and the digestion tubes placed in a hot block. Samples are digested at 508C for 1 h and then at 958C for an additional hour. Boiling chips are added to ensure that the samples do not “bump” during digestion. Sample volumes are adjusted (25 ml) with reagent water and then thoroughly mixed. The resulting solutions are analyzed by ICP-OES [24]. For the determination of mercury in waters and wastewaters, samples are digested with a mixture of oxidizing agents to ensure that any organic mercurials are converted to inorganic mercury that can be quantitated by CV-AAS. The MOE method involves the addition of 1.2 ml sulfuric acid, 0.5 ml nitric acid, potassium persulfate (0.3 ml of 5% w/v K2S2O8), and potassium

941

P. Drouin and R. Clement

dichromate (0.3 ml of 5% w/v K2Cr2O7) to a 25 ml sample, which is then heated by water bath (87 ^ 38C) in a borosilicate culture tube (with caps) for 2 h [25]. Samples are allowed to cool to room temperature and are then analyzed by using a continuous ﬂow CV-AAS system. This is similar to the US EPA method 7470A, except potassium dichromate is used as an oxidizing agent instead of potassium permanganate [26]. Method 3015A is the microwave digestion method for water samples (drinking, surface, ground and waste) and extracts (TCLP). The original 3015 method was designed as a nitric acid-only procedure but the revised method allows for the addition of HCl [1,12]. This modiﬁcation allowed for improved recoveries for certain elements (Al, Sb, Ba, Be, Cr, Fe, Mg, Ag, and V). Samples (45 ml) are placed in a PFA, TFM, or quartz microwave vessel. Either nitric acid (5 ml) or a nitric acid (4 ml)/HCl (1 ml) combination is added to samples. A programmed microwave oven is used to rapidly increase sample temperature to 170 ^ 58C over 10 min, which is held for an additional 10 min. Samples are allowed to cool, and then are analyzed by an appropriate analytical method (F-AAS, GF-AAS, ICP-OES, ICP-MS, CV-AAS). Most modern microwaves feature preprogrammed US EPA methods. Samples that contain signiﬁcant amounts of organic material should be diluted with water because the evolution of gaseous digestion products may cause the pressure release valve to rupture, resulting in the potential loss of sample or volatile elements. Even with diluted samples, the same ﬁnal sample size (45 ml) is recommended. Link reported on the development of the 3015A method [1]. Studies involved the use of simulated waste water samples by mixing solid certiﬁed reference materials (CRMs) [e.g. ,400 mg of SRM 2704 (Buffalo River sediment), SRM 4355 (Peruvian soil), SRM 1084A (Wear-metals in lubricating oil), SRM 1634C (Trace metals in fuel oil)], with high purity water to produce solutions with about 0.9% solids. Samples were digested in a microwave oven with either nitric acid or a nitric acid/HCl mixture, heated to 170 ^ 58C in 10 min, which was held for 10 min. Results from the simulated waste water samples indicated that the method compared well to the 3051A and 3050B leach results. Addition of HCl to the digestion procedure increased recoveries for silver, antimony, and vanadium. This work also allowed for the validation of results for boron, mercury, and strontium, which have been incorporated into the method. 28.4.2 Solid sample types – US EPA methods For solid samples (sediments, sludges, and soils), the US EPA has three different digestion methods [15–17]. Method 3050B is based on hot plate digestion, and method 3051A employs microwave digestion. Method 3052 also uses microwave digestion, but includes HF and is designed to provide a “total” digestion. For method 3050B, samples (1 ^ 0.01 g dry weight) are weighed into digestion vessels. After addition of 10 ml HNO3 (1:1), samples are heated

942

Sample preparation for industrial waste analysis

(95 ^ 58C) and allowed to reﬂux for 10–15 min. Samples are cooled, and 5 ml concentrated HNO3 is added. Samples are reheated and reﬂuxed for 30 min. If brown fumes emanate from samples when the nitric acid is added, the reaction is not complete and additional aliquots of nitric are to be added until brown fumes are no longer produced. Samples are then heated (95 ^ 58C) for 2 h, or until the volume has been reduced to , 5 ml. After cooling, 2 ml water and 4 ml hydrogen peroxide (30% H2O2) are added to samples, which are then carefully heated until effervescence subsides, and are once more allowed to cool. The process of adding H2O2 is repeated until effervescence is minimal. Sample volume is reduced to less than about 5 ml by heating (95 ^ 58C). If analysis by GF-AAS or ICP-MS is chosen, samples are diluted to 100 ml, and are ﬁltered or centrifuged prior to analysis. For samples to be analyzed by F-AAS or ICP-OES, 10 ml HCl is added to samples, which are then reﬂuxed (95 ^ 58C) for 15 min. Digestates are then ﬁltered (Whatman No. 41 or equivalent) and taken to volume (100 ml) with water. For samples that are found to have poor recoveries for antimony, barium, lead, and silver, the following modiﬁcation to the method may be followed. Samples (1 ^ 0.01 g dry weight) are digested with nitric acid (2.5 ml of HNO3) and HCl (10 ml of HCl) at 95 ^ 58C for 15 min. Samples are ﬁltered (Whatman No. 41 or equivalent) and ﬁlter papers washed with 5 ml of hot (95 ^ 58C) HCl followed with 20 ml hot (95 ^ 58C) water. Filter papers and corresponding samples are then placed back into their digestion vessels and heated (95 ^ 58C) in 5 ml HCl until the ﬁlter paper is dissolved. After samples are ﬁltered, corresponding ﬁltrates are combined and allowed to cool. If a precipitate occurs upon cooling, 10 ml HCl may be added to help redissolve the precipitate. Samples are then diluted to volume (100 ml) with nitric acid. The MOE also uses aqua regia to digest industrial solid wastes [27]. The MOE procedure involves placing samples (0.5 g) in digestion tubes and adding 1.5 ml nitric acid and 4.5 ml HCl. After allowing samples to digest cold for 15 min, they are placed in a digestion hot block. Samples are gently heated (508C) for an hour and then at 958C for an additional 3 h. 28.4.3 Reported studies—hot plate digestion A number of authors have reported on variations of method 3050B for a variety of sample types. Open vessel aqua regia digestions have been used for the determination of phosphorous and rare earth elements (REE) in manure and fertilizer [28]. Freeze-dried manure samples (2 –3 g) were combined with 5 ml water and 10 ml aqua regia. Solutions were allowed to cold digest overnight, after which they were reﬂuxed for 1.5 h. The resulting solutions were diluted (100 ml) and analyzed by ICP-MS. REE concentrations in manure samples were all , 2.5 ppm, which was lower than concentrations in the fertilizer samples. In another study, aqua regia (4:1 HCl:HNO3) was used to analyze soil samples for phosphorous [29], where phosphorous results were compared to

943

P. Drouin and R. Clement

results obtained by a sodium carbonate fusion. Both dissolution techniques produced acceptable results for the CRMs analyzed (SO-1 and SO-4). Estonian soil samples were digested using an open vessel hot plate by employing a nitric acid/hydrogen peroxide mixture, and compared to results obtained by aqua regia digestion [30]. In the nitric acid/hydrogen peroxide procedure, 0.5 g soil samples were added to a conical ﬂask with 10 ml nitric acid. After a night of cold digestion, samples were heated (708C) for 2–5 days. Each morning 2 ml hydrogen peroxide was added. For the aqua regia digestion, 0.5 g samples were added to a conical ﬂask with 2.5 ml nitric acid and 7.5 ml HCl. The aqua regia mixture was allowed to cold digest overnight. Samples were then heated (708C) on a hot plate for 1–5 days. The resulting solutions were analyzed for copper, calcium, iron, magnesium, manganese, lead, and zinc by F-AAS. The nitric acid/hydrogen peroxide mixture showed no signiﬁcant difference in the extraction of copper, manganese, or zinc compared to aqua regia digestions, but the aqua regia digestion extracted more calcium, magnesium, and iron. After only one day, about 80 –90% of the total recoverable metals were extracted. In another study, a hot block aqua regia digestion procedure was used to analyze soil samples for thallium [31]. Samples (5.0 g) were weighed into a digestion tube and mixed with 2 ml water and 7 ml nitric acid, and then allowed to sit overnight to cold digest. Twenty milliliters of HCl were added and samples were then digested at 1508C for 2 h. After ﬁltering and dilution to the ﬁnal volume (100 ml) with water, samples were analyzed by ICPMS. In the analysis of several samples from the Wageningen Evaluating Programmes for Analytical Laboratories (WEPAL) International Soil Exchange Program, overall results were in good agreement with the reported study results. Myo¨ha¨nen et al. determined arsenic, cadmium, chromium, and lead using a simultaneous GF-AAS (Perkin Elmer SIMAAS 6000) for soil sample digested in an open vessel hot block with aqua regia [32]. They added 7.5 ml HCl and 2.5 ml nitric acid to 1.0 g samples after wetting samples with 1–2 ml water. Samples were allowed to stand overnight to cold digest and then heated (1408C) for 2 h. The resulting digestates were ﬁltered and the ﬁlters washed with warm nitric acid (5 £ 10 ml of 14% HNO3). They also reported on a sample digestion procedure using larger sample volumes (3.0 g) and increased acid volumes (22.5 ml of HCl and 7.5 ml of HNO3). No signiﬁcant differences in the results were observed between the two sample sizes examined. Masson examined the results from a number of different digestions to analyze thallium in soil and vegetation samples (maize, cabbage, and rye) [33]. Aqua regia, nitric acid/hydrogen peroxide, and a nitric/sulfuric acid mixture were used to digest soil samples, whereas vegetation samples were only digested using aqua regia and nitric acid/hydrogen peroxide. For soil samples, the nitric/sulfuric acid digestion produced low results for thallium. The aqua regia digestion resulted in results that were biased low, which may be due to a loss of volatile thallium complexes (TlCl3·4H2O). Nitric acid/hydrogen peroxide digestions produced the best results for both soil and vegetation samples.

944

Sample preparation for industrial waste analysis

28.4.4 Reported studies—microwave digestion methods US EPA method 3051 is a closed vessel microwave digestion method. Samples (0.5 g) are digested in a PFA, TFM, or quartz vessels with 10 ml nitric acid. Smaller sample sizes (0.25 g) can be used if the sample is contaminated with oil. This microwave-assisted 10-min digestion includes a 5.5 min interval to raise the temperature to 175 ^ 58C, followed by a 4.5 min hold time. Because digestions using only nitric acid may result in poor recoveries for certain elements (Ag, Sb, and Sn), method 3051A was developed. The overall procedure uses the same nitric acid (10 ml of HNO3) or mixture of nitric acid (9 ml of HNO3) and HCl (3 ml of HCl). Link et al. reported on the development and validation of the US EPA method 3051A [2] comparing the draft method to the hot plate digestion method 3050B and the microwave digestion method 3051 [2]. The addition of HCl, along with nitric acid, required digestion vessels capable of withstanding 30 atm (435 psi), and resulted in better recoveries of antimony, iron, and silver. Other authors have reported similar results [3,4]. It was also noted that more accurate temperature control resulted in more precise determinations. As part of their validation of the method, they created a simulated sludge by mixing SRM 2704 (0.25 g of Buffalo River sediment) and SRM 1634c (0.25 g of trace metals in fuel oil). They found that the method provided adequate recoveries and the results compared well to those arising from method 3050B [2,16]. Other authors have examined problems with the analysis of soils contaminated with oil and grease, comparing a nitric acid/hydrogen peroxide digestion to a nitric acid/perchloric acid digestion [34]. The nitric acid/hydrogen peroxide digestion involved adding 15 ml nitric acid to 0.2 g samples and allowing them to cold digest overnight. Samples were then heated (1508C) in a block digester until evolution of brown fumes stopped (3 –8 h). After solutions were cooled and 5 ml hydrogen peroxide added, samples were heated (1208C) and 1 ml aliquots of hydrogen peroxide added until effervescence stopped. In the second procedure, 1 g samples were cold digested overnight in 15 ml nitric acid by heating 1208C by using a block digester. The temperature was increased to 1508C, and held until the evolution of brown fumes ceased. During this step, nitric acid was added to maintain a constant volume of about 10– 15 ml. After solutions were allowed to cool, 5 ml perchloric acid were added. Samples were then heated at 1608C for 2 h in a block digester. The digestion was considered complete when the perchloric acid began to fume (white fumes). Solutions were brought to volume (100 ml) and analyzed by ICP-OES and F-AAS. The nitric/perchloric acid digestion provided almost complete digestion of CRM NIST 2710 for cadmium, copper, nickel, lead, and zinc. There was no statistical difference between results obtained from the two digestion procedures. There was an exception for cadmium, but the nitric/perchloric acid digestion did not extract a signiﬁcant level of cadmium (i.e. below instrumental detection limits) [34].

945

P. Drouin and R. Clement

Zhang and Hattey [35] compared an open vessel microwave digestion to a hot block for the digestion of cattle and chicken manure. For the hot block digestion, 0.5 g samples were placed in a Pyrex digestion tube. A nitric/ perchloric acid mixture was added and samples were taken through a stepwise digestion procedure (45 min at 1308C; 15 min at 1708C; 90 min at 2158C). Larger sample sizes (5 g) were used with the open vessel microwave system. The microwave system employed automatically added acids to samples. Results for the two digestion approaches were comparable but the microwave technique offered a signiﬁcant increase in analytical speed. The larger sample sizes that the open microwave digestion system can accommodate are beneﬁcial for the analysis of heterogeneous industrial waste samples. Maaskant described the development of a certiﬁed sewage sludge reference material [36], in which an aqua regia digestion procedure was compared to the US EPA method 3050A. The aqua regia digestion resulted in greater extraction of metals. De Abreu analyzed anaerobically digested sewage sludge, swine manure, dairy manure, pond sediment no. 2 (NIES, National Institute for Environmental Studies), and municipal compost following a number of different digestion procedures [37]. An open vessel digestion procedure was employed in which 0.5 g samples were added to a Pyrex tube with 5 ml nitric acid. After allowing the sample/acid mixture to cold digest for 30 min at room temperature, 1 ml perchloric acid was added and the temperature increased to 2108C and held until the digestion was complete. The resulting solution was diluted to 50 ml with water for analysis. Two digestion procedures were compared: one procedure was similar to US EPA method 3051, wherein 0.5 g samples are digested with nitric acid, and a second procedure was a microwave digestion that employs aqua regia. For analysis of the pond sediment no. 2 CRM, both microwave digestion procedures examined extracted higher concentrations of the metals of interest (Cd, Cr, Cu, Fe, Mn, Ni, Pb, and Zn) than the nitric/perchloric acid digestion. Both microwave digestion procedures produced comparable results except for manganese and zinc, for which the aqua regia results were greater. Microwave digestion with aqua regia resulted in the highest metal concentrations for the various sample types analyzed. Depending on the sample type, those digested by microwave employing aqua regia had chromium concentrations two- to fourfold higher than samples digested in the hot block using nitric and perchloric acid. Only aqua regia was an effective acid mixture for digesting organic wastes (i.e. manure) in these studies [37]. Xing and Veneman compared the standard US EPA method 3051 to a modiﬁed version in which hydrogen peroxide was added in addition to nitric acid [38]. Samples were initially digested according to method 3051with 2 ml hydrogen peroxide being added after cooling, after which microwave digestion was repeated. Except for manganese in a couple of samples, there was no statistical difference in results obtained for the two digestion procedures. In another study, aqua regia microwave digestions were used for the analysis of silver, lead, and tin in sediments by GF-AAS [5]. A pyrolytic

946

Sample preparation for industrial waste analysis

graphite platform treated with ruthenium (Ru) was used a permanent modiﬁer. Excellent agreement with certiﬁed total values for three CRMs (LKSD-3, MESS-2, STSD-2) was obtained. In method 3052, solid samples are weighed (0.5 ^ 0.001 g) into a Teﬂon PFA vessel, and nitric (9 ml) and hydroﬂuoric (3 ml) acids are added [17]. The amount of hydroﬂuoric acid added may vary (0.5 –5 ml of HF), depending on the amount of silicon dioxide present in the samples. The method also allows for the addition of HCl to help stabilize silver, barium, and antimony. With the microwave temperature program used, the temperature is increased to 180 ^ 58C over 5.5 min, and then maintained for an additional 9.5 min, after which samples are allowed to cool to room temperature. For ICP-OES analysis, boric acid may be added to samples to complex ﬂuoride and protect the quartz torch. Alternatively, HCl and hydroﬂuoric acid volumes may be reduced by evaporating samples to near dryness. This may result in the loss of some volatile elements (i.e. arsenic, antimony, and zinc). Chen and Ma used three standard reference materials (SRMs) (NIST 2704, 2709, 2711) and 20 Florida soil samples to evaluate hot plate and microwave digestions using aqua regia in addition to an aqua regia/hydroﬂuoric acid mixture [7,39]. Subsamples of 0.5 g were weighed into a Teﬂon PFA vessel to which 12 ml freshly prepared aqua regia were added. Samples were then digested at a pressure of 0.69 £ 106 Pa for 5.5 min. The aqua regia/hydroﬂuoric acid mixture was used by ﬁrst adding 4 ml hydroﬂuoric acid to samples and letting them digest overnight in the PFA vessel. Twelve milliliters of aqua regia were then added, and samples were further digested in a microwave oven at a pressure of 0.83 £ 106 Pa for 20 min. Boric acid (2 g) was added to each sample to complex the free HF. For both digestion procedures, samples were ﬁltered and brought to volume (100 ml) with double distilled water. The open vessel digestion involved adding 12 ml freshly prepared aqua regia to 0.5 g samples in a beaker and heating at 1108C for 3 h. Samples were then heated to near dryness and diluted with 2% nitric acid. Samples were ﬁltered and diluted to volume (100 ml) with water. A number of elements were determined by ICPOES (Al, Ba, Ca, Fe, K, Mg, Mn, P, and Zn) and GF-AAS (As, Cd, Cr, Cu, Ni, Pb, and Se). The average precision (% RSD) for all elements examined was comparable for data generated employing aqua regia hot plate (4.6%) and aqua regia microwave (5.2%) digestions, which were slightly higher than the data precision observed for the aqua regia/HF digestion (3.7%). Poor precision for selenium, arsenic, and cadmium was attributed to the low concentrations of these elements, and to chemical interferences observed in the GF-AAS analyses. It was observed that the hot plate digestions extracted approximately 70% of the total metals available, whereas the microwave/aqua regia digestions extracted 80% of the total metals available. The aqua regia/hydroﬂuoric acid mixture microwave digestion procedure recovered 94% of the total metals available. Low chromium recovery for all digestion techniques is attributed to the presence of insoluble chromium minerals (i.e. chromite, FeCr2O4).

947

P. Drouin and R. Clement

The microwave digestion procedure extracted higher concentrations than the hot plate digestion, but the difference was only signiﬁcant for aluminum, calcium, iron, magnesium, barium, lead, and zinc. Mester et al. compared results for six different acid mixtures in two microwave digestion systems to a hot plate digestion procedure for the analysis of ﬂy ash [40]. The hot plate digestion procedure involved adding the 36 g samples to a 2 l beaker and adding 400 ml aqua regia. Solutions were heated at 708C until a dense sludge were formed. Then 20 ml HCl was added and samples were boiled for 2 h, after which 500 ml water was added. The resulting solutions were ﬁltered and the volumes adjusted to 2 l with water. The microwave digestion procedures involved digesting the sample (0.2 g) in nitric acid (4 ml), nitric acid (3 ml)/hydroﬂuoric acid (1 ml), nitric acid (1.5 ml)/ hydroﬂuoric acid (1 ml)/hydrogen peroxide (1.5 ml), nitric acid (1.5 ml)/ hydroﬂuoric acid (1 ml)/HCl (1.5 ml), and nitric acid (1.5 ml)/hydroﬂuoric acid (1 ml)/perchloric acid (1.5 ml) mixture. For samples containing hydroﬂuoric acid, boric acid (1.5 ml of H3BO3) was added once the samples were cool. These were then reheated for an additional 7 min. The samples were analyzed by F-AAS for chromium, copper, nickel, lead, and zinc. Copper, lead, and zinc concentrations did not vary more than 10% between the six microwave digestions procedures tested. The hot plate digestion resulted in lower recoveries of chromium, copper, and nickel. Bettinelli et al. compared results obtained using an aqua regia and a nitric/hydrochloric/hydroﬂuoric acid microwave digestion of soil and sewage samples [6]. For the aqua regia digestion, 0.25 g samples were added to the PTFE digestion vessel and allowed to cold digest overnight in 8 ml aqua regia. Samples were then digested in a microwave oven for 14 min at three different power settings (250, 400, and 500 W). The digested samples were then ﬁltered and diluted to 50 ml with water. For the nitric/hydrochloric/hydroﬂuoric acid digestion procedure, 2 ml hydroﬂuoric acid were added to the samples in the digestion vessels. Nitric acid (2 ml) and 6 ml HCl were then added, and samples were cold digested overnight. During the digestion of samples by microwave, power was increased stepwise (250, 400, and 600 W) for 8, 4, and 6 min, respectively. Samples were allowed to cool and 2 ml boric acid was added to each sample, after which samples were reheated (300 W) for three additional minutes. The aqua regia results were compared to the certiﬁed aqua regia results generated using an open vessel digestion procedure (DIN 38414-S7). Percent recoveries ranged from 89 to 110%. Statistical analysis of the data indicated that the variance was related to instrumental measurements. For the nitric/hydrochloric/hydroﬂuoric acid digestion procedure, elemental recoveries were 83 –108%. The instrumental analysis was the greatest source of variance in the results. A number of authors have reported results for the analysis of soil samples using digestion procedures employing HF acid [41–43]. Liu et al. described a digestion designed to address a low bias in the total recovery of chromium in

948

Sample preparation for industrial waste analysis

marine sediment (BCSS-1) [8]. Results from open vessel, hot plate digestion, and closed vessel microwave digestions were compared. The open vessel digestion involved adding 3 ml nitric acid, 3 ml HF, 1 ml perchloric acid to the 0.5 g samples. Samples were placed on a hot plate and heated at 2208C for 4 days. They were redissolved by adding nitric acid (20 ml of 10% HNO3) and heating for 1 h. For the microwave digestion procedure, the 0.25 g samples were digested 20 min after addition of 3 ml nitric acid, 3 ml HF, and 1 ml perchloric acid, at an elevated pressure (140 psi). Once samples were cooled, vessels were uncapped and heated to near dryness to remove silica and HF. The samples were reconstituted with 1 ml nitric acid and 5 ml water. The same digestion procedure was performed using a high-pressure digestion system, wherein samples were digested for 2 min at 250 psi, followed by 1.5 min at 1000 psi, 3 min at 600 psi, and 20 min at 250 psi. With these multi-acid digestion procedures only 112 mg/g of chromium was recovered, compared to the certiﬁed value of 125 mg/g. By using the same procedures, spike addition recoveries averaged 98%, and good agreement with certiﬁed values was obtained for a second reference material (132 mg/g versus 135 mg/g for NIST 2704, Buffalo River sediment). These results illustrate that refractory elements may be difﬁcult to digest, and metal recoveries are dependent on the speciﬁc sample matrix. 28.4.5 Ultrasound-assisted extractions Ultrasonic extraction can be an effective means of extracting metals from various sample types, and has been used to extract metals from soils, dust, paint, and air ﬁlters [44]. The concepts have been treated in detail by Ashley in Chapter 12. Arambarri et al. used an aqua regia/hydroﬂuoric acid mixture to determine tin in sediments [45]. They employed an ultrasonic bath to help in the digestion process. Aqua regia (5 ml) was added to 0.35 g samples in a Teﬂon tube, which was placed in an ultrasonic bath for 15 min at 208C. Samples were then heated to 558C for 20 min, after which HF was added. Samples were then heated to 688C for an additional 68 min. Samples were diluted with 2 ml nitric acid to the ﬁnal volume of 25 ml, for analysis by GF-AAS. Excellent agreement with the PACS-2 CRM was achieved. 28.4.6 Alkaline digestions The US EPA method 3060A is an alkaline digestion procedure designed to extract soluble and insoluble hexavalent chromium (Cr6þ) compounds from soils, sludges, and sediment [18]. The method was designed to extract all soluble, adsorbed, and precipitate forms of hexavalent chromium while ensuring that no trivalent chromium is oxidized, and that no hexavalent chromium is reduced. The resulting solutions may be analyzed by colorimetry

949

P. Drouin and R. Clement

(method 7196A) or ion chromatography (method 7199) [46,47]. The method involves digesting 2.5 g samples with sodium carbonate/sodium hydroxide (0.28 M Na2CO3 and 0.5 M NaOH) at 90 –958C for 1 h. For samples containing high concentrations of trivalent chromium, the addition of magnesium in a phosphate buffer can be used to minimize method-induced oxidation. Prompt analysis of the neutralized, extracted sample is recommended to ensure that hexavalent chromium is not reduced, which would result in concentrations that were biased low. Species speciﬁc isotope dilution mass spectrometry (SS-IDMS) is an alternative detection method that uses isotopic enriched spikes to help correct for any oxidation or reduction of chromium compounds, which may be effective in reducing analytical bias [48]. 28.4.7 Laboratory safety Digestions involve the use of concentrated acids, high temperatures, and—in the case of microwave digestions—high pressures that could potentially result in personal injury. Proper safety precautions, including a thorough understanding of the reagents (i.e. material safety data sheets), the safe operation of equipment used, and the use of personal protective equipment (e.g. gloves, lab coat, and safety glasses), must be practiced at all times. All open vessel digestions must be performed in a properly functioning fume hood to ensure that corrosive and toxic fumes are removed. Microwave vessels should be vented in a fume hood after vessels have cooled (see manufacturer’s instructions). When perchloric acid is being used, a properly designed perchloric acid fume hood must be used. Special caution must be followed when using perchloric acid because perchloric acid may react violently with samples containing high organic components, which frequently occur in industrial waste applications. HF causes severe tissue damage and bone destruction.

28.5

LEACH PROCEDURES

A number of laboratory scale batch leach tests have been developed to determine whether industrial wastes pose an environmental hazard because of the potential leaching of metals and other toxic contaminants to the environment, especially to ground water [49–54]. These leach tests are used to determine whether a sample is a hazardous waste (as per deﬁned regulations). Leach tests are different from acid digestions in a number of ways. First, leach procedures are generally designed to mimic the environment to which the waste will be exposed and therefore are less aggressive than acid digestions. Leach solutions are typically deionized water or dilute acid (pH .2) solution. Second, because of the heterogeneous nature of many industrial wastes, large sample sizes (70– 100 g) are employed. Third, a large liquid to

950

Sample preparation for industrial waste analysis

solid ratio is used (20:1), to ensure that the extraction is not limited by solubility. Finally, leach tests are performed at room temperature and are characterized by long contact times (e.g. 18 h). One of the most widely used leach procedures, the US EPA’s TCLP, is detailed below [49]. 28.5.1 Toxicity characteristic leaching procedure In 1984, the US Congress passed the Hazardous and Solid Waste Amendment (HSWA) that directed the US EPA to address some of the limitations of its previous leach test, the extraction procedure (EP) [55]. The new leaching procedure was required to be applicable to all waste types, allow for the analysis of volatile organic compounds (VOCs), minimize operational difﬁculties, and improve the precision of the data, and address practical considerations (time, effort, and cost). In the development of the TCLP test, municipal landﬁll leachates were collected using large-scale lysimeters, and used to leach a variety of industrial wastes. A number of laboratory leach tests, both batch and column leach tests, were performed and compared to the results of the municipal landﬁll leachate. A statistical comparison of results showed that a buffered acetate solution mimicked the results of the column leaching of the municipal landﬁll leachate and was chosen as the extraction solution for the TCLP test. While the TCLP method is operationally similar to the EP, the TCLP method was found to be more rugged, more precise, and allowed for the determination of VOCs [56]. 28.5.2 TCLP regulatory limits The TCLP method regulatory limits were determined by using a backcalculation from an acceptable chronic exposure level at a well that has been contaminated by leachate from a municipal landﬁll based upon the US EPA Composite Model for Landﬁlls (EPACML) [56]. The TCLP regulatory limits included eight inorganic elements and 20 organic compounds [57]. The inorganic limits are listed in Table 28.2. A number of other jurisdictions have adopted the TCLP method, including the Ontario MOE in 2000 [58]. The MOE added a number of inorganic and organic parameters to the original list of analytes, and their regulatory limits are listed in Table 28.2. Spanish regulators employ the TCLP method to determine the ecotoxicity parameter (EC50) [54,59]. Samples are processed using the TCLP method and the leachate is ﬁltered. The EC50 parameter is evaluated using a luminescence bioassay with the marine bacterium Photobacterium phosforeum in a Microtox analyzer. When the luminescent bacteria are exposed to toxic materials that interfere with the metabolism of the growing organism, the light emission output decreases in proportion to the toxicity of the sample. The Spanish regulations deﬁne a hazardous waste as a sample with EC50 lower than 3000 mg/l;

951

P. Drouin and R. Clement TABLE 28.2 The TCLP regulatory limits for the US EPA and the Ontario MOE (Environmental Protection Act, Regulation 558 (amendment to regulation 347) Schedule 4) [57,58] Element

US EPA regulatory limit (mg/l)

Ontario MOE regulatory limit (mg/l)

Arsenic (As) Barium (Ba) Boron (B) Cadmium (Cd) Chromium (Cr) Lead (Pb) Mercury (Hg) Selenium (Se) Silver (Ag) Uranium (U)

5.0 100

2.5 100 500 0.5 5.0 5.0 0.1 1.0 5.0 10

1.0 5.0 5.0 0.2 1.0 5.0

moderately hazardous wastes as samples with EC50 between 3000 and 30,000 mg/l; and samples with EC50 greater than 30,000 mg/L, are not considered hazardous. 28.5.3 TCLP method summary The TCLP procedure was designed for both inorganic and organic compounds including VOCs. Specially designed zero headspace extraction (ZHE) vessels are used for the determination of VOCs. For the purposes of this chapter, the use of TCLP for metals is emphasized. Readers are directed to the method for speciﬁc requirements for the determination of organics and VOCs [49]. Collected samples for TCLP evaluation are not preserved, but are stored at 48C until analysis. If a sample contains a liquid component, the sample is ﬁltered through a glass ﬁber ﬁlter (0.6 –0.8 mm). The ﬁltrate is retained and is combined later with the TCLP leachate. A common waste management treatment process involves the solidiﬁcation and stabilization of waste by mixing the waste with cement to form monoliths. To allow for the analysis of wastes that have been processed in this manner, the TCLP procedure allows for particle size reduction if necessary (all particles should be able to pass through a 9.5 mm standard sieve). Two possible extraction ﬂuids can be used: a buffered acetic acid solution with pH 4.93 or an acetic acid solution with pH 2.88. To determine which extraction ﬂuid is to be employed, a 5 g subsample of the solid is mixed with water (96.5 ml). If the resulting water has a pH , 5.0, then the pH 4.93 extraction ﬂuid is used; if the pH . 5.0, then the pH 2.88 extraction

952

Sample preparation for industrial waste analysis

ﬂuid is used. The solid component of the sample is added to extraction vessel and the appropriate extraction ﬂuid is added (at a ﬂuid:sample ratio of 20:1). Samples are mixed by rotation in a mechanical mixer at 30 ^ 2 rpm for 18 ^ 2 h. Extractions are performed at room temperature (22 ^ 28C). Leachates are passed through a glass ﬁber ﬁlter (0.6–0.8 mm) and combined with the initial liquid component (if they are compatible), the pH recorded, and samples are then acidiﬁed with nitric acid (pH , 2). Leachates are digested (method 3010A, 3015A or equivalent), and analyzed with appropriate instrumentation (e.g., F-AAS, ICP-OES, ICP-MS). If the initial liquid component and the leachate are not compatible, then each component is analyzed separately and the results are combined mathematically based on the volume-weighted average of the two components. With each analytical batch, at least one blank must be analyzed and one matrix spike must be analyzed for each waste type being analyzed [49].

28.5.4 TCLP applications A wide variety of industrial wastes ranging from wastewater treatment sludge [60], tannery wastes [59,61,62], contaminated soils [63–68], plating wastes [54, 69], foundry wastes [70,71], ﬂy ash [53,72– 76], and sewage sludge [77,78] have been analyzed using the TCLP method. The TCLP procedure is a useful tool in evaluating the effectiveness of hazardous waste treatment programs. Ramesh and Kozin´ski showed that by increasing the ashing temperature from 1000 to 14008C, they were able to reduce the amount of chromium, lead, and cadmium leached from synthetic paper mill sludge [79]. Chen et al. used mineral apatite (Ca10(PO4)6(OH,F,Cl)2) to absorb lead, cadmium, and zinc from synthetic waste water and contaminated soil samples [80]. They reported that the mineral apatite was able to adsorb 151 mg of lead per gram of apatite. TCLP leachate of apatite showed that lead levels would be below the regulatory limit. Some success was observed for cadmium and zinc but the results were not as promising for all metals. Mixing apatite with lead-contaminated soil reduced the amount in the TCLP leachate so that the soil would not be classiﬁed as contaminated. Sletten et al. reported on the development of a treatment process for metal-contaminated leachate from a municipal solid waste dump [60]. Metals in leachate were precipitated by increasing the sample pH. The resulting sludge was tested with the TLCP method and did not fail TCLP guidelines. Ekhter et al. examined the effect of long cure times on solidiﬁcation/ stabilization of arsenic salts [74]. They found that Portland cement, alone, was effective at reducing the arsenic concentration (As3þ and As5þ) in the TCLP leachate to less than 5 mg/l. The addition of ﬂy ash to the Portland cement resulted in an increase in the amount of arsenic leached. A slight decrease with increasing time of the amount of arsenic extracted from Portland cement by the TLCP procedure was observed for the three cure times reported (28 day, 1 year,

953

P. Drouin and R. Clement

3 years). Peters reviewed the current methods for the remediation of contaminated soils with chelating agents [63]. After successful remediation, the decontaminated soils passed both the TCLP and US EPA Total Extractable Metal limits. Rinehart et al. reported on how chromium’s oxidation state affected the amount of chromium in the TLCP extract [68]. A soil sample from one site had a “total” chromium of 478 mg/kg and the soil sample from second site had a “total” chromium concentration of 1272 mg/kg (nitric acid digestion). The use of Portland cement to stabilize the soil only resulted in a slight decrease in the amount of chromium extracted in the TCLP procedure (2.0 mg/l). These results show that solidiﬁcation/stabilization procedures alone are not always an effective waste treatment process. Janusa et al. examined the effect of particle size and contact time on the reliability of TCLP for solidiﬁed/ stabilized waste [81]. Although the TCLP method speciﬁes that samples must be rotated for 18 h, it is not speciﬁed how long the sample should be in contact with the leachate before the rotation is started or how long after the rotation is completed (before ﬁltering). Experiments indicated that excessive contact times (14–28 days) resulted in increased leaching of lead. Although the authors considered the contact times used in their study to be excessive, it is clear that ﬁltering leachates should be completed as soon as extractions are completed. Janusa et al. also showed that as the particle size is reduced, the percent of lead leached is increased. While the TCLP method requires that particle size of the sample be reduced to , 9.5 mm, this study showed that more aggressive particle size reduction would not be appropriate, especially in the case of solidiﬁed/stabilized wastes. Kendall provided a case study where metallic iron was employed to reduce the amount of lead extracted from sand molds used for brass casting [71]. The waste treatment procedure involved adding approximately 10% metallic iron (w/w) to the sand molds. During TCLP extraction, the metallic iron reduced the lead (Pb2þ to Pb0), which lowered lead concentrations below regulatory limits. In the case study, drill core samples were obtained from a waste site where the sand molds had been disposed. The TCLP procedure was performed on the drill core samples and relatively high concentrations of lead (66 ppm) were measured in the leachate from the deepest drill core sample (i.e. the samples that had been in the landﬁll the longest). In other words, the iron used to stabilize the lead waste was oxidized over a period of time, and therefore the lead became available for environmental leaching. Kendall noted that the use of metallic iron is not an effective waste treatment process because the waste is not permanently stabilized.

28.5.5 TCLP and sequential extractions In a number of studies, the TCLP method has been used to evaluate contaminated sediments. Hardaway et al. used the TCLP method to examine

954

Sample preparation for industrial waste analysis

contaminated sediment samples taken from the Bayou d’Inde in Louisiana [65]. Results were compared to a nitric acid/hydrogen peroxide microwave digestion of the sample. The results of the microwave digestion showed that the sediments collected were contaminated with chromium (12 –230 ppm), copper (10–660 ppm), mercury (0.03 –11 ppm), lead (11–640 ppm), and zinc (16–390 ppm). The TCLP method was only able to extract 7–40% of the corresponding metals concentration of the digested sample. Phillips and Chapple used the TCLP method to evaluate contaminated soil samples and compared the TCLP results to a four-step sequential extraction [66]. This extraction approach is detailed by Rauret in Chapter 39. The ﬁrst step was designed to determine metals adsorbed to exchangeable sites involving shaking of the soil sample (1 g) in a magnesium chloride solution (8 ml of 1 M MgCl2, pH 7) for 1 h at room temperature. Metals associated with the carbonate fraction of the soils were determined by using the residue from the ﬁrst step. It was then shaken for 6 h in a sodium acetate solution (8 ml of 1 M NaOAc, pH 5) at room temperature. The third step was designed to determine metals associated with the iron and manganese oxides. This involved taking the residue from the second step and extracting it with a hydroxylamine hydrochloride/acetic acid solution (20 ml of 0.04 M NH2 OH·HCl in 25% HOAc) for 6 h at 968C. To determine the metals associated with the organic fraction of the soil, the residue from the third step was digested with nitric acid/hydrogen peroxide (3 ml of 0.02 M HNO3 þ 5 ml 30% H2O2) for 2 h at 858C followed by the addition of more hydrogen peroxide (3 ml 30% H2O2), and an additional 3 h of heating (858C). An ammonium acetate/nitric acid (5 ml of 3.2 M NH4OAc in 20% HNO3) solution was added and samples were shaken for 30 min, then diluted with 20 ml deionized water. Extracts were analyzed for chromium, copper, iron, manganese, lead, and zinc using F-AAS. The concentrations of copper, chromium, iron, and lead extracted in the TCLP method compared well to the concentrations found in the exchangeable fraction (1st extraction step). The majority of copper was associated with the organic fraction. Chromium concentrations in the soil were low, but chromium was associated with the organic and oxide fractions. Lead was found to be associated with the iron and manganese oxides. Therefore, the TCLP method extracts metals associated with the exchangeable and carbonate fractions of the soil. Hale et al. provided a detailed description of the evaluation of a contaminated industrial site where soil had been contaminated with arsenic and chromium [64]. In addition to the analysis of the total arsenic and chromium concentrations in soil samples, the group evaluated the samples by performing the TCLP method and a sequential extraction procedure on the samples. The ﬁrst step of the sequential extraction procedure involved a water leach to extract arsenic and chromium associated with pore water and water-soluble salts. Samples were then leached with hydrogen peroxide to determine the arsenic and chromium concentrations associated with the

955

P. Drouin and R. Clement

soil’s organic matter. An ammonium oxalate/oxalic acid buffer (pH 3) was used to extract arsenic and chromium associated with amorphous iron hydroxides. The ﬁnal extraction involved hydrogen peroxide and HCl and was designed to remove the acid-soluble metal as well as metals adsorbed to the soil, without dissolving the mineral component of the soil. While the total arsenic and chromium concentrations were higher in clayey soil (at the top of the soil depth proﬁle), the TCLP method extracted more arsenic and chromium from the sandy soil (lower in the soil depth proﬁle). The selective extraction and grain-sized data indicated that arsenic was primarily associated with iron hydroxides, whereas the chromium was partitioned equally between the iron hydroxides and organic matter when the clay content was high (,28.1%). At the lower clay levels (. 28.1%), chromium was associated with pore water. Snyman compared the TCLP method to an ammonium ethylenediaminetetra-acetic acid (NH4EDTA) extraction method (used by the agricultural industry to determine the potentially available metal concentration for crop production), for the evaluation of sewage sludge [77]. Snyman reported that the TCLP method was only able to extract , 20% of metals (cadmium, copper, lead, and zinc), compared to aqua regia digests. In contrast, the NH4EDTA method extracted 36% of the copper and 70% of lead, compared to the aqua regia digests. Either NH4EDTA methods or aqua regia digestions should be used to classify and characterize sewage sludge before application to farmland because the use of the TCLP method underestimated metal concentrations in sewage sludge. The TCLP test should only be used for applications as deﬁned in the applicable legislation. 28.5.6 TCLP limitations The TCLP method was designed to simulate leaching of samples from a municipal solid waste dump. Industrial wastes are often disposed of in speciﬁcally designed monoﬁlls. In such cases, the TCLP method may not be the most appropriate test. Hassett suggested the composition of the leaching solution be site speciﬁc [53]. He described the synthetic groundwater leaching procedure (SGLP) that employs a buffered sodium sulfate/sodium bicarbonate solution, which has a similar composition to the groundwater found in central and western North Dakota, to evaluate the leaching of coal combustion by-products [53]. Egemen and Yurteri analyzed ﬂy ash and bottom ash from coal ﬁred power plants and compared the EP to the TCLP and ASTM D3987 methods [75]. They reported that the EP method extracted greater amounts of cadmium, copper, nickel, lead, and zinc than the TCLP and that the TCLP extracted more than the ASTM D3987 method [52]. However, the authors noted that results from the ASTM D3987 method were more consistent with the results from actual leachate collected from the ﬂy ash monoﬁll.

956

Sample preparation for industrial waste analysis

28.6

CERTIFIED REFERENCE MATERIALS

CRMs or SRMs are materials that are well characterized with certiﬁed chemical or physical properties. CRMs are often used during method development and validation to ensure the accuracy of the method and their development and application has been described in detail by Quevauviller in Chapter 4. CRMs can also be employed as part of a laboratory’s quality control and quality assurance programs by periodic analysis during routine operation. The CRM sample matrix chosen should match the sample matrix being analyzed as closely as possible (i.e., choose a sewage CRM for sewage analysis, if available). The CRM should contain as many of the target analytes as possible, at concentration ranges similar to expected concentrations of analytes in the samples being analyzed. The digestion procedure originally used to produce the certiﬁed value should be similar to the digestion procedure being used to analyze the samples. Some CRMs list total metal concentrations that may not be comparable to metal concentrations derived from methods that employ acid digestions. Fortunately, many CRMs are becoming available with certiﬁed or provisional values derived from a number of different digestion procedures [82]. Various CRM sample types are available from a number of international agencies, including water (surface water, ground water, sea water, rain water), landﬁll leachate, sediments (river, lake marine, estuarine), soils (contaminated and sewage sludge amended), ﬂy ash, sewage sludge, coal, plating sludges (copper and chrome plating sludges), furnace slag, and mine tailings (Table 28.3). Despite the number of CRMs now available, few industrial waste sample types are represented. When CRMs are not available, it may be possible to prepare synthetic samples by combining two separate CRMs, as was the approach employed to develop and validate the US EPA method 3015A and 3051A [1,2]. For sample types not represented by commercially available CRMs or SRMs, it is still possible to develop consensus value reference materials through round-robin studies or by statistical analysis of data from large, homogenized samples, which have been analyzed repeatedly over a signiﬁcant period of time.

28.7

SUMMARY AND FUTURE DEVELOPMENTS

Microwave digestions offer a number of advantages over conventional hot plate/block digestions. The high temperature/pressures that are produced inside the sealed digestion vessels result in faster digestion, less possibility of sample contamination, and retention of volatile elements (e.g. Hg). While the analytical results arising from a hot plate/block digestion and a microwave digestion system will be comparable for most elements and sample types, microwave digestion systems will generally result in a slightly more aggressive

957

958 TABLE 28.3 Examples of CRMs available and their producers Name (description)

Producer

LKSD-1,2,3,4 STSD-1,2,3,4 SO-2,3,4 TILL 1,2,3,4 RTS 1-4 UTS 1-4 SL1 SLEW 2

Lake sediment Stream sediment Soil Soil Sulﬁde tailings sample Uranium tailings samples Blaster-furnace slag Estuarine water

CANMET, Sales Manager, CCRMP, CANMET Mining and Mineral Sciences Laboratories, Minerals and Metals Sector, Natural Resources Canada, 555 Booth Street, Ottawa, Ontario, Canada, K1A 0G1. Tel.: (613) 995-4738, fax: (613) 943-0573, E-mail: [email protected]

SRM 1633B SRM 1632B SRM 2704 SRM 2710 SRM 2711

Coal ﬂy ash Trace elements in coal Buffalo River sediment Montana soil no. 1 Montana soil no. 2

NIST, Standard Reference Materials Group, National Institute of Standards and Technology, 100 Bureau Drive, Stop 2322, Gaithersburg, MD 20899-2322, USA. Tel.: (301) 975-6776, fax: (301) 948-3730, E-mail: [email protected]

LGC 6138

Polluted soil coal-carbonisation site soil Landﬁll leachate

LGC Promochem, Queens Rd, TEDDINGTON, Middlesex, TW11 0LY, United Kingdom. Tel.: þ44 (0)20-8943-7000, fax: þ44 (0)20-8943-2767, E-mail: [email protected]

LGC 6175

continued

P. Drouin and R. Clement

Code

TABLE 28.3 (continuation) Name (description)

Producer

CRM 143R CRM 483 CRM 484

Sewage sludge amended soil Sewage sludge amended soil Sewage sludge amended (Terra Rossa) soil Sewage sludge (domestic origin) Sewage sludge (mixed origin) Sewage sludge (industrial origin) Coal Fly ash Fly ash on artiﬁcial ﬁlters

BCR, Institute for Reference Materials and Measurements, Reference Materials Unit attn. BCR Sales, Retieseweg, B-2440 Geel, Belgium. Tel.: þ32-(0)14-571-704, fax: þ 32-(0)14-590-406, E-mail: [email protected]

Ash (industrial incinerator) Ash (municipal incinerator) Paint sludgePlating sludge no. 1 (copper electroplating) Electroplating sludge no. 2 Plating sludge no. 3 (chrome plating)

RT Corporation, 2931 Soldier Springs Road, P.O. Box 1346, Laramie, Wyoming 82070. Tel.: 307-742-5452, fax: 307-745-7936, E-mail: [email protected]

CRM 144R CRM 145R CRM 146R CRM 040 CRM 038 CRM 128 RT012 RT019 RT006 RT009 RT010 RT011

Sample preparation for industrial waste analysis

Code

959

P. Drouin and R. Clement

digestion. The purchase of a microwave digestion system represents signiﬁcantly higher capital expenditure than hot plate/block systems, and highpressure digestion vessels are also more expensive than the beaker/ﬂask/tube equipment used for hot plate/block systems. Most commercial closed vessel microwave digestion systems can only accommodate 8–12 digestion vessels, which limit the sample throughput in commercial analytical laboratories. The capital cost and sample limit capacity of current microwave systems could potentially be offset by increases in operational efﬁciency because a single microwave digestion may replace multiple hot plate/block digestions currently required to recover different elements (e.g. Hg) from a single sample. As both method detection limits and regulatory limits reach lower levels, sample digestion techniques capable of minimizing contamination and producing precise digestions will become increasingly important. The TCLP procedure is useful for evaluating industrial waste materials to determine if they are potentially harmful to the environment. The TCLP method was designed to mimic the environment inside a municipal solid waste dump and its applicability to other types of landﬁlls has been questioned. Selecting an appropriate leach procedure for a speciﬁc waste site may not represent the ideal situation; however, this will have to be balanced with increasing regulatory complexities and the cost associated with ensuring compliance. The toxicity and the mobility of an element depend on its chemical form and therefore methods that are capable of speciating the chemical form of elements to accurately access potential environmental or human health impact are required [66,68]. The US EPA method 3060 is an example of an analytical procedure capable of speciating hexavalent chromium in solid sample types [18, 48]. Future development of methods speciﬁc to other elements and organometallic compound to properly assess the potential impact on the environment will be an area of ongoing research and development. Although speciation can be performed with instrumental techniques (X-ray absorption near edge structures, XANES), methods similar to US EPA method 3060 or involving multistep selective leach procedures will be developed [83].

28.8

USEFUL WORLD WIDE WEBSITES

A number of useful Internet sites have been developed to help disseminate information about the determination of metals in various environmental sample types. The US EPA Ofﬁce of Solid Waste website (http://www.epa.gov/ osw/) contains all of the SW-846 method listed and can be downloaded in pdf format. The American Society for Testing and Materials ASTM International website can be found at http://www.astm.org. The European Environmental Agency website is designed to help transfer provision of timely, targeted, relevant, and reliable information to policy-making agents and the public

960

Sample preparation for industrial waste analysis

(http://www.eea.eu.int/). Kingston’s SamplePrep website (http://www. sampleprep.duq.edu/) is an excellent source of information on microwave digestion techniques, acid decomposition and safety. Cheatham’s PlasmachemL is a Listserver dedicated to plasma chemistry (ICP-OES and ICP-MS), serving as an electronic bulletin board where subscribers can post question and receive responses from other member of the Listserver. While not strictly related to sample preparation, problems and solutions related to preparation of samples for analysis by ICP-OES and ICP-MS are frequently discussed. The Plasmachem Listserver postings are archived and can be searched (http:// listserv.syr.edu/archives/plasmachem-l.html). The European Network for Harmonization of Leaching/Extraction Tests is a project designed to help leaching and extraction tests used by various European Union countries to help consolidate the different procedures (http://www.leaching.net/). LGC Promochem is an independent company that produces and distributes CRMs (formed by the merger of LGC’s reference materials business and Promochem Group of Companies) (http://www.lgcpromochem.com). Other websites of CRM producers include NIST (http://www.ts.nist.gov) and CANMET (http://www. nrcan.gc.ca/mms follow links to Canada’s Canadian Certiﬁed Reference Materials Project, CCRMP) and NRC (http://inms-ienm.nrc-cnrc.gc.ca follow links to research and development and then to chemical metrology).

Acknowledgments The authors acknowledge the following people for their help in the preparation of this chapter: T. Crawford for her assistance in procuring references; J. Caron, G. Kanert, and R. Moody for useful conversations concerning the TCLP method. REFERENCES 1 2 3 4 5 6 7 8 9 10 11

D.D. Link, P.J. Walter and H.M. Kingston, Environ. Sci. Technol., 33 (1999) 2469. D.D. Link, P.J. Walter and H.M. Kingston, Environ. Sci. Technol., 32 (1998) 3628. A. Robbat Jr. and R.L. Simpson III, Fresenius J. Anal. Chem., 364 (1999) 305. D. Florian, R.M. Barnes and G. Knapp, Fresenius J. Anal. Chem., 362 (1998) 558. J.B. Borba da Silva, M.A. Mesquita da Silva, A.J. Curtius and B. Welz, J. Anal. At. Spectrom., 14 (1999) 1737. M. Bettinelli, G.M. Beone, S. Spezia and C. Bafﬁ, Anal. Chim. Acta, 424 (2000) 289. M. Chen and L.Q. Ma, Soil Sci. Soc. Am. J., 65 (2001) 491. J. Liu, R.E. Sturgeon, V.J. Boyko and S.N. Willie, Fresenius J. Anal. Chem., 356 (1996) 416. G.E.M. Hall and J.C. Pelchat, J. Anal. At. Spectrom., 12 (1997) 103. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 3005, Revision 1, 1992. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 3010A, Revision 1, 1992.

961

P. Drouin and R. Clement 12 13 14 15 16 17 18 19 20 21 22 23 24

25

26 27

28 29 30 31 32 33 34 35 36 37 38 39

962

US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 3015A, Revision 1, 1998. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 3020A, Revision 1, 1992. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 3031, Revision 0, 1996. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 3050B, Revision 1, 1998. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 3051A, Revision 2, 1996. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 3052, Revision 0, 1996. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 3060A, Revision 1, 1992. C.Y. Zhou, M.K. Wong, L.L. Koh and Y.C. Wee, Anal. Chim. Acta, 314 (1995) 121. I. Lavilla, B. Pe´rez-Cid and C. Bendicho, Fresenius J. Anal. Chem., 361 (1998) 164. K.E. Levine, J.D. Batchelor, C.B. Rhoades Jr. and B.T. Jones, J. Anal. At. Spectrom., 14 (1999) 49. E.M.L. Lorentzen and H.M. Kingston, Anal. Chem., 68 (1996) 4316. C.J. Mason, M. Edwards and P. Riby, Analyst, 125 (2000) 327. Ontario Ministry of the Environment, The Determination of Metals in Final Efﬂuents, Industrial Waste and Landﬁll Leachates by Inductively Coupled PlasmaAtomic Emission Spectrometry (ICP-AES), 2002. Ontario Ministry of the Environment, The Determination of Mercury in Liquid Waste, Landﬁll Leachate and Sewage Samples by Cold Vapour-Flameless Atomic Absorption Spectrophotometry (CV-FAAS), 2002. US EPA, Test methods for Evaluating Solid Waste (SW-846), Method 7470A, Revision 1, 1994. Ontario Ministry of the Environment, The Determination of Metals in Solid Industrial Wastes by Inductively Couple Plasma-Atomic Emission Spectrometry (ICP-AES), 1999. Y. Hu, F. Vanhaecke, L. Moens, R. Dams, P. del Castilho and J. Japenga, Anal. Chim. Acta, 373 (1998) 95. A.R. Crosland, F.J. Zhao, S.P. McGrath and P.W. Lane, Commun. Soil. Sci. Plant Anal., 26 (1995) 1357. H. Ho¨dreja¨rv and A. Vaarmann, Anal. Chim. Acta, 396 (1999) 293. ˇ izˇma´rova´ and P. Ne˘mec, Commun. Soil. Sci. J. Zbiral, P. Medek, V. Kuba´nˇ, E. C Plant Anal., 31 (2000) 2045. T. Myo¨ha¨nen, V. Ma¨ntylahti, K. Koivunen and R. Matilainen, Spectrochim. Acta Part B, 57 (2002) 1681. P. Masson, A. Gomez and A. Tremel, Commun. Soil. Sci. Plant Anal., 27 (1996) 109. N. Cook, M.C. Turmel and W.H. Hendershot, Soil Sci. Soc. Am. J., 64 (2000) 609. H. Zhang and J.A. Hattey, Commun. Soil. Sci. Plant Anal., 31 (2000) 2959. J.F.N. Maaskant, A.H. Boekholt, P.J. Jenks and R.D. Rucenski, Fresenius J. Anal. Chem., 360 (1998) 406. M.F. de Abreu, R.S. Berton and J.C. de Andrade, Commun. Soil. Sci. Plant Anal., 27 (1996) 1125. B. Xing and P.L.M. Veneman, Commun. Soil. Sci. Plant Anal., 29 (1998) 923. M. Chen and L.Q. Ma, Soil Sci. Soc. Am. J., 65 (2001) 491.

Sample preparation for industrial waste analysis 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70

Z. Mester, M. Angelone, C. Brunori, C. Cremisini, H. Muntau and R. Morabito, Anal. Chim. Acta, 395 (1999) 157. D. McGrath, Talanta, 46 (1998) 439. J. Kucˇera, V. Sychra and J. Koubek, Fresenius J. Anal. Chem., 360 (1998) 402. M. Chen, L.O. Ma and G. Harris, Soil Sci. Soc. Am. J., 66 (2002) 632. K. Ashley, Trac, 17 (1998) 366. I. Arambarri, R. Garcia and E. Milla´n, Analyst, 125 (2000) 2084. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 7196A, Revision 1, 1992. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 7199, Revision 0, 1996. D. Huo, Y. Lu and H.M. Kingston, Environ. Sci. Technol., 32 (1998) 3418. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 1311, Revision 0, 1992. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 1312, Revision 0, 1994. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 1320, Revision 0, 1992. ASTM, Annual Book of ASTM Standards, Water Environ. Technol., Standard Test Method for Shake Extraction of Solid Waste with Water (D 3987-85), 1994. D.J. Hassett, 15th Annual Waste Testing & Quality Assurance Symposium, 1999, p. 66. J. Viguri, R. Iban˜ez, A. Andres, L. Ortiz and A. Irabien, Environ. Technol., 20 (1999) 171. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Method 1310A, Revision 0, 1992. US EPA, Proceeding of the Environmental Protection Agency Public Meeting on Waste Leaching, 1994. US EPA, Test Methods for Evaluating Solid Waste (SW-846), Ch. 7, Revision 3, 1996, p. 15. Ontario Ministry of the Environment, Environmental Protection Act, Regulation 558, Schedule 4, 2000. J. Ferna´ndez-Sempere, M.L. Barrueso-Martı´nez, R. Font-Montesinos and M.C. Sabater-Lillo, J. Hazard. Mater., 54 (1997) 31. R.S. Sletten, M.M. Benjamin, J.J. Horng and J.F. Ferguson, Water Res., 29 (1995) 2376. M.C. Chuan and J.C. Lu, Water Res., 30 (1996) 932. E.E. Chang, P.C. Chiang, P.H. Lu and Y.W. Ko, Chemoshpere, 45 (2001) 91. R.W. Peters, J. Hazard. Mater., 66 (1999) 151. J.R. Hale, A. Foos, J.S. Zubrow and J. Cook, J. Soil Contam., 6 (1997) 371. C. Hardaway, K. Gauthreaux, J. Sneddon and J.N. Beck, Microchem. J., 63 (1999) 398. I. Phillips and L. Chapple, J. Soil Contam., 4 (1995) 311. X. Chen, J.V. Wright, J.L. Conca and L.M. Peurrung, Water, Air, Soil Pollut., 98 (1997) 57. T.L. Rinehart, D.G. Schulze, R.M. Bricka, S. Bajt and E.R. Blatchley III, J. Hazard. Mater., 52 (1997) 55. I.B. Singh, M. Mudgal, A. Mishra, D. Dawar and B. Chakradhar, Indian J. Chem. Technol., 4 (1997) 298. M.C. Ruiz, A. Andres and A. Irabien, Environ. Technol., 21 (2000) 891.

963

P. Drouin and R. Clement 71 72 73 74 75 76 77 78 79 80 81 82 83

964

D.S. Kendall, 15th Annual Waste Testing & Quality Assurance Symposium, 1999, p. 72. Ming-Chi Wei, Ming-Yen Wey, Jiann-Harng Hwang and Jyh-Cherng Chen, J. Hazard. Mater., 57 (1998) 154. M.R. Iba´n˜ez, A. Andre´s, J.R. Viguri, I. Ortiz and J.A. Irabien, J. Hazard. Mater., A79 (2000) 215. J.O. Eckert Jr. and Q. Guo, J. Hazard. Mater., 59 (1998) 55. E. Egemen and C. Yurteri, Waste Manage. Res., 14 (1996) 43. X.D. Li, C.S. Poon, H. Sun, I.M.C. Lo and D.W. Kirk, J. Hazard. Mater., A82 (2001) 215. H.G. Snyman, Water Sci. Technol., 44 (2001) 107. R. Brobst, EPA Region VIII, Biosolids Management Handbook Part II, 1994, p. 2.3-2. A. Ramesh and J.A. Kozin´ski, Environ. Pollut, 111 (2001) 225. X. Chen, J.V. Wright, J.L. Conca and L.M. Peurrung, Water, Air, Soil Pollut., 98 (1997) 57. M. Janusa, J.C. Bourgeois, G.E. Heard, N.M. Kliebert and A.A. Landry, Microchem. J., 59 (1998) 326. In: R.E. Clement, L.H. Keith and K.W.M. Siu (Eds.), Reference Materials for Environmental Analysis. CRC Press/Lewis Publishers, Boca Raton, FL, 1997. V.J. Zatka, J.S. Warner and D. Maskery, Environ. Sci. Technol., 26 (1992) 138.

Chapter 29

Sample preparation for semiconductor materials Katsu Kawabata, Yoko Kishi, Fuhe Li and Scott Anderson

29.1

INTRODUCTION

As integration of semiconductor devices continues to increase and junctions become shallower, contamination control of chemicals and materials used in the semiconductor industry is becoming more critical and important. The International Technology Roadmap for Semiconductors (ITRS) [1,2] has been commonly used as an indication of trends in the semiconductor industry. In 2001, the ITRS suggested that critical metallic impurity levels in the ultrapure water (UPW) and the process chemicals should be lower than 20 and 10 ppt, respectively, and that the required impurity level of critical surface metals on Si wafers should be lower than 1 £ 1010 atoms/cm2. These requirements will be further lowered in the near future, as is shown in Table 29.1, and a new manufacturing process, that uses a larger variety of chemicals shown in Table 29.2, will be required to meet the future device geometry. Chemical impurities remain on a wafer to be later trapped in a device which can lead to defects that are shown in Table 29.3 [3]. There are several analytical techniques used to determine impurities in chemicals and materials used in the semiconductor industry. Total reﬂection X-ray ﬂuorescence (TXRF) has been used for in-line monitoring of surface contamination on Si wafers because of its non-destructive measurement capabilities. However, the impurity levels required for many semiconductor applications are near the detection capability of TXRF. Vapor Phase Decomposition (VPD) has been commonly used as a sample preconcentration technique for surface analysis of Si wafers [4,5]. A small amount of the sample solution obtained by the VPD technique is analyzed by TXRF, Inductively Coupled Plasma Mass Spectrometry (ICP-MS), or Graphite Furnace Atomic Absorption Spectrometry (GF-AAS). Process chemicals, which are by nature, solutions, can be analyzed by ICP-MS, GF-AAS and Inductively Coupled Plasma Optical Emission Spectrometry (ICP-OES). Each technique has its characteristic strengths and weaknesses. GF-AAS can be used to determine Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

965

K. Kawabata et al. TABLE 29.1 Technology requirements for wafer environmental contamination control—near-term [1] Year of production

DRAM 1/2 pitch (nm) MPU/ASIC 1/2 pitch (nm) Critical particle size (nm) Ultrapure Water Total Silica (ppb) Critical cation, anion, metals (ppt, each) Liquid chemicals HF, H2O2, NH4OH: Fe, Cu (ppt, each) Critical cation, anion, metals (ppt, each) HCl, H2SO4: all impurities (ppt) BEOL solvents, Strippers: K, Li, Na (ppt, each)

2001

2002

2003

2004

2005

2006

2007

130 150

115 130

100 107

90 90

80 80

70 70

65 65

65

58

52

45

38

35

33

0.1 ,20

0.1 ,20

0.1 ,20

0.1 10

0.05 10

0.05 10

0.05 10

,150

,135

,110

,100

,90

,50

,50

,10

,10

,10

,5

,5

,5

,1

,1000

,1000

,1000

,1000

,1000

,1000

,1000

,1000

,1000

,1000

,1000

,1000

,1000

,1000

very small amounts of sample and have good detection limits for most of the elements, but cannot determine many elements simultaneously. ICP-OES can be used to determine many elements simultaneously and handle samples containing higher dissolved solids directly, however, the detection limits, which are typically at ppb levels, are not good enough for pure chemicals. ICP-MS is getting accepted as the technique for both process chemicals and wafers since it can detect nearly all the metallic elements at the ppt level or lower. However, traditional ICP-MS is not a universal technique because of interference and matrix suppression issues. In order to overcome interferences arising from isobaric and polyatomic ions, several techniques have been developed. ICP-MS with a quadrupole mass ﬁlter (ICP-QMS) can be used with the cool plasma technique to reduce interferences [6] while higher resolution ICP-MS (HR-ICP-MS) uses a magnetic sector that literally

966

Sample preparation for semiconductor materials TABLE 29.2 Chemicals and materials commonly used in the semiconductor industry Process

Chemical compounds

Function

Poly-Si manufacturing

HCl, SiHCl3

Si epitaxial SiO2 epitaxial TiN CVD Ta2O5 CVD Quartz BPSG, PSG Pure metal SC-1 or APM cleaning SC-2 or HPM cleaning DHF cleaning DHF/HCl cleaning

SiH4 TEOSa TiCl4, TDMATb, NH3 Ta2O5, Ta(OC2H5)5 SiH4, SiH2Cl2 SiH4, PH3, B2H6 Al, Cu NH4OH, H2O2, DIW HCl, H2O2, DIW HF, DIW HF, HCl, DIW

Piranha or SPM cleaning Drying Buffered Oxide Etching or BOE Nitride Etching Silicon Etching or HNA

H2SO4, H2O2

Source of Si wafer, Solar cell, TFT, LCD Si deposition SiO2 deposition TiN deposition Ta2O5 deposition Furnace tube Passivation Contact and interconnect Remove particles Remove metallic impurities Remove SiO2 Remove SiO2 and metallic impurities Remove organic impurities

IPA NH4F, HF, DIW

Remove water Etch SiO2

H3PO4 HF, HNO3, CH3COOH, DIW HF, HNO3, DIW HCl, HNO3 HCl, HNO3 H3PO4, HNO3, CH3COOH, DIW PR, NMP, PGME, PGMEA, NBA TMAH, NMP H2SO4, NMP, DMSO ARC

Etch Si3N4 Etch Si

Silicon Etching Aluminum Etching Gold Etching Metal Etching Lithography Resist Developer Resist Stripper Anti-reﬂective Coating Chemical Mechanical Polishing or CMP

Colloidal Silica

Etch Si Etch Al Etch Au Etch Al, Ni, Ag Make circuit pattern Develop resist Remove resist Reduce substrate reﬂectivity phenomena Make ﬂat surface

a

Tetraethoxysilane.

b

Tetrakis(dimethylamino)titanium.

967

K. Kawabata et al. TABLE 29.3 Defect on Si wafer [3] Contaminants Alkali and alkali-earth elements: Li, Na, K, Ca, Mg, Ba Transition and heavy metals: Fe, Cr, Ni, Cu, Zn, Au, Mn, Pb Doping elements: B, Al, P, Ga, As, In

Defect on Si based semiconductor devices Reduce the breakdown voltage Reduce the lifetime of carrier, causing higher dark current Shift the device operating point

separates analyte and interfering ions by using higher spectral resolution [7]. Since the cool plasma technique can eliminate some of the critical interferences encountered in semiconductor applications, such as Arþ, ArOþ, ArHþ and ArCþ, it has been widely used for routine analysis in the semiconductor industry. However, the cool plasma technique cannot overcome other important interferences when samples contain Si, Ti, H2SO4, H3PO4 and HCl matrices. In addition, the cool plasma technique suffers from matrix suppression and insufﬁcient decomposition of refractory elements such as Si, Ca, Ti, Zr, Mo and W due to the lower energy of the plasma. When used properly, the cool plasma technique requires either the elimination of the matrix or the use of matrix-matched standard solutions [8,9]. HR-ICP-MS can overcome most of these interferences, although combination with the cool plasma technique is recommended for analysis of single ppt levels of Na, Ca, K and Fe because it drastically reduces the background from the cones. A more recent innovation, ICP-QMS with a reaction cell system, can also be used to eliminate most of these interferences as well as Arþ, ArOþ, ArHþ and ArCþ interferences without the need of the cool plasma technique [10]. The sample preparation scheme used is dependent upon the analytical technique that is to be used to make the measurement. As noted earlier, elimination of the matrix will be required if an interference exists while using the cool plasma technique. On the other hand, only a simple dilution will be needed if the analytical instrument has the capability to eliminate the interference problem. In order to determine ultratrace levels of impurities in semiconductor chemicals and materials, contamination control is the most critical issue. This chapter describes how to avoid contamination arising from the environment and equipment during sample preparation, and what kind of sample preparation is recommended for ICP-MS analysis.

968

Sample preparation for semiconductor materials

29.2

CONTAMINATION CONTROL

29.2.1 Clean room Since airborne metallic and particulate impurities can cause defects in the ﬁnished product, semiconductor devices are typically manufactured in a cleanroom environment. Consequently, the sample preparation for chemicals and materials used in the semiconductor industry should be performed in a similar environment. The cleanroom is speciﬁed based on the number of particles of diameter of 0.5 mm or larger that exist in a cubic foot, as deﬁned by the US Federal Standard 209E. This standard is shown in detail in Table 29.4 [11]. While a class 1000 cleanroom should have fewer than 1000 particles of 0.5 mm size per cubic foot, it may contain particles as large as 5 mm. In a non-cleanroom environment, the number of particles is typically more than 1 million per cubic foot. These particles can contain Si, Al, K, Ca, Fe, Na, Mg and B. To demonstrate the impact of airborne contamination, let us envision one particle 5 mm in size dropping into a 100 ml solution. This particle will impart a concentration in the solution of approximately 5 pg/ml or 5 ppt. If that same particle drops into a 1 ml solution, the concentration of that solution is increased by 500 ppt. Depending on the concentration to be analyzed and the sample volume, a lab environment must be properly designed to ensure the cleanliness of the samples to be analyzed. In order to maintain a particle-free cleanroom environment, air passes through a mechanical ﬁlter device from the ceiling and exits through the bottom of the sidewall or through a grating in the ﬂoor. The air pressure in the clean room must be higher than that outside to avoid entrainment of unﬁltered air. For most semiconductor works, it is recommended that an

TABLE 29.4 Airborne particulate cleanliness classes in cleanrooms and clean zones, US Federal standard 209E [11] Class

1 10 100 1000 10,000 100,000

Measured particle size (mm) 0.1

0.2

0.3

0.5

5

35 350 NA NA NA NA

7.5 75 750 NA NA NA

3 30 300 NA NA NA

1 10 100 1000 10,000 100,000

NA NA NA 7 70 700

969

K. Kawabata et al.

analytical instrument be installed in at least a class 1000 cleanroom in which a class 100 or better clean draft is available for the sample preparation area. There are two types of mechanical ﬁlters that are used to create the clean environment: high-efﬁciency particle air (HEPA) ﬁlters and ultra low penetration air (ULPA) ﬁlters. The HEPA ﬁlter eliminates more than 99.99% of particles larger than 0.3 mm and the ULPA ﬁlter eliminates more than 99.999% of particles larger than 0.12 mm. Although the mechanical ﬁlter efﬁciently removes particulate matter, it cannot remove gaseous components. A HEPA ﬁlter made of borosilicate glass, for example, used in air containing HF vapor will generate gaseous boron compounds, e.g. BF3, which will produce boron contamination in the clean room, as shown in Table 29.5 [12–14]. Additionally, contamination by phosphorous from a sealant used in adhering ﬁlter media to support modules has been reported in the literature [15]. For these reasons, the ULPA ﬁlter made of polytetraﬂuoroethylene (PTFE) is strongly recommended for down-ﬂow hoods in the sample preparation area. Once a class 1000 or better grade of cleanroom is set up, the generation of particulate matter from an operator should be taken into account. It may be surprising to learn that the emission rate from normal clothing is about TABLE 29.5 Comparison of contaminants outside and inside a cleanroom [14] Compounds

Outside cleanroom (ng/m3)

Inside cleanroom (ng/m3)

Al B Ca Cu Fe K Mn Na Pb Zn NO3 2

190 17 450 4 230 170 110 510 17 46 8600 2100

,1 130 2.5 ,1 ,1 ,1 ,1 ,1 ,1 ,1 940 ,10

1600 ,100 ,100 3500

,10 ,10 ,10 2700

SO4 22 Cl2 F2 PO4 32 NH3

970

Sample preparation for semiconductor materials

100,000 particles per minute (p/min) when a person is sitting or standing motionless, about 500,000 p/min during hand, forearm and head movement, and up to 10,000,000 p/min during normal walking [16]. The proper use of cleanroom clothing is thus required to maintain the quality of the cleanroom environment. 29.2.2 Equipments, reagents and standards In order to analyze chemicals and materials at the ppt level of contamination, all materials such as plastic or quartz bottles, gloves and pipette that directly touch chemicals and materials should be considered as sources of contamination [17–26]. Special attention should be paid to the selection of equipment and thorough cleaning procedures should be followed prior to use. 29.2.2.1 Gloves Skin, hair, exhaled air and perspiration from a body are the largest contamination sources in the cleanroom. Consequently, handling of chemicals and materials must always be performed while wearing gloves. Several different types of gloves are commercially available, many are designed for different purposes, and attention must be paid to such things as their chemical property, particle generation, electrostatic discharge, and comfort of ﬁt. Powder-free gloves must be used in order to avoid particle contamination even though gloves with powder are easier to put on. Unfortunately, new gloves commonly used in a cleanroom have particles so that thorough washing with pure water is required after putting them on. The gloved hands should be washed diligently before handling samples because the gloves are easily contaminated when they are in use [26]. 29.2.2.2 Bottles and containers Quartz (QZ), polyethylene (PE), polypropylene (PP), polystyrene (PS), PTFE and perﬂuoroalkoxy (PFA) are materials commonly used as bottles and containers for semiconductor samples. QZ is the preferable material for sample preparation that requires higher temperature, but it is not a good choice when using hydroﬂuoric acid (HF). On the other hand, PE, PP, PS, PTFE and PFA can be used for HF, but these materials are not suitable when temperature changes are expected. Their porous surface causes interactions, such as adsorption, desorption, leaching and diffusion of elements between chemicals and material. PE, PP and PS materials often suffer from contamination by Al, Cr, Mg and Zn from catalysts used in production processes. Fortunately, PFA does not use any problematic catalyst during the manufacturing process so that a relatively high level of purity is obtainable. PFA is a preferable material for ppt level analysis, although B, Na, Al, K, Fe, Cr, Ni and Mo may be present from the stainless steel mold used in manufacturing. Four different PFA materials

971

K. Kawabata et al. TABLE 29.6 Grades of quartz [28] Material

Normal grade High purity grade

Al

Ca

Cr

Cu

Fe

K

Li

Mg

Na

,20 0.05

0.6 0.05

,0.02 0.01

,0.06 0.005

0.2 0.02

0.3 0.01

0.6 0.01

0.1 0.005

0.2 0.05

Unit: mg/g (ppm).

were tested for extractable metallic impurities and there was a signiﬁcant difference between the materials [27]. There are different purities of QZ materials, as shown in Table 29.6 [28]. All bottles used for sample preparation must be of the highest purity material and cleaned prior to use, although the nature of the cleaning procedure will depend on the chemicals to be analyzed. The following is a procedure used at one of the chemical manufacturers in Japan for QZ and PFA bottles [27]. 1. soak in (1 þ 1) HCl for one week; 2. soak in (1 þ 1) HNO3 for one week; 3. heat with the chemical to be stored for 3 h (under the boiling point of the chemical); 4. heat with UPW for 3 h (under the boiling point of UPW). After completing the cleaning procedure described above, the bottles can be used for storing ultra pure chemicals at room temperature; however, they should not be used with any preconcentration procedures. When these bottles were used for the preconcentration of HF, a few tenth ppt levels of Na, Al and Fe contamination were found, even after the cleaning. The contamination levels ﬁnally decreased to a few ppts after it had been used 14 times [27]. If bottles or containers are used for digestion or preconcentration, several repetitive cleaning procedures with heat are required if a single ppt level analysis is to be attempted. If bottles or containers are used only for dilution of chemicals, heating steps 3 and 4 can be replaced with soaking for several days with the chemical to be used, followed by soaking with UPW. All cleaned bottles and containers must be thoroughly washed with plenty of UPW prior to use. All cleaned bottles must be used only for ultra trace level analysis and each bottle should have a logged identiﬁcation in order to avoid cross-contamination. It should be noted that a label with adhesive attached to PFA bottle causes contamination due to permeation of adhesive vapor into the bottle [27]. Carving a number on the bottle is, therefore, recommended.

972

Sample preparation for semiconductor materials

29.2.2.3 Volumetric ﬂask A volumetric ﬂask is commonly used for dilution of chemicals or preparation of standard solutions and the same attention should be paid to their cleaning as that for bottles. A volumetric ﬂask can be used for a sub-ppb level of standard solution, but is not recommended for sample preparation of semiconductor chemicals and materials. As noted earlier, the atmosphere and human beings are the source of contamination, which usually enters an open sample vessel between the cap and the mouth. When a sample solution is poured into a volumetric ﬂask, the sample solution readily touches the mouth and washes the contamination down into the ﬂask. In addition, a volumetric ﬂask should be shaken rather than swirled after ﬁlling up to the graduated line in order to have better mixing, although it causes the contamination from the cap. For this reason, a gravimetric method using a wide-mouthed bottle is recommended for the preparation of standard solutions at the low ppt level. A detailed preparation procedure will be discussed in Section 29.3. 29.2.2.4 Pipette To avoid Na, B and Si contamination, a glass pipette must not be used for the preparation of semiconductor chemicals and materials. A pipette with disposable plastic tips is the preferred tool. Some tips use coating material to have better delivery of solution, but it can be a source of contamination. A metal free colorless tip made of polypropylene is commercially available, but it is not clean enough for sampling of pure chemicals, especially higher concentrations of acid and alkali. Such a pipette tip should be used only for the sampling of standard solutions. Recently, a pipette tip made of PFA has become commercially available which might, after thorough cleaning, be used for sampling of pure chemicals. It is recommended that the PFA tip be dedicated to just one chemical and not used with other chemicals to minimize sample cross-contamination. There are several ranges of variable volume pipettes on the market, the most convenient being from 10 to 100 ml or 100 to 1000 ml. Although much smaller volumes, such as from 1 to 10 ml are available, it is often difﬁcult to achieve complete release of the solution, due to its surface tension. A micropipette uses a stainless steel plunger, which, over time, might be corroded by acid vapor and cause contamination with the metallic components. Soaking a pipette tip in 1% HNO3 is good practice to minimize contamination from the tip. The tip should not be soaked for more than a few days because the low adhesion properties of the tip surface will be degraded, which causes memory of the solution in the tip. As mentioned earlier, the gravimetric method is preferred in order to avoid contamination, and the use of a pipette should be limited to the standard solution. The cleaning of the tip with deionized water (DIW) just before the use

973

K. Kawabata et al.

is acceptable in many cases, but the following cleaning procedure is recommended for the preparation of a single ppt level calibration standard: If an 80 ml volume of standard solution is sampled, for example, – set the volume of the pipette initially at 100 ml; – wash the tip by aspirating a 1% HNO3 solution several times; about 2/3 of the tip should be inserted in the wash solution; – wash the tip by aspirating DIW several times; about 2/3 of tip should be inserted in the wash solution to wash the outside of the tip; – set the volume of the pipette at 80 ml; – wash it by aspirating the standard solution once; the tip must not be inserted in the solution deeper than 1/2 of the tip to avoid contamination from the outside of the tip; – aspirate the standard solution; – dispense the standard solution into a sample bottle. The tip must not touch any part of the bottle. When a PFA bottle is used, the tip should be inserted deeper than the neck of the bottle to avoid the static electricity phenomenon. If the tip is not deep enough and the sample is dispensed, droplets of sample solution might be attracted toward the neck of the bottle. 29.2.2.5 Balance To avoid contamination of a sample solution, dilution by the gravimetric method is the best approach, since a pipette might cause a contamination problem, especially when dispensing highly concentrated reagents. A balance with an accuracy of one-hundredth gram is recommended. 29.2.2.6 Reagents and standards Reagents, including pure water and standard solution used for the analysis of semiconductor chemicals and materials, should be of very high quality. Pure water is used for dilution of chemicals and standard solutions. There are several pure water systems on the market. Distilled water that uses evaporation and condensation of water is the lowest cost solution. Unfortunately, this method cannot effectively eliminate impurities because water is evaporated at the boiling point and most of the impurities are vaporized with the pure water and subsequently condensed. DIW obtained by ion exchange techniques shows impurity levels lower than 1 ppt for most of the elements, as shown in Table 29.7, and it is the ﬁrst choice for most applications. The cost of DIW is usually higher than for distilled water. DIW can be further puriﬁed by means of a sub-boiling system that evaporates the DIW below the boiling point, avoiding evaporation of impurities. The purity of the water can be better than DIW, but it takes time for the initial cleaning of the system and produces only a few liters of pure water per day.

974

Sample preparation for semiconductor materials TABLE 29.7 Pure water (Milli-Q Element) results Element

Concentration found (ppt)

Element

Concentration found (ppt)

Li Be B Na Mg Al K Ca Ti V Cr Mn Fe Ni Co Cu Zn Ga

,0.2 ,0.9 ,0.8 ,0.2 ,0.2 ,0.1 ,3 ,0.1 ,2 ,0.04 ,0.1 ,0.5 ,0.4 ,0.2 ,0.04 ,0.1 ,1 ,0.05

Ge As Sr Zr Mo Ag Cd In Sn Sb Ba Ta W Au Tl Pb Bi U

,0.6 ,2 ,0.02 ,0.04 ,0.1 ,0.1 ,0.1 ,0.02 ,0.9 ,0.08 ,0.04 ,0.05 ,0.07 ,0.05 ,0.01 ,0.09 ,0.01 ,0.01

Obtained by DRC-ICP-MS.

Nitric acid is the most commonly used acid for preparation of samples and standard solutions. It is also added to weak acids or alkali solutions to stabilize elements in the sample and to avoid adsorption onto the plastic sample bottles. Since its impurity level affects the practical detection capability, electronic grade HNO3 should be used. Several acid manufacturers guarantee the quality of HNO3 at better than 10 ppt for each element [29,30]. A single stock standard element solution designed for AAS has been used for making calibration standard solutions. It is only designed to certify the concentration of the element but it is not speciﬁed or controlled for other impurities. When AAS grade single standard solutions are mixed and used for multi-element analysis, cross-contamination, especially by B, Na, K, Ca and Fe, is commonly found. This problem can be easily checked by comparing the single standard solution and the multi-element standard solution. To avoid this problem, a single stock element solution that limits the impurity of other elements should be mixed to make a multi-element standard solution. A multi-element standard solution designed for ICP-MS is also commercially available [31,32].

975

K. Kawabata et al.

29.3

SAMPLE PREPARATION

The best way to avoid contamination during analyses is to analyze samples directly, without any sample preparation. However, sample preparation is often required in order to overcome the following issues: – some samples add severe matrix effects and complex spectral overlapping problems to an analysis. In this case, sample matrix needs to be eliminated before an accurate analysis can be performed by ICP-MS; – the detection limits that are attainable by directly analyzing samples are not low enough for some applications, and preconcentration of analytes prior to the analysis is needed; – certain types of samples cannot be directly introduced into the ICP for accurate mass spectrometric analysis. As noted earlier, sample preparation, regardless of analyte preconcentration or matrix elimination, should be carefully performed in a clean environment with a closed system rather than an open one. A closed evaporation system basically applies the same principle as the sub-boiling system using a ﬁltered gas or a vacuum system to evacuate vaporized material and exhaust it to the atmosphere or a scrubber [33 –36]. Such a closed system, used for evaporation or puriﬁcation of chemicals, consists of high purity PFA containers with caps, a heating device and a temperature controller. Table 29.8 represents the ppt-level spike recovery data obtained by evaporating three acids commonly used in semiconductor manufacturing [33]. Most of the elements show good spike recoveries, regardless of the type of acid. Some elements form volatile species, which are lost during the evaporation. For example, B and Hg are readily lost when evaporating these acids; Ge and Sb form volatile chlorides in HCl and are difﬁcult to recover during the evaporation of HCl. The volatility of B and Hg can be controlled by making complexation with mannitol [37] and the addition of gold [38], respectively. However, addition of these chemicals may cause contamination errors and, consequently, degrade the desirable detection limits that are attainable if the reagents are not pure enough. For that reason, direct ICP-MS analysis or simple dilution followed by ICP-MS analysis is always preferable for liquid chemicals if the required detection limits can be achieved. Solid materials are usually digested by pure chemicals prior to analysis by ICP-MS. A closed digestion system is also preferable in order to avoid possible contamination during the digestion. However, a high pressure closed microwave digestion technique is not recommended for ultra trace level analysis because the high pressure opens up the porous surface of PFA vessels and leaches out contaminants from the vessel itself. The leaching process can last for a very long time. Additionally, the high pressure can also push some analytes into the interior wall of the vessels, causing analyte loss.

976

Sample preparation for semiconductor materials TABLE 29.8 Spike recovery (%) data with Evacoclean 12P35 [33] Element

49% HF

36% HCl

69% HNO3

Al Sb As Ba Be Bi Cd Ca Cr Co Cu Ga Ge Au Fe Pb Li Mg Mn Mo Ni Nb K Ag Na Sr Ta Tl Sn Ti V Zn Zr Spike concentration

78 98 96 98 89 98 100 83 98 99 101 99 99 99 95 95 – 103 98 94 85 99 88 91 83 99 117 97 83 85 95 99 96 50 ppt

102 – 92 100 101 98 99 97 107 92 94 92 – 95 105 99 118 108 100 87 81 93 100 95 91 95 109 99 100 82 91 88 95 100 ppt

89 94 86 106 98 101 92 94 103 96 93 95 95 98 82 104 88 83 104 97 101 96 101 99 104 98 99 105 99 78 81 96 500 ppt

977

K. Kawabata et al.

The maximum matrix concentration after digestion of the solid depends on the plasma temperature and the elements of the matrix. Gaseous samples may be analyzed directly by ICP-MS [39], but the standardization is difﬁcult due to the limited availability of gaseous standards. Trapping by a Teﬂon membrane ﬁlter, followed by dissolution with acids, is a convenient approach for removing metallic impurities from particulate matter in gases [40]. However, the Teﬂon membrane ﬁlter is normally located at a point of use in a manufacturing line and impurities that pass through the ﬁlter are of greater concern. For that reason, a bubbling technique is used as an alternative approach. Gasses are passed through an acid solution and metallic impurities are trapped in the acid [41]. Although the trapping efﬁciency is difﬁcult to assess, this method has been widely used for the evaluation of cleanroom environments. 29.3.1 Preparation and analysis of samples Once samples with adequate matrix concentration are prepared, as is described later, they can be analyzed by ICP-MS. ICP-MS is not an absolute analytical technique and needs to be calibrated with standards before quantitative analysis is performed. There are two calibration methods typically used: external calibration and standard additions calibration. 29.3.1.1 External calibration Calibration curves are obtained by analyzing standard solutions typically prepared in a few percent HNO3, by which the quantitative results for samples are calculated from the relative sensitivity. External calibration curves are used for analysis of other chemicals and generally prove to be the most convenient method. When using a single external calibration for a variety of chemical matrices, it is important to verify the sensitivity in these matrices is similar if accurate results are to be obtained. Additionally, hotter plasma conditions are preferable to minimize any matrix suppression arising from different matrices [42]. 29.3.1.2 Standard addition calibration When using this method, a standard solution is directly added to a sample and quantitative results are obtained by measuring the background equivalent concentrations (BECs). This technique is basically not affected by the sample matrix and allows accurate results when a background signal from the analytical instrument is negligible. When the background signal arising from the analytical instrument is high relative to the analyte signals, a positive bias occurs because the result is partially contributed by the background. This problem is especially critical for analysis using the cool plasma technique. In this case, the standard additions method should be separately used for both the acid and the digested samples with subsequent subtraction of the two

978

Sample preparation for semiconductor materials

concentration results. In most situations, the standard additions method should be performed for each sample, which makes this a time-consuming and labor intensive procedure. 29.3.1.3 Preparation procedure of calibration standard The manner of preparation of calibration standards varies depending on the concentration of the calibration standard, cleanliness of bottles, and sample handling skill. Since the multi-element stock standard solution used for ICP-MS is prepared at a higher concentration, e.g. 10 ppm, it should be ﬁrst diluted to obtain an adequate intermediate concentration. If the concentration of the calibration standard planned is 100 ppt, the preparation of a 100 ppb intermediate stock standard is recommended. There are two ways to prepare the standard solutions, depending on how many bottles are used. 29.3.1.4 Use of several bottles – weigh a larger empty bottle, e.g. 500 ml; – add DIW to the bottle directly from the DIW system; DIW must not touch the mouth of the bottle and the volume should occupy about half the volume of the bottle; – reweigh the DIW bottle and calculate the amount of DIW in the bottle; – add an adequate amount of HNO3 to the bottle directly without using a pipette; it must not touch the mouth of the bottle and a few percent of HNO3 is recommended; – reweigh the bottle and calculate the amount of HNO3 added; – close the cap of the bottle and swirl the contents carefully and thoroughly, do not shake the bottle in order to avoid contamination from the cap and mouth; this solution is used as diluent for the preparation of calibration standard solutions; – weigh several empty bottles, e.g. four 100 ml bottles, for the calibration standards; – pour the HNO3 diluent solution into each empty bottle; it must not touch the mouth of the bottle and it should ﬁll about half the volume of the bottle, e.g. 50 g; – weigh each bottle; – add an adequate amount of intermediate multi-element standard solution to each bottle according to the weight of HNO3 diluent solution in each bottle. If a 100 ppt standard solution is prepared and the weight of HNO3 diluent solution is 50 g, add 50 ml of a 100 ppb multi-element standard solution: 50 ml of standard solution is about 0.05 g, which is negligible compared with the 50 g sample. Each bottle represents a different concentration level of the standard; – close the cap of the bottle and swirl the contents thoroughly.

979

K. Kawabata et al.

This is the most common and conventional way of preparing standard solutions over a few tenth ppt levels. The contamination from each bottle will be critical for a single ppt level analysis, therefore, the bottles must be cleaned prior to use. 29.3.1.5 Use of one bottle – weigh empty bottle; – add DIW to the bottle directly from DIW system; DIW must not touch the mouth of the bottle and the volume should occupy about half the volume of the bottle; – reweigh the DIW bottle and calculate the amount of DIW in the bottle; about 50 –100 g is recommended, depending on a sample uptake rate of the ICP-MS; – add an adequate amount of HNO3 directly to the bottle without using a pipette; it must not touch the mouth of the bottle; – reweigh the bottle and calculate the amount of HNO3 added; – close the cap of the bottle and swirl the contents carefully and thoroughly, do not shake the bottle in order to avoid contamination from the cap and mouth; this solution is used as the calibration blank solution; – aspirate the blank solution into the instrument; – after the analysis, measure the weight of the blank solution; – add an adequate amount of intermediate multi-element standard solution to the blank solution; this solution is the lowest concentration of calibration standard; – close the cap of the bottle and swirl the contents thoroughly; – aspirate the standard solution into the instrument; – repeat the procedure until all concentration levels of calibration standards are measured. This is the best procedure to obtain a single ppt level calibration curve because variation of contamination from each bottle can be avoided. However, each standard solution must be individually prepared after each measurement. This procedure is used for the method of standard additions. 29.3.1.6 Preparation procedure for semiconductor reagents and materials Each reagent or material requires a different sample preparation procedure and some examples are presented below. DIW DIW seems, at ﬁrst, to be the easiest reagent to analyze. However, DIW is among the most difﬁcult of samples in the list shown in Table 29.2 since it is the purest reagent and used as a blank solution. Since, as noted earlier, the ICP-MS is not an absolute analytical technique, the method of standard

980

Sample preparation for semiconductor materials

additions is the best means of direct analysis of DIW. Table 29.7 presents results obtained for analysis of DIW from a Milli-Q Element system (Bedford, MA, USA). Since there is no blank solution, the results are expressed as , BEC in ppt. In order to analyze impurities at much lower concentration levels, preconcentration via evaporation is required. In this case, DIW without preconcentration is used as the blank and the preconcentrated DIW is analyzed as a sample. It is recommended that a small amount of ultrapure HNO3 (about 1%) be added to UPW to dissolve and stabilize any elements that may be present in the form of oxides or hydroxide particulates and to minimize possible analyte adsorption during sample introduction and transport processes. H2O2 A 30% solution of H2O2 can be directly analyzed by ICP-MS. It should be noted that H2O2 becomes explosive when the concentration exceeds 50%. Therefore, preconcentration of H2O2 must not be performed. Since the dissociation constant of H2O2 is very small, like water, the addition of HNO3 is recommended to stabilize elements in solution. HNO3 A 35% solution of HNO3 can be directly analyzed by ICP-MS. Preconcentration of HNO3 is possible using an evaporation device in order to improve the detection limits, but possible loss of B and Hg should be taken into consideration. HCl A 20% solution of HCl can be directly analyzed by ICP-MS. When using a conventional ICP-QMS, the elimination of HCl is required for the determination of K, V, Cr, Ga, Ge, As and Se since Cl related polyatomic ions, such as ClH2, ClO, ClOH, ClO2, Cl2 and ArCl, interfere with the determination of these elements. The elimination of HCl can be done by means of an evaporation device, however, care should be taken to prevent the possible loss of B, Ge, As, Sb, Sn and Hg [43]. On the other hand, DRC-ICP-MS and HR-ICP-MS can eliminate most of these interferences and the elimination of HCl prior to the analysis may not be required [44], unless much lower detection limits are needed. HF A 25% solution of HF can be directly analyzed by ICP-MS. Preconcentration of HF is possible using an evaporation device in order to improve the detection limits, but possible loss of B, As, Se, Si and Hg should be taken into consideration [43].

981

K. Kawabata et al.

H3PO4 A 1% solution of H3PO4 can be directly analyzed by ICP-MS. Phosphorus related polyatomic ions, such as PO, PO 2 and PO2H, interfere with determination of Ti, Cu and Zn, respectively, using conventional ICP-QMS. The elimination of H3PO4 is difﬁcult because H3PO4 dehydrates with heat and crystallizes forming metaphosphoric acid at around 3008C, and it eventually sublimates at higher temperature. Therefore, the elimination of H3PO4 matrix is not readily achieved. HR-ICP-MS and DRC-ICP-MS can eliminate these interference issues [45]. H2SO4 A 10% solution of H2SO4 can be directly analyzed by ICP-MS. Sulfur related polyatomic ions, such as S2, SO, SOH, SO2 and ArS, interfere with determination of Ti, V, Cr, Zn, and Ge, respectively, using conventional ICPQMS. Evaporation and preconcentration of H2SO4 is possible using an evaporation device, but possible loss of B, As and Hg should be taken into consideration. HR-ICP-MS and DRC-ICP-MS can eliminate these interference issues [46]. NH4OH A 6% solution of NH4OH can be directly analyzed by ICP-MS. Since the dissociation constant of NH4OH is small, addition of HNO3 (about 1%) is recommended to stabilize elements in solution. Preconcentration of NH4OH is possible using an evaporation device, but possible loss of B and Hg should be taken into consideration. Tetramethylammonium Hydroxide (TMAH) A 5% solution of TMAH can be directly analyzed by ICP-MS. Preconcentration of TMAH is possible using an evaporation device, but possible loss of B and Hg should be taken into consideration. Isopropanol (IPA) and Acetic acid (HAc) Undiluted IPA and HAc can be analyzed by ICP-MS following addition of oxygen into the plasma to avoid carbon build up at the tip of the sampling and skimmer cones. The concentration of oxygen needed to oxidize the carbon is about 3% of total nebulizer gas ﬂow. If the addition of oxygen is not possible, a ﬁve-fold dilution with DIW is required. Since the dissociation constant of IPA and HAc is small, addition of HNO3 (about 1%) is recommended to stabilize elements in solution. Preconcentration of IPA and HAc is possible using an evaporation device, but possible loss of B, As, Hg and Pb should be taken into consideration. Since IPA is a very good dehydrating agent, it absorbs moisture from the atmosphere while transferring from one bottle to another. A clean room environment is strongly recommended.

982

Sample preparation for semiconductor materials

29.3.1.7 NMP, PGMEA, PGME, NBA, EL, DMSO and other organic solvents Many organic solvents have been used in the semiconductor industry for various purposes and it is impossible to consider all organic solvents in this section, so general concepts for their analysis are presented below. Common organic solvents include N-methyl pyrrolidone (NMP), propylene glycol monomethyl ether acetate (PGMEA), propylene glycol monomethyl ether (PGME), n-butyl acetate (NBA), ethyl lactate (EL), and dimethyl sulfoxide (DMSO). With the exception of NBA, all are water miscible and can be analyzed by ICP-MS after a ﬁve- to ten-fold dilution with DIW. These reagents can be analyzed by ICP-MS without dilution, but the sample uptake rate should be very low and the addition of oxygen (about 5% of total nebulizer gas ﬂow) is required as in the case of direct analysis of IPA. Since the vapor pressure of organic solvents is high, it is very easy to evaporate. However, some volatile organometallic compounds will be lost during the evaporation step. If other organic solvents are analyzed by ICP-MS, the following steps are general considerations: – if an organic solvent is water miscible, dilute it with 1% HNO3 and analyze; – if an organic solvent is not water miscible, analyze it directly with the addition of oxygen; – if the vapor pressure of the organic solvent is too high, dilute it with NMP or PGMEA and analyze it directly with the addition of oxygen; – if the viscosity of the organic solvent is too high, dilute it with NMP or PGMEA and analyze it directly with the addition of oxygen; – if an organic solvent is not soluble in NMP or PGMEA, dilute it with xylene and analyze it directly with the addition of oxygen. When an organic solvent is diluted with NMP or PGMEA, an aqueous standard solution can be added to the diluted organic solvent. If an organic solvent cannot be diluted with NMP or PGMEA, dilution may be done using xylene; in this case, the aqueous standard solution cannot be used because it will be isolated from the organic solvent and an organic standard solution, such as Conostan [47], may be used. Another alternative is to use microwave digestion with HNO3. 29.3.1.8 Photoresist (PR) and anti-reﬂective coating (ARC) There are many different types of PR and ARC used in various applications. PR and ARC can generally be diluted with some organic solvents, such as NMP or PGMEA. Therefore, PR and ARC can be analyzed by ICP-MS with the addition of oxygen to the plasma after dilution with NMP or PGMEA. The maximum concentration of PR and ARC allowed in the ﬁnal sample solutions varies depending on their composition. In general, PR and ARC should be diluted to about 3 and 10%, respectively.

983

K. Kawabata et al.

When the concentration of PR is too high, carbon deposition occurs in the injector tube of the ICP-MS torch. Some of the organic chemicals used in the semiconductor industry contain S in their molecular structure, which will cause S related interferences like H2SO4 analysis on ICP-MS. 29.3.1.9 Si wafer Impurities in reagents used for cleaning and manufacturing of devices can be trapped in a thin surface oxide layer on the Si wafer. The impurities in the oxide layer represent one of the major contamination issues during semiconductor manufacturing. It is very important to monitor for those impurities and track down their contamination sources. There are two types of oxide layer: native and thermal oxides. The native oxide layer is generally a few tenths of an Angstrom thick whereas the thermal oxide is over a few thousand Angstroms thick. In order to digest the surface oxide layer, VPD has been developed as the preconcentration technique [48]. This technique has been widely used for both TXRF and ICP-MS analyses. The basic principle of VPD is as follows: a Si wafer is placed in a Teﬂon container and exposed to HF vapor. After the SiO2 layer decomposes in the HF vapor phase, a small volume, e.g. 200 ml, of mixed HF and H2O2 solution is dropped onto the Si wafer and scanned across the entire surface of the wafer. The impurities dissolved from the decomposed oxide layer are collected by the scanning process and then analyzed by either TXRF or ICPMS. There are several automatic wafer surface preparation systems on the market and successful results have been published [49]. Wafer scanning can also be done manually, the results obtained are comparable with those obtained using automatic systems, if not better. Compared to the automatic system, manual scanning appears to be much more cost effective. A small volume of mixed HF and H2O2 solution can also be used to selectively etch the oxide layer on the Si wafer, without exposing the surface to a HF vapor, since HF reacts with SiO2 but not Si. The resultant small volume of sample solution can be analyzed directly by ICP-MS using a low ﬂow nebulizer. It should be noted that some elements which are more electronegative than Si have a tendency to redeposit on the Si wafer surface when the appropriate concentration of H2O2 is not present in the HF solution [50]. In addition, thicker thermal oxides can produce very high concentrations of Si in the ﬁnal sample solutions. The concentration can be as high as 1000 ppm, which will certainly cause Si related interferences and matrix suppression problems when using cool plasma ICP-MS for analysis. The Si matrix can, however, be eliminated quite easily by heating, because Si in the sample solution exists in the hydrated form of SiF4, which can be readily evaporated. However, potential loss of B, Se, As and Hg should be considered.

984

Sample preparation for semiconductor materials

29.3.1.10 Poly-Si and quartz A mixture of concentrated HF and HNO3 (1 þ 1) is used for digestion of Poly-Si and quartz. Although the decomposition occurs at room temperature, gasses such as SiF4, escape from the solution, and an appropriate ventilation system is required. When a higher concentration of Si sample is analyzed by ICP-MS, deposition of SiO2 occurs at the tip of sampler and skimmer cones since the plasma temperature is important for decomposition of SiO2. In addition, Si related polyatomic ions, such as SiO, SiF, SiO2 and SiO2H, interfere with the quantitation of Ti, Cu and Zn, respectively. In the past, HR-ICP-MS was the only technique used to overcome these interferences. Recently, the reduction of these interferences by DRC-ICP-MS with hot plasma has been reported, which has also shown better matrix tolerance. The concentration of Si in the matrix prior to analysis by ICP-MS should be lower than 0.2% [51]. Elimination of the Si matrix is recommended when using cool plasma ICP-MS.

29.3.1.11 Colloidal silica A simple dilution with a 5– 10% HF solution is recommended for analysis of a colloidal silica sample. After dilution, the solution is indistinguishable from the ﬁnal solutions prepared by digesting poly-Si and quartz.

29.3.1.12 TiCl4, SiH2Cl2, SiHCl3, TEOS, TDMAT, organometallic compounds These reagents can be analyzed by ICP-MS after preconcentration by evaporation. However, because these reagents are used as source material for chemical vapor decomposition (CVD), impurities in the vapor phase are of greater concern than those in the residue. For this reason, a simple dilution technique is preferable. Some compounds, such as TiCl4, can be diluted with DIW but some others may react with water. The following are general guidelines for analysis of these chemicals: – if a reagent is water soluble, dilute with DIW and then analyze it; – if a reagent is not water soluble, dilute with an organic solvent and analyze it; – if a reagent contains a refractory element, such as Al, Si, Ti, Mo or W, dilute with acids so that the maximum concentration of these refractory elements in the ﬁnal solution is less than 0.1%; these refractory elements form oxides easily and their oxides can deposit on the tip of a cone if their concentration in the ﬁnal solution being introduced into the plasma is high. Last but not the least, some organometallic compounds react with water spontaneously. Material safety data sheet (MSDS) must be checked prior to their digestion.

985

K. Kawabata et al.

29.4

CONCLUSION

ICP-MS is an indispensable technique for quality control of chemicals and materials used in the semiconductor industry. The contamination control is the most critical issue in order to determine ultratrace levels of impurities, and this chapter described several key points to avoid contamination arising from the environment and equipment during the sample preparation. Since the sample preparation is highly dependent upon the analytical technique, it is important to understand capabilities and limitations of analytical techniques prior to the sample preparation.

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22

986

Semiconductor Industry Association, The International Technology Roadmap for Semiconductors, Yield Enhancement, 17, 2001. Semiconductor Industry Association, The International Technology Roadmap for Semiconductors, Front End Process, 16, 2001. D. Burkman, C. Peterson, L. Zazzera and R. Kopp, Microcontamination, November (1988) 57– 62. C. Neumann and P. Eichinger, Spectrochim. Acta B, 46(10) (1991) 1369–1377. J. Fucsko, S.S. Tan and M.K. Balazs, J. Electrochem. Soc., 140 (1993) 1105 –1109. K. Sakata and K. Kawabata, Spectrochim. Acta B, 49(10) (1994) 1027–1038. N. Bradshaw, E. Hall and N. Sanderson, J. Anal. At. Spectrom., 4 (1989) 801– 803. M. Radle, H. Lian, B. Nicoley and A. Howard, Semiconductor Int., 8 (2001) 217– 224. Application Note, Analysis of Ethyl Lactate solvent for the semiconductor industry, Thermo Elemental, Cheshire, UK, April, 2000. V. Baranov and S. Tanner, J. Anal. At. Spectrom., 14 (1999) 1133–1142. Airborne particulate cleanliness classes in cleanrooms and clean zones, US Federal standard 209E, 1992, p. 5. M. Inoue, T. Noda, M. Togawa, H. Inaba, H. Gomi, T. Yoshida and T. Okada, Proceedings of the 12th ISCC in Yokohama, (1994) 111 –116. J. Muller, L.A. Posta-Kelty, H.W. Krautter and J.D. Sinclair, Solid State Technol., 37(9) (1994) 61– 72. Technical News, Evaluation methods for trace impurities in clean room air, (KH0001) 2-A0-(1), Sumika Chemical Analysis Service. E.J. Mori, J.D. Dowdy and L.E. Shive, Microcontamination, November (1992) 35– 43. A. Lieverman, Contamination Control and Cleanrooms. Van Nostrand, New York, 1992. F. Lee, E.M. Howard and D. DeMuynck, Microcontamination, March (1994) 33. A. Hartzell, J. Rose, D. Liu, P. McPherson and M. Oshaughnessy, Microcontamination, October (1996) 69. P.J. Paulsen, E.S. Beary and D.S. Bushee, Anal. Chem., 61(8) (1989) 827– 830. J.B. Goodman and P.M. Van Sickle, Microcontamination, 9(11) (1991) 21– 25. J. Dahmen, K. Englert and G. Giebenhain, International Laboratory—paciﬁc rim edition, April/May, 1997. Y.W. Heo and H.B. Lim, Bull. Korean Chem. Soc., 20(2) (1999) 226– 228.

Sample preparation for semiconductor materials 23

24 25 26 27 28 29 30 31 32 33 34 35 36 37 38

39 40 41 42 43 44 45 46 47 48 49 50 51

R. Hegde, V. Anantharaman and J. Zahka, Measurement of Ionic Extractables from Millipore’s All-Teﬂon Wafergard PF Filter Products, Millipore Application Notes MA018, 1992. M. Cole, R. Van Ausdal and J. Waldman, Microcontamination, 7 (1989) 37. T. Gutowski and A. Shucosky, Semiconductor Pure Water and Chemical Conference, (2002) 100–109. R.W. Welker, Micro, 61(September) (1999). A. Ono, Tsukuba Seminar, Society of Plasma Spectrometry, July 2002, pp. 25– 34. Properties of high purity quartz, Heraeus Amersil Inc., GA, USA. TAMAPURE-AA-10, Tama Chemicals Co. Ltd, Tokyo, Japan. ULTRAPURE, Kanto Chemical Co. Inc., Tokyo, Japan. SPEX CertiPrep Inc., Metuchen, NJ, USA. Inorganic Ventures, Lakewood, NJ, USA. Evapoclean, ANALAB SA, Hoerdt, France. Anton Paar GmbH, Graz, Austria. Milestone S.R.L., Sorisole, Italy. CEM Corporation, Mathews, NC, USA. T. Ishikawa and E. Nakamura, Anal. Chem., 62 (1990) 2612–2616. USEPA method 200.8, Methods for the Determination of Metals in Environmental Samples Supplement 1 (EPA/600/R-94/111), United States Environmental Protection Agency, 1994. R. Hutton, M. Bridenne, E. Coffre, Y. Marot and F. Simondet, J. Anal. At. Spectrom., 5 (1990) 463– 466. S. Laly, K. Nakagawa, T. Kimijima, S. Taguchi, T. Ikeda and S. Hasaka, Anal. Chem., 68 (1996) 4312– 4315. T. Fujimoto, N. Takeda, T. Taira and M. Sado, Semiconductor Pure Water and Chemical Conference, (1996) 325–331. K. Kawabata and Y. Kishi, Semiconductor Pure Water and Chemical Conference, (2001) 241–253. H.M. Kingston and S.J. Haswell, Am. Chem. Soc., Washington, DC, 1997, pp. 55– 222. Y. Kishi and K. Kawabata, Presented at Federation of Analytical Chemistry and Spectroscopy Societies, 2001. K. Kawabata, Y. Kishi, Presented at Winter Conference on Plasma Spectrochemistry, 2002. K. Kawabata, Presented at Tsukuba Seminar of Society of Plasma Spectrometry, February 2002. Conoco Inc., Ponca City, OK, USA. A. Shimazaki, Electrochem. Soc., 91-9 (1991) 47. L. Lovejoy and S. Hues, Future Fab, (2002) 13. F. Meyer and J. White, Semiconductor Int., 137(July) (1999). A. Porche, Y. Kishi and R. Wolf, Analysis of Si Wafers Using the ELAN DRC-ICPMS, Application note D-6444, PerkinElmer Instruments, 2001.

987

This page intentionally left blank

SECTION - 3

This page intentionally left blank

Chapter 30

Sampling and sample treatment in the analysis of organotin compounds in environmental samples Roberto Morabito

30.1

INTRODUCTION

Organotin compounds are molecules characterized by a central Sn atom covalently bound to one or more organic substituents. Their general formula is Rn SnXð4-nÞ where R is an alkylic or arylic group (such as methyl, butyl, phenyl, octyl) and X is an inorganic anion (such as halides, oxide, hydroxide). The Sn –C bond is quite stable in aqueous solutions, in the presence of oxygen and heat. It can be broken by high temperatures (higher than 2008C), UV and g-ray, as well as in the presence of microbiological activities or strong chemical oxidants. The toxicity of these compounds increases with the increasing of the number of organic substituents, reaching a maximum for n ¼ 3; at the same time, their solubility in water decreases and the solubility in lipids increases. Di- and mono-substituted compounds are mainly used as stabilizers for rigid PVC whereas triorganotin compounds are mainly used as biocides in pesticide formulations [mainly triphenyltin (TPhT)] and, above all, in antifouling paints [mainly tributyltin (TBT) but also TPhT]. In Table 30.1 are reported some of the physical –chemical properties of butyl- and phenyl-tin compounds taken out by Blunden et al. (1984), Blunden and Evans (1990) and Hoch (2001) [1 –3]. Antifouling paints are applied on the hulls of boats and in general, on surfaces in prolonged contact with water to inhibit the growth of foulant organisms, such as barnacles, tube worms, etc., and to reduce their deleterious effects such as the slowing of cruise speed, increasing of fuel consumption, increasing of maintenance costs, etc. Starting from the 70s, TBT-based paints replaced copper-based paints due to a superior performance in terms of efﬁcacy and duration (about 5–7 years while copper paints are effective for no more than two years) [4– 7]. Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

991

R. Morabito TABLE 30.1 Physical – chemical properties of butyl- and phenyl-tin compounds [1–3] Compound

Formula

Melting point (8C)

Boiling point (8C)

Tetrabutyltin Tributyltin chloride Dibutyltin dichloride Monobutyltin trichloride Tetraphenyltin Triphenyltin chloride Diphenyltin chloride Monophenyltin chloride

Sn(C4H9)4 (C4H9)3SnCl (C4H9)2SnCl2 (C4H9)SnCl3 Sn(C6H5)4 (C6H5)3SnCl (C6H5)2SnCl2 (C6H5)2SnCl2

297 263 39– 41

145 (1.3 kPa) 172 (3.3 kPa) 135 (1.3 kPa) 93 (1.3 kPa) .420

226 106 42

333–337

Solubilitya (ppm)

5– 50 4– 92

1– 5 50

a

In distilled and sea water, from different references.

TBT is directly released into aquatic environment and its release can be both continuous (release from the hulls of boats) or intermittent (release from dockyard activities as paint removal, cleaning, painting, etc.). Environmental persistence and fate of TBT are strictly correlated to the speciﬁc characteristics of the aquatic ecosystem such as temperature, salinity, pH, suspended matter, microbial populations, ﬂushing rates, etc. Distribution of TBT among the different environmental compartments is regulated by (i) physical mechanisms (including volatilisation, adsorption, etc.), (ii) chemical mechanisms (including photochemical reactions) and (iii) biological mechanisms (including uptake and transformation) [7 –10]. In general, TBT, after its release into aquatic environment, shows a great tendency to be accumulated onto particulate matters, sediments being the ﬁnal sink. At the same time, both TBT and TPhT can be degraded, through stepwise dealkylation mechanisms, down to inorganic tin by biological, chemical and physical pathways [11–14]. The inorganic compounds of tin are practically non-toxic for all living systems, due to their low solubility in lipids [15,16] and extremely low reactivity at physiological pH [17]. On the contrary, the organic compounds of tin show high toxicity that reaches a maximum for the tri-substituted compounds [18,19]. The inorganic substituents do not signiﬁcantly affect the toxicity of the compounds, unless they are strongly coordinating groups [20]. Toxicological data on organotin compounds have been published [20 –27]. The bioaccumulation of TBT in marine organisms has been largely documented [23]. In particular, bivalves can accumulate dissolved TBT from sea water, presumably directly into exposed tissues such as gills, followed by

992

Sampling and sample treatment in the analysis of organotin compounds

migration to other tissues, or by ingesting tainted food. Very high concentrations can be reached in these organisms, because they are not capable, due to a low activity of the mixed function oxidase system, to metabolise a wide range of xenobiotics, including organotins [28,29]. Bioconcentration factors calculated in the ﬁeld [14,30,31] resulted to be higher than those predicted on the basis of octanol –water partition coefﬁcient or calculated from laboratory experiments for oysters [32] and for mussels [33,34]. The high toxicity of TBT together with its tendency to be accumulated in marine organisms can produce heavy damage particularly to mollusc and gastropod populations. The ﬁrst evidence of environmental damage by TBT released by antifouling paints appeared in Arcachon (France) where, from 1975 to 1982, oyster production was severely disturbed by a lack of reproduction and the appearance of shell calciﬁcation anomalies in adult oysters with high economic losses [35–37]. Decline of gastropod population has been registered worldwide as a consequence of the induction of the imposex effect by TBT, consisting of a superimposition of male sexual characteristics on female organisms [38 –41]. High occurrence of imposex has been evidenced in the North Sea, Atlantic Ocean and Mediterranean Sea in Europe as well as along the coasts of USA, Japan, India, Australia, Chile, etc. [42–58]. France was the ﬁrst country to restrict the use of TBT-based antifouling paint by a legislation that inhibited the use of these paints on boats with hulls less than 25 m long [59]. Similar regulations were enacted by countries such as UK, USA, Canada, Italy and many others [60– 63]. Recently, a review of organotin regulatory strategies has been published by Champ [64]. TBT monitoring is today required by many EC legislations, e.g. under the Council Decisions 75/437/EEC (Marine Pollution from Land-based Sources), 77/585/EEC (Mediterranean Sea) and 77/586/EEC (Rhine River), the Council Directive 80/68/EEC (Groundwater) and the Water Framework Directive 2000/60/EC [65]. As a consequence of the relevance of the “TBT case”, many analytical methods for the determination of organotins in environmental matrices have been developed and a huge amount of environmental data have been produced worldwide. Organotin concentration levels in various environmental samples have been determined along the coasts of the Mediterranean Sea [31,55,66 –73], Baltic and North Sea [49,74– 77], Atlantic Ocean [78–82], USA [83–85], Canada [86,87], Oceania [88,89], Asia [89–94] and even Greenland [95]. However, the analytical difﬁculties related to speciation analysis together with the low organotin environmental concentrations and the complexity of the environmental matrices make necessary the step-by-step validation of the applied methods and the evaluation of the quality of the produced analytical data. A number of books or book chapters and reviews dedicated to organotins have been published in the last years [3,23 –24,27,64,65,96 –104].

993

R. Morabito

30.2

CRITICAL STEPS IN ORGANOTIN ANALYSIS

Analytical methods developed for speciation analysis, as for environmental and chemical analysis in general, include a number of single operations; each of them must be considered as a potential source of errors. Schematically, these single operations can be grouped in different steps from sampling to reporting. The total uncertainty budget associated to the ﬁnal results can be considered as a sum of different contributions coming from the single operations or, more schematically, from the different steps in which an analytical method can be schematically divided: 2 2 2 2 UTotal ¼ ðUSampling þ UStorage þ UTreatment þ UAnalysis þ …Þ1=2

The largest contributions arise usually from at least four critical steps including sampling, storage, treatment and instrumental analysis. As state of the art, it is largely agreed that the contribution coming from the ﬁnal analysis is generally in the range 1–20% whereas the treatment step can contribute up to 60–70% to the total uncertainty and the sampling uncertainty can represent 99.999…% of the total budget. Recently, many efforts have been put in the evaluation and the improvement of the quality of analytical measurements. As consequence of the high performances of the new instrumental apparatus and the care for quality put in the analytical operations, the ﬁnal analysis (and its uncertainty) can be considered under control or can be put under control by following adequate precautions. The sample treatment, however, cannot be considered under control in many cases even if tools like reference materials (RMs) and certiﬁed reference materials (CRMs), quality control charts implementation, availability of derivatised calibrants have signiﬁcantly improved the knowledge and reduced the problems associated with this step. Sampling remains, unfortunately, far from being under control. Despite this ﬁgure, many analytical chemists still devote all their efforts to instrumental analysis and to a less extent to sample treatment without or with low care for sampling and/or for the origin of the samples to be analysed in their laboratories. Problems related to the critical steps of organotin speciation analysis have been reported in literature [104–106]. 30.2.1 Sampling There is a large agreement in the scientiﬁc community in considering sampling as the most critical step in environmental analysis. Problems related to sampling include contamination, losses of analytes and change in speciation during the collection of samples. However, the most critical issue regards the representativeness of sampling.

994

Sampling and sample treatment in the analysis of organotin compounds

In order to reduce and minimize these problems, a sampling strategy must be developed for each particular case. This strategy should depend on the ﬁnal aim of the study going on and, starting from a deep knowledge of the chemical– physical characteristics of the investigated analytes, the physical and geochemical characteristics of the sampling site, the anthropic sources, etc., should be able to answer at least ﬁve questions: which and how many samples should be collected and how, when and where the samples should be collected. 30.2.1.1 Which samples should be collected As already mentioned, TBT, after its direct release into the water, is accumulated by marine organisms and onto suspended matter, sediments being the ﬁnal sink. Thus, for understanding the fate and the distribution of TBT in the investigated ecosystem, it is necessary to collect water, sediment and biological samples at each sampling station. Only the knowledge of the organotin concentration levels in all these compartments allow considerations about the “contamination history” of the site and the potential risk for the ecosystem. The knowledge of only the water concentration does not allow a full evaluation of the risk as it was shown in the past: low TBT water concentrations, ranging from few units to few tenths of ppt, were able to produce heavy contamination (ppm levels) in mussels sampled at two different sites, La Spezia and Taranto Gulfs (Italy) [14,31,66,68]; in another Italian site (Olbia Gulf–Sardinia island), similar levels of TBT water concentrations (around 10 ppt) resulted in a signiﬁcantly lower contamination of mussels (tenths of ppb) [107]. Mussel samples collected at the same site of Olbia Gulf few months later showed the same high contamination levels of Taranto and la Spezia Gulfs (ppm levels) despite substantially similar water concentrations (from 10 to 30 ppb) [107]. The data pointed out that there is no direct correlation between water and marine organisms’ TBT concentrations (a better correlation can be found between sediment and biota TBT concentrations) and, thus, there is a need for analysing all the different matrices and no extrapolation can be done. Furthermore, with regard to water samples, many authors perform the analysis after ﬁltration at 0.45 mm, to determine the organotin concentration in the dissolved phase. As the toxicity values of organotin compounds are generally referred to their concentration in dissolved phase, such a procedure is widely accepted to provide information about the contamination levels. However, the strong tendency of TBT to be accumulated onto suspended matter can lead to an underestimate of the true contamination of the site [99]. Depending on the particular site, the concentration of adsorbed TBT can be equal and in some cases higher than in the dissolved phase. Moreover, the contribution of the adsorbed TBT in the bioaccumulation and its toxic effects towards marine organisms cannot be neglected being the particulate matter the primary source of entrance of TBT in feeding ﬁlter organisms. On the other side, particulate matters could contain chip paints that could lead to an unreal overestimation of the TBT contamination level of

995

R. Morabito

the site [108], the storage of non-ﬁltered samples can lead to TBT degradation as mentioned below in the text, the analyses of non-ﬁltered samples provide results not easily comparable due to the variety of the extraction techniques used for organotin analysis. So, despite the lack of agreement between the monitoring authorities of the different countries, the determination of both the concentration of dissolved TBT, after ﬁltration of samples, and of adsorbed TBT, by analysing the ﬁlters, should be recommended. With regard to sediment samples, only the undisturbed surface layer (0 –2 cm) should be sampled. Box corer, or some other low disturbance sampler, should be used for sediment sampling. The sieving of the sediment samples at 60 mm mesh is a widely accepted procedure in order to compare results obtained on sediments collected in other sites [99]. A full characterization of sediments should be done in order to improve the comparability of data collected in different sites. Biota samples should be classiﬁed according to species, size, sex, state of health, position in the water column, etc. 30.2.1.2 How many samples should be collected This issue affects directly the representativeness of both the sampling station and sampling area. In general, the higher the number of collected samples the higher the representativeness. However, the higher the number of collected samples the longer the time of analysis and the time necessary to achieve data and information, and the higher the costs; so, compromises must be done. With regard the representativeness of sampling area, there are different statistical approaches (e.g. use of transects, square grid, ray grid, etc.), depending on the characteristics of the sampling area and its hydrodynamic, that can be used in order to minimize the number of samples to be collected maintaining a sampling area representativeness. With regard to the representativeness of the sampling station, there are no systematic approaches. Conclusions of an Interagency workshop on aquatic sampling and analysis of organotins, held in 1986 in USA, recommended the collection of 3 samples for each station and the analysis of two of them; if the results are different by more than 15%, the third sample should be analysed and all the results of the three samples should be used to calculate the mean [109]. 30.2.1.3 How the samples should be collected Contamination from the hulls of the sampling boats treated with organotinbased antifouling paints could lead to overestimation of organotin concentrations in water. For this reason the use of rubber boats for sampling is strongly recommended. Monobutyltin (MBT) and dibutyltin (DBT), primary degradation products of TBT, are largely used as heath stabilizers of rigid PVC. Release of these compounds from PVC was evidenced [110]. Thus, PVC bottles should not be used as sampling bottles in order to avoid possible changes in the organotin

996

Sampling and sample treatment in the analysis of organotin compounds

speciation due to the release of DBT and/or MBT from the walls of the containers. Losses of organotins due to the adsorption on the container walls are minimized by using polycarbonate bottles [111]. Teﬂon sheets, glass plates and stainless steel mesh frames have been used for organotin sampling but risks of adsorptive losses have not been studied [112]. Rotating polycarbonate drum sampler has been used to minimize losses [112]. Surface microlayer and bottom water should not be collected as reported below; the use of close –open –close samplers with a remote-controlled stopper would allow collection of water at controlled depth. 30.2.1.4 Where the samples should be collected Signiﬁcant differences of TBT and MBT concentrations usually occur in the water column. TBT shows a great tendency to accumulate in the surface microlayer reaching concentrations much higher (1 or 2 order of magnitude) than in water column [112,113]. Thus, superﬁcial water samples should be collected at about 50 cm under the water surface. Organotin concentrations usually decrease as the water depth increases [14,68,113]. However, higher concentrations of organotins could be found at the interface water–sediment. The primary degradation product of TBT in sediments seems to be MBT [14, 114] that, because of its hydrophilicity, may enter the water column in a short time after its formation. Moreover, sediment resuspension phenomena can lead to resolubilization of adsorbed organotins. Recent results by Amouroux et al. [115] would strengthen the above point. Diffusion of organotin compounds through volatilisation processes should lead to signiﬁcant enrichments just above the sediment–water interface. Hence, depth water samples should be collected at a depth sufﬁciently above the bottom to avoid direct inﬂuence from resuspension or degassing out of the sediment. Signiﬁcant differences of TBT concentrations were registered between enclosed and open sampling areas both for water and sediments [66,116]. Sediments collected in the range of few tenth of meters can show signiﬁcant differences in organotin concentrations due to chip paints arising from dockyard activities [117]. 30.2.1.5 When the samples should be collected Seasonal variation in the organotin concentration often occurs due to the increase of the anthropic sources during summer (due to leisure boats) and different extension of the degradation phenomena between hot and cold season. Higher concentrations of organotins are usually determined during spring and summer time [14,113]. Strong seasonal ﬂuctuations in TBT concentrations have been evidenced in the south and southwest of England [118]. Sampling carried out during low or high tide can lead to an over- or underestimation of the organotin concentration in the dissolved phase due to the dilution effect of tide. High differences in organotin concentrations in water

997

R. Morabito

samples collected before and after a tidal cycle have been reported [119]. Slack tide is the time recommended by US EPA. General theory and recommended approaches to sampling have been recently published in English by Pierre Gy [120]. 30.2.2 Storage Storage of samples is one of the critical aspects of the whole analytical procedure, during which physico-chemical alterations occurring in the samples could affect the ﬁnal determination. Adsorption (losses) and desorption (contamination) on and from the container walls, losses of analytes for volatilisation, degradation due to biological activity and/or UV irradiation can occur during the storage of samples affecting directly their representativity. This is particularly true for organotins in biological and sediment samples where degradation phenomena are well documented. The main parameters controlling the stability of chemical species during storage are pH, temperature, light conditions and the material of containers in case of water samples whereas humidity and temperature are the main parameters for solid samples being the light conditions and the material of containers much less critical in this case. Regarding the stability of organotin compounds in stored environmental samples, relatively few studies have been carried out so far, and most of them have been reviewed [104 –106,121,122]. Organotin stability was evaluated on seawater samples stored at different temperatures (þ48C and room temperature), under different light (daylight and dark) and pH (acidiﬁed or not) conditions, in different materials (glass, PTFE, PVC, polycarbonate), under different chemical –physical conditions (dissolved in the sample and adsorbed on C18 cartridges) [13,111,123–127]. Stability of organotins in stored natural freshwater samples was investigated too [128]. TBT resulted to be stable over 4 months in samples of seawater ﬁltered and acidiﬁed at pH 2 and stored in the dark both at þ4 and þ208C; DBT and MBT resulted to be stable only at þ48C [13]. TBT concentration in water did not signiﬁcantly change in samples stored in pyrex glass at þ48C for 4–5 months [13,111,127]. These results have been conﬁrmed in a successive study where all butyltins showed good stability over 7 months in both acidiﬁed and unacidiﬁed samples stored at þ 48C whereas, in the same conditions, phenyltins displayed large instability starting from the ﬁrst month of storage [126]. Adsorption of organotins on PTFE and glass container walls was evidenced [123,124] whereas the use of polycarbonate bottles instead of pyrex bottles led to an increase of butyltin stability as well as the use of C18 cartridges even at room temperature [126]. PVC containers must be avoided for sample storage. Leaching of DBT and MBT from PVC containers has been experimentally evidenced [110,129].

998

Sampling and sample treatment in the analysis of organotin compounds

Stability of organotins in sediments have been tested under different temperatures of storage (from 2 20 to þ 40 8C) as they are (wet) and after different drying procedure (air and oven drying, freeze drying, pasteurisation) [13,126,127,130–132]. Studies on the stability of sediment extracts showed that no signiﬁcant variation in the TBT and DBT concentrations was observed in samples stored at þ4 and þ208C; on the contrary, a signiﬁcant degradation of MBT was observed [131,132]. Other studies showed degradation of all butyltin compounds in sediment stored at temperatures higher than þ48C [126,130]. Butyltins in sediments stored at both þ4 and 2 208C resulted to be stable over a period of 4 months [13] and 1 year [130], respectively. However, sediment storage at þ48C for periods longer than 1 year can lead to instability problems as testiﬁed by the need of recertiﬁcation of BCR 462 [133]. This CRM (coastal sediment) was originally stored at þ48C but after two years there was a clear evidence of degradation and recertiﬁcation was needed. Pasteurisation and air- and oven drying led to degradation of organotins also in samples stored at þ48C. No signiﬁcant degradation was observed in freeze-dried sediment samples stored at þ4 and 2208C [126]. The stability of organotin compounds in freeze-dried mussel samples stored under different conditions of light and temperature was investigated over 4 years. The samples were stored at four different temperatures in the dark: 220, þ 48C, room temperature and þ408C. Furthermore, at room temperature, samples were stored in the daylight as well. TBT resulted to be stable over 6 months in samples stored at þ48C and over 4 years at 2208C. On the contrary, samples stored at þ408C and at room temperature displayed signiﬁcant degradation particularly if stored in the light. More pronounced degradation phenomena were observed for TPhT. Signiﬁcant decrease of TPhT concentration was observed also in samples stored at þ48C, while in samples stored at room temperature in the light TPhT was completely degraded. Complete details of this study are reported elsewhere [134,135]. Other studies on the stability of organotins in biological matrices led to similar conclusions [126]. Recently, stability of butyltins in a oyster candidate reference material has been tested over 18 months in samples stored at temperatures from 220 to þ408C. The compounds resulted to be stable at þ 4 and 2 208C. On the contrary, 50% TBT degradation was observed in samples stored 1 month at þ408C and 12 months at room temperature [136]. As a conclusion, stability of organotin compounds in stored environmental samples has been demonstrated to be crucial and further studies are needed to optimise all the storage parameters reducing the risk of degradation. However, some indication and recommendation can be provided on the basis of the work done.

999

R. Morabito

In case of water samples, samples should be ﬁltered and then stored in polycarbonate bottles at þ48C in the dark; pH should not be lower than 2 and the samples should not be stored for more than one month. Sediment and biota samples should be stored wet or (better) freeze-dried at 2208C. In these conditions, butyltins are stable for years while phenyltins are stable for several months. A long-term stability both for butyl- and phenyl-tin compounds can be achieved by storing freeze-dried samples at 2 708C. 30.2.3 Sample treatment Most of the analytical methodologies for organotin determination envisages an extraction of these compounds from the matrix, their derivatisation in case of gas chromatographic separation and a puriﬁcation step. Sample treatment can be certainly considered one of the most critical steps, being prone to errors. Contamination of samples can arise from glassware, reagents, the laboratory and the operator itself. In case of organotin analysis, plastic glassware, and in particular PVC (containing di- and mono-butyltin) and plastic gloves (sometimes containing trioctyltin), must be avoided. However, the crucial point is in the choice of the most suitable extraction and derivatisation reagents. It is trivial to remind, in fact, that low extraction efﬁciencies and/or low derivatisation yields lead directly to an underestimation of the real contamination. Furthermore, changes in the real speciation during treatment due to degradation must be taken into account, optimising not only the choice of the most suitable reagents but also the experimental conditions like temperature, light and treatment time. 30.2.3.1 Extraction Extraction is primarily used: (i) to separate the analyte from the matrix eliminating or reducing the interferences from the other components; (ii) to concentrate the analyte up to detectable concentration level. In speciation analysis, extraction should be performed not only in such a way that the analyte is separated from the interfering matrix without loss or contamination, but also without changes in speciation [137]. Low extraction yields, losses of analytes and contamination directly affect the quality of the results. Moreover, changes in the organotin speciation occurring during this step lead not only to wrong information about the contamination levels but also to wrong considerations about the extension of the ongoing degradation phenomena. Thus, recovery should be carefully tested for each organotin compound. The best approach should be the use of CRMs even if spiking experiments are mostly applied. As reported below, however, spiking experiments can lead to an overestimation of the real extraction efﬁciency, and the best strategy and

1000

Sampling and sample treatment in the analysis of organotin compounds

experimental conditions under which spiking experiments should be carried out are still strongly debated within the scientiﬁc community. Anyway, some approach may be recommended [105]. It is worth to stress, on the other hand, that the use of CRMs leads to an evaluation of the performance of the overall analytical method rather than an evaluation of the extraction efﬁciency only, as the differences between the certiﬁed and the found values are usually imputable not only to low extraction efﬁciency but also to, e.g. low yields of derivatisation, losses during the clean-up step, etc. Full details on the evaluation of the extraction efﬁciency in organometallic speciation are given elsewhere [138]. A large number of extraction methods for organotin determination in environmental samples have been reviewed [101,139– 144]. Recently, a comparison of 12 selected extraction methods for butyltin and phenyltin species determination has been carried out [142]. Their extraction efﬁciency was compared by analysing a certiﬁed reference material for organotins in mussel tissue (BCR 477). The use of a CRM guarantees that differences in the results cannot be imputable to inhomogeneity between the analysed samples. Furthermore, the comparison with the certiﬁed values allows the evaluation of the performance of the method. Following the application of the different extraction methods, the same, previously optimised derivatisation and clean-up steps have been applied before GC –MS analysis [145]. Sample treatment and analyses were carried out in the same laboratory and by the same operator. In these conditions, the performances of the overall methods can be reasonably considered strictly related to the efﬁciencies of the extraction method applied that can be then compared and evaluated. The selected methods envisaged the use of solvents of different polarity such as methanol [98,142,146 –148], dichloromethane [86,147,149,150], pentane [80,149], hexane [149–152], toluene [153] in the presence [80,86,98,101, 105,136–147,148–150,151–153] or not [142,148,150] of a complexing agent like tropolone and with the addition of acids such as HCl [98,148,151,152,154, 155], HBr [80,147,149] and acetic acid [148] or without [86,142,146,147,149, 150]. Results showed that acidic conditions together with the use of tropolone and a polar organic solvent enhance the extraction efﬁciency for these compounds. The presence of acidic conditions seems to be necessary for all the considered organotin compounds. The only exception is represented by the simultaneous use of methanol and tropolone that in case of butyltins provided excellent results despite the absence of acidic conditions. Acid concentrations higher than 1 mol l21 provided lower results probably due to some degradation of the tri- and mono-substituted compounds in particular. On the contrary, the nature of the acid does not seem to strongly inﬂuence the extraction efﬁciency even if hydrochloric and hydrobromic acids provided best results than acetic acid. The presence of tropolone has a strong inﬂuence on the extraction of di- and monosubstituted compounds while, as expected, it does not inﬂuence the extraction

1001

R. Morabito

of the tri-substituted compounds. Finally, the higher the polarity of the organic solvent the higher the extraction efﬁciency probably due to a better “wettability” of the matrix. Alkaline extractions with NaOH and KOH in ethanol have been proposed by Waldock and Waite [156] and Nagase and Hasebe [157], respectively. Extraction after solubilization of the matrix with tetramethylammoniumhydroxide (TMAH) [152,158,159] and after enzymatic digestion (generally using lipase and protease) [152,160,161] have also been proposed for organotin. Recently, the use of accelerated solvent extraction systems (ASE) has been proposed also for organotin analysis [162,163]. The extraction methods described above are generally applied to both sediment and biological matrices even if, depending on the different methods, the recoveries can vary from sediment to biota. The only exception are represented by the methods based on the use of solubilizers and enzymes whose application is restricted to biological samples. Recovery evaluation. The evaluation of the extraction recovery of the considered method can be done by using CRMs or surrogates such as in the isotope dilution approach, spiking experiment, internal standard [138]. In the absence of reference materials, the most commonly applied approach is to assess the recovery by a spiking procedure. The procedure consists of the addition of a known amount of the analyte (spike) to the matrix. The recovery of the spike can be evaluated by analysing the matrix before and after the spike addition. This approach suffers from some drawback, i.e., the spiked compounds may not be in effective equilibrium with the incurred ones; a weaker binding of the spike onto the matrix in comparison to the native compound means that the recovery will likely be overestimated, which will lead to a low bias in the corrected result. The major drawback is that the spike is not always bound the same way as the naturally occurring compounds; generally, as the spiking compounds are bound to the matrix in a weaker way than the native compounds, spiking experiments lead to an overestimation of the real efﬁciency of the tested method. Results of spiking experiments should be used in a conservative way: if quantitative recoveries are obtained on spiked samples, this does not mean that the same efﬁciency will be achieved on unknown samples, but if quantitative recoveries are not obtained on spiked samples, it is sure that no quantitative extraction will be achieved on unknown samples. This means that spiking experiments should be used to have the proof that you are getting wrong but not the proof that you are getting right. In general, the best experimental conditions under which spiking experiments can be suitably carried out are still debatable. Some recommendations are given [105]: –

Long equilibration time in case of solid samples (e.g. soils, sediments, biota) is usually necessary to simulate as well as possible a “natural” adsorption of the spike. Equilibration time of 24 h, or at least “overnight”,

1002

Sampling and sample treatment in the analysis of organotin compounds

– –

is generally recommended. However, the equilibration time should be decided case-by-case, taking into consideration the nature of the species and/or the matrix to be spiked. Possible degradation or transformation of the spike during the equilibration, particularly in case of sediment samples, could occur. Recovery tests are to be carried out for each single species as the extraction efﬁciency changes from one species to another. Real matrices as similar as possible to the unknown sample should be used. Furthermore, these matrices should be free from the analyte(s) of concern or at least with a content below the detection limit of the determination technique used. Sediment and sea water samples collected in open sea or in remote areas, can meet this last requirement for many species (e.g. organotins); adversely, biological samples and especially feeding ﬁlter organisms usually present signiﬁcant concentrations of the analyte(s) of concern. In this case, the level of the incurred compound(s) must be accurately evaluated before spiking and the percentage of recovery is usually referred to the sum of the original content plus the spike or directly to the spike amount after subtraction of the original content. It is worth to stress, that the two methods lead to a different evaluation of the recovery and that both of them do not lead to a correct recovery evaluation.

Other considerations have been given by Quevauviller and Morabito [138]. One further shortfall of this approach is the lack of harmonised, welldeﬁned, spiking protocol. A recent attempt has been made to use a common procedure for evaluating the recovery of organotin compounds during the certiﬁcation of reference materials. The BCR guidelines for the production of CRMs [164] request that the results provided by the participating laboratory to the certiﬁcation campaign be corrected for recovery. In order to overcome problems due to the differences in the recovery evaluation procedures, it was decided, during the development of the BCR 477 [165] project, to provide all participants with guidelines for spiking experiments. The protocol, suggested by the Coordinator (ENEA, Italy), was discussed in a technical meeting before the start of the certiﬁcation campaign. The participants agreed on the following points: (a) A preliminary analysis should be performed in order to have an idea about the organotin concentration levels in the sample; (b) Underivatised spiking compounds (pure organotin salts) should be used; (c) Standard additions at three different levels should be carried out. The ratio among the different spike additions should be 1 (same concentration level found during the preliminary analysis):2:4. (The concentrations of the spike levels was strongly debated because some participants indicated that 4 as third level was to be considered too high).

1003

R. Morabito

If the experiment is correctly carried out, a straight regression line should be obtained: y ¼ a þ bx where y is the amount found, x is the added spike amount and b ¼ recovery/100 (d) The same amount of sample should be taken for all spiking experiments; (e) 1–2 ml of methanol should be added to each aliquot (100 –500 mg) for sample rewetting. Then, 1 ml of methanol containing the spiking compounds should be added dropwise to the aliquots. It is recommended to use only freshly prepared spiking solutions; (f) All the aliquots should be left overnight under mechanical agitation; they should be analysed on the same day; (g) The spiking solvent, if still present, should be evaporated by a gentle stream of nitrogen. Basically, the same protocol, improved by the suggestions of the laboratories participating in the projects, has been considered also for the certiﬁcation of BCR 646 [166] and candidate BCR 710 [136]. 30.2.3.2 Derivatisation Due to the low volatility of the organotins, a derivatisation reaction is required when a gas chromatographic-based technique is going to be used. Furthermore, derivatisation also allows the reduction of the occurrence of possible interferences during the subsequent analytical steps and particularly at the detection stage. It is worth to stress that derivatisation must be considered, as well as extraction, one of the most critical steps in organotin analysis. Low yields of derivatisation and/or degradation phenomena (especially in case of phenyltins) can heavily affect the quality of the results. A validation or at least a careful study of the procedure being used in the laboratory for the particular matrix with its particular interferences is necessary [145,168]. This validation is, however, almost always hindered by a lack of commercially available derivatised standards. Recently, the synthesis of derivatised standards for ethylation and Grignard derivatisation carried out at the Free University of Amsterdam [135] within the frame of Standards, Measurements and Testing Programme ﬁnanced projects allowed the establishment of optimisation and validation studies. The most common derivatisation reactions applied for organotin analysis are: hydride generation with NaBH4, ethylation with NaBEt4, and alkylation with Grignard reagents [145]. Performances and suitability of these techniques for organotin determination in environmental matrices have been recently critically evaluated [168].

1004

Sampling and sample treatment in the analysis of organotin compounds

Hydride generation and ethylation with NaBEt4 are particularly suitable for aqueous samples, both presenting the main advantage to be directly applicable to the samples. Simultaneous in situ derivatisation/extraction is possible, reducing the number of analytical steps and then the potential sources of errors. Ethylation, contrarily to hydride generation, presents high yields of derivatisation not only for butyltin compounds but also for phenyltin ones. In case of solid samples such as sediment and biological samples, hydride generation is often hindered by the presence of severe interferences whereas the main advantage of NaBEt4 (its stability in water) is not anymore exploitable. Furthermore, NaBEt4 in presence of strong acids, often applied in the extraction of organotin compounds from solid samples, is not stable and decomposes. In this case, the organotins have to be extracted into an organic solvent prior to the derivatisation reaction. Grignard derivatisation offers the advantage of being able to be used for the determination of most of the organotin compounds present in the environment (methylated, butylated and phenylated species) in a large variety of matrices (water, sediments, biota). Grignard derivatisation can be performed by methylation, ethylation, propylation, butylation, pentylation and hexylation. Among the different Grignard reactions, hexylation and pentylation present the advantage of providing derivatised compounds with relatively low volatility, which allows preconcentration steps in the sample pretreatment without taking special precautions, even if, due to the low volatility, condensation problems in the interphase have been described when the coupling GC –AAS is used [140]. The use of propyl, ethyl, and, above all, methyl Grignard reagents offer the advantage of a higher reactivity even if the high volatility of their products can lead to volatilisation losses during the preconcentration steps and careful precautions must be taken. Methylation has an additional disadvantage, it cannot be applied to the determination of methylated species. Same precautions must be taken as well in case of ethylation with NaBEt4. Independently on the applied derivatisation technique, it should be necessary to check the derivatisation yields in the chosen experimental conditions. Detailed information of the three techniques, when applied to organotin speciation, are reported in the previous mentioned paper [168].

30.3

IMPROVING THE QUALITY OF ORGANOTIN MEASUREMENTS IN EUROPE

Following a consultation of European expert and according to the relevance of the problem, the European Union started in 1988 a “Tin speciation” project [169].

1005

R. Morabito

The aim of this project was to evaluate the state of the art of the analytical performances of the European laboratories involved in such analysis, improve these performances and certify reference materials. The project, still going on, allowed the participating laboratories to individuate, along the analytical method, at least three main sources of error (excluding sampling and storage): extraction, derivatisation and detection. The European state of the art of the analytical measurements in the organotin ﬁeld can be evaluated looking at the results obtained in the last 10 years within the EU “Tin speciation” project. This project, like other European projects, followed the classical “BCR step-by-step approach”. The BCR approach consists of a series of intercomparisons of increasing difﬁculties, starting from easy intercomparisons on standard solutions (to check the performance of the analytical techniques), on cleaned extract(s) (to detect sources of error occurring at the separation step), on raw extract(s) (to detect sources of error occurring at the clean-up step), on spiked samples (to detect sources of error occurring at the extraction step) and arriving to very complicate intercomparisons on real matrices (to check the performance of the whole method). This type of improvement scheme enables to detect and remove errors speciﬁcally occurring at an analytical step. With respect to the “Tin speciation” project, the improvement scheme focused on intercomparisons on standard solutions, spiked samples and real matrices. The ﬁrst exercise asked participants to perform analyses on four standard solutions of TBT in presence or not of other tin compounds such as DBT, MBT, TPhT and Sn(IV). Fifteen laboratories from seven EC countries participated in this exercise. The results of this intercomparison were found to be in good agreement, which indicated that possible systematic errors did not arise from the techniques of ﬁnal determination. The second exercise was undertaken in 1989 on the determination of TBT in a spiked sediment (collected in the Lake Maggiore, Italy, and prepared at the EC Joint Research Centre of Ispra). The sediment could be considered a low polluted sediment and preliminary analyses showed no detectable organotin concentration. The sediment was then spiked with 3.3 ppm of TBT. The results of this interlaboratory trial did not reveal any systematic errors in the different analytical methods compared. Fifteen out of the 15 participating laboratories were able to provide results. The coefﬁcient of variation (CV) obtained between laboratories (25%) was considered to reﬂect the state of the art at that stage, and the ratio between the highest and the lowest set of values was around 2. The group of experts recommended to proceed with the organisation of a certiﬁcation campaign. The harbour sediment to be used in the interlaboratory study (RM 424) was collected in the Sado Estuary (Portugal) in the vicinity of the harbour of Setu´bal. This sediment could be considered a high polluted sediment. A very large scatter of data, ranging from less than 10 to more than 300 ng g21 of TBT, was

1006

Sampling and sample treatment in the analysis of organotin compounds

unfortunately obtained with a ratio between the highest and the lowest set of values of more than 30. Six laboratories were not able to report values as the detection limits of the techniques used were too high. Detailed discussions were necessary to explain the sources of discrepancies that were individuated at each stage of the procedure (extraction, derivatisation and separation/detection). The methods using hydride generation and AAS as ﬁnal determination were found to be in considerable difﬁculty with this complicated matrix, due to unknown interferences, either at the derivatisation step or in AAS detection, or detection limits which were too low. As observed by the participants, the laboratories using gas chromatographic separation and detection either by FPD or MS tended to agree, which would conﬁrm that these methods would be more suited to the determination of TBT in this particular material. New results obtained by independent laboratories using GC-MIP-OES, SFE-GCFPD and GC-FPD agreed with the results found during the intercomparison by the chromatographic technique. Although the analytical methods involving hydride generation were successfully applied in the interlaboratory study on TBT in spiked sediment, it was concluded that the low TBT mass fractions and the complicated matrix did not allow the use of these methods for an accurate TBT determination in this material. It was stressed that although certiﬁcation was contemplated at the start of the interlaboratory study, this material would probably not be suited as a CRM, and it is now available on the BCR catalogue as RM 424 with indicative values of TBT, DBT and MBT [170]. Additional information is reported in Table 30.2. The intercomparison highlighted that laboratories (and analytical techniques) able to detect high amount of TBT in easy matrices (3.3 ppm of TBT in the low polluted Lake Maggiore sediment) may fail in the determination of low amount of TBT in difﬁcult matrices (20 ppb in the high polluted Sado Estuary sediment). Further studies on the applicability of hydride generation techniques to the determination of organotins in sediments were envisaged. An attempt to certify butyltins in sediment was tried again two years later. The sediment was collected in the Arcachon bay (France). The material TABLE 30.2 RM 424—harbour sediment Species

Indicative value (mg/kg)

Accepted sets of results

Techniques used

GC– MS, GC–FPD, HPLC– ICP-MS, GC– MIP-OES HG-GC– AAS, GC –MS, GC –FPD GC– MS, GC–FPD

TBT

20 ^ 5

8

DBT MBT

53 ^ 19 257 ^ 54

6 3

1007

R. Morabito

(BCR 462) was successfully certiﬁed for its TBT and DBT content at values of 70 ^ 14 and 128 ^ 16 ng g21, respectively [171]. However, the technical discussion of the certiﬁcation results highlighted that poorer extraction recoveries for TBT in the candidate CRM were often observed in comparison to recoveries obtained from other sediment materials. Some laboratories found lower recoveries if the spike was allowed to equilibrate longer. A good agreement was generally observed for DBT. In the case of MBT, furthermore, problems in the extraction step reported by many laboratories and a high scatter of results prevented certiﬁcation. Despite the success of the TBT and DBT certiﬁcation, the discussion stressed that at least two steps of the analytical method, i.e., extraction and derivatisation, could not be considered under control and dedicated studies were ﬁnanced. CRM 462 was stored at room temperature but, after two years, some doubt regarding its long-term stability arose. The material was withdrawn from the market and recertiﬁed as BCR 462R [133]. Additional information on BCR 462R is reported in Table 30.3. This fact demonstrated the necessity to store CRMs for organotins at 2 208C in order to increase their long-term stability. Following the studies on sediments, a certiﬁcation of butyl- and phenyl-tin species in mussel tissue was attempted. Mussels were collected in the La Spezia Gulf where is located one of the main Italian mussel farm. The results were extremely satisfying for butyltins and TBT, DBT and MBT could be certiﬁed and the material is available in the BCR catalogue as BCR 477 [135,165]. It is worth stressing that in the case of TBT, results from 18 out of 19 participating laboratories were accepted and that the RSD was around 8% only, testifying the high level of performances reached by the laboratories participating in this project. Additional information on BCR 477 is reported in Table 30.4. On the other hand, phenyltins’ certiﬁcation was hindered by a lack of longterm stability of these compounds and the high scatter of results among

TABLE 30.3 BCR 462R—coastal sediment Species

Certiﬁed value (mg/kg))

Accepted sets of results

Techniques used

TBT

54 ^ 15

5

DBT

68 ^ 12

7

HG-GC–AAS, GC– MS, GC–FPD, HPLC–ID-MS HG-GC–AAS, GC– MS, GC–FPD, GC –MIP-OES, GC–ICP-MS, HPLC–ICP-MS

1008

Sampling and sample treatment in the analysis of organotin compounds TABLE 30.4 BCR 477—mussel tissue Species

Certiﬁed value (mg/kg)

Accepted sets of results

Techniques used

TBT

2.20 ^ 0.19

18

DBT

1.54 ^ 0.12

15

MBT

1.50 ^ 0.28

8

GC– QF-AAS, HG-GC–AAS, GC– MS, GC– FPD, GC–MIP-OES, HPLC–ID-ICP-MS, HPLC–ICP-MS, HPLC–ICP-OES, HPLC-Fluo GC– QF-AAS, HG-GC–AAS, GC– MS, GC– FPD, GC– MIP-OES, HPLC–ICP-MS GC– QF-AAS, HG-GC–AAS, GC– MS, GC– FPD, GC– MIP-OES, HPLC–ICP-OES

laboratories (RSD up to 50%) testifying that further efforts are needed to reach comparability of data in phenyltin analysis [172]. Following this campaign, a certiﬁcation campaign for organotins in freshwater sediments has been carried out. As the previous projects had pointed out the lack of derivatised calibrants, the preparation of a series of calibrants of veriﬁed purity and stoichiometry for distribution to the participants in the certiﬁcation campaign was envisaged in the project. Ethylated and pentylated butyltins as well as their chloride salts have been prepared and distributed among the participants. The project was successful and the material has been certiﬁed for butyl- and phenyl-tins and is now available as BCR 646 [173]. Further details are reported in Table 30.5. Finally, the MULSPOT project (preparation of a certiﬁed oyster tissue reference material for species of tin, mercury and arsenic—candidate BCR 710) has been successfully concluded in 2002, proposing TBT and DBT (besides arsenobetaine and methylmercury) for certiﬁcation. Four oyster materials have been prepared for intercomparisons (T37 and T38), recovery evaluation (T36) and certiﬁcation (T34). Twentytwo laboratories from 10 European Countries and one laboratory from Canada participated at the certiﬁcation. The results of the MULSPOT certiﬁcation campaign are schematically reported in Table 30.6 whereas the schematic description of the analytical methods used by the participating laboratories is reported in Table 30.7. By now, the “Tin speciation” project allowed the production of one reference material and four certiﬁed reference materials (one of which is at the certiﬁcation committee stage), the development of new analytical methods for organotin speciation, the preparation of pure derivatised

1009

R. Morabito TABLE 30.5 BCR 646—freshwater sediment Species

Certiﬁed value (mg/kg)

Accepted sets of results

Techniques used

TBT

480 ^ 80

14

DBT

770 ^ 90

13

MBT

610 ^ 120

10

TPhT

29 ^ 11

7

DPhT

36 ^ 8

8

MPhT

69 ^ 18

6

GC– MS, GC– FPD, GC–MIP-OES, GC– AED, GC –QF-AAS, HPLC-Fluo GC– MS, GC– FPD, GC–MIP-OES, GC– AED, GC –QF-AAS GC– MS, GC– FPD, GC–MIP-OES, GC– AED, GC –QF-AAS GC– MS, GC– MIP-OES, GC– AED, HPLC-Fluo GC– MS, GC– FPD, GC–MIP-OES, GC– AED GC– MS, GC– FPD, GC–MIP-OES, GC– AED

calibrants, the optimisation of extraction and derivatisation techniques, leading to a general improvement of the quality of organotin analysis in environmental samples. Further information on the “Tin speciation” project has been published by Quevauviller [174] and Quevauviller and others [175]. In addition to the BCR reference materials, other CRMs are available from the National Research Council of Canada (NRCC) and the Japanese National Institute for Environmental Studies (NIES). The list of the CRMs for organotin analysis available on the market is reported in Table 30.8.

TABLE 30.6 Candidate BCR 710-oyster tissuea Species

Certiﬁed value (mg/kg)

Accepted sets of results

Techniques used

TBT

135 ^ 27

10

DBT

82 ^ 19

9

GC–MS, GC –FPD, GC–ICP-MS, HPLC–ID-ICP-MS, GC–AED GC–MS, GC –FPD, GC –ICP-MS, GC–AED

a

The candidate BCR 710 has been proposed also for the certiﬁcation of arsenobetaine and methylmercury.

1010

TABLE 30.7

Lab Analytical code technique

Species

Sample mass (mg)

Sample pretreatment

Calibration

Extraction: 2.5 ml methanol; mechanical shaking at 420 rpm (2 h), þ 12.5 ml HCl (0.12 M) in methanol, ultrasonication (1 h); neither drying nor clean-up of the extract; no preconcentration. After extraction the sample is directly introduced in a derivatisation reactor; involving a one-step simultaneous ethylation and extraction, using NaBEt4 in the presence of an iso-octane layer Extraction: sample þ 15 ml methanol/tropolone 0.03% þ 1 ml HCl (12 M); soniﬁcation (30 min), repeated once; centrifugation: transfer to methylene chloride (15 ml), separation in separatory funnel, twice; organic phase dried with anhydrous Na2SO4; volume reduction to 1 ml solvent exchange with isooctane; Derivatisation: Grignard alkylation with 1 ml pentylmagnesium bromide (2 M) in ethyl ether (room temperature 5 min). Clean-up: ﬂorisil column. Separation: fused column (25 m £ 0.25 mm £ 0.45 mm) CP SIL 8CB

1000 mg/l (as tin) st. stock solutions in methanol: MBT 95%, DBT 97%, TBT 96% (all Aldrich) TPrT 98% as Int. St. (Strem Chemicals)

02

GC– PFPD TBT, DBT, MBT

500

03

GC– MS

500

TBT, DBT, MBT

Pentylated calibrants. Pure calibrant TPrT (.99%), MBT (.98%), DBT (.99%), TBT (.98%), all IVM. TPrT as internal standard

continued

Sampling and sample treatment in the analysis of organotin compounds

Schematic description of the analytical methods used by the laboratories participating in the certiﬁcation campaign of the MULSPOT Project

1011

1012 TABLE 30.7 (continuation) Species

Sample mass (mg)

Sample pretreatment

Calibration

10

TBT, DBT, MBT, TPhT, DPhT, MPhT

400

Samples are redissolved in 5 ml water þ 0.5 g NaCl þ 0.05 ml 37% HCl. Extraction using 12 ml 0.02% tropolone in diethylether, ultrasoniﬁcation (5 min), manual shaking (5 min); ether phases are combined in a tube with anhydrous Na2SO4; reduction of ether volume; evaporated to dryness in a 408C water bath; Samples are redissolved in 1 ml toluene; cooled on ice, reaction with 0.6 ml 2 M pentylmagnesium bromide in diethylether (5 min); samples are quenched with 1.2 ml 1 M NH4Cl, the toluene phase is removed, evaporation to dryness under N2; redissolution in a known mass of toluene. Separation: 15 m £ 0.53 mm £ 1.5 mm SBP-1 column

MBT (95%), DBT (96%), TBT (96%), MPhT (98%), DPhT (96%) (all Aldrich); TPhT (.97%, Fluka); 1000 ppm stock solutions in both methanol and hexane. Methanol standards used for spiking and standard addition calibration; hexane standards are used for recovery evaluation. TeBT (93%, Aldrich) as internal standard

GC–ICPMS

R. Morabito

Lab Analytical code technique

TABLE 30.7 (continuation) Species

Sample mass (mg)

Sample pretreatment

Calibration

11

TBT, DBT, MBT, TPhT, DPhT, MPhT

1000

Sample þ 50 ml of water/HBr (1:1, v/v), magnetic stirring (60 min); extraction with 50 ml 0.04% (w/v) tropolone in CH2Cl2 (2 h); centrifugation (10,000 rpm, 10 min); organic phase dried with anhydrous Na2SO4; volume reduction to about 1 ml by rotatory evaporation; solid residue þ 5 ml hexane; shaking (5 min); centrifugation; organic phase used to re-extract aqueous phase (3 min); hexane extract dried, added to concentrated dichloromethane extract; volume reduction to 0.5 ml by rotatory evaporation; derivatisation by 4 ml 1 M pentylmagnesium bromide in ether (1 h, room temperature). Clean-up: excess reagent removed with 0.5 M H2SO4 in a cold water bath; centrifugation (10,000 rpm, 5 min); aqueous phase extracted twice with 5 ml pentane, combined with former extract; drying with anhydrous Na2SO4, reduction of volume to 0.5 ml (rotatory evaporation); puriﬁcation by Florisil column (10 ml pentane). Preconcentration: extract reduced to 1 ml, then with N2 ﬂow to just dryness; redissolution in 100 ml of the internal standard solution (dimethyldipentyltin in hexane). Separation: 30 m £ 0.32 mm £ 0.25 mm SPB-1 column

Internal standard Me2Pe2Sn; TBT 96%, DBT 97%, MBT 95%, MPhT 98%, DPhT 96%, TPhT 95%. All Aldrich

GC–FPD

continued

Sampling and sample treatment in the analysis of organotin compounds

Lab Analytical code technique

1013

1014

TABLE 30.7 (continuation) Species

Sample mass (mg)

Sample pretreatment

Calibration

12

HPLC– ID-ICPMS

TBT

350 –400

Extraction: sample þ 11 ml stainless steel ASE cell; mixed with “hydromatrix” to increase surface area for extraction; spiking with 117Sn enriched TBTCl; overnight equilibration; extraction with Dionex ASE 200 (5 cycles of 5 min, 1008C, 1,500 psi; 15–17 g 1 M sodium acetate/1 M acetic acid in methanol extraction solvent); dilution with ultrapure water. Separation: C18 reversed phase column; mobile phase 65:23:12 v/v/v acetonitrile/water/acetic acid with 0.05% vv TEA

14

GC–AED

300

Extraction: sample þ 10 ml TMAH 20% (60 min); addition of HCl to pH 7; addition of sodium acetate/ acetic acid buffer (pH 4); extraction with hexane; derivatisation with NaBEt4

17a

GC–MS

TBT, DBT, MBT, TPhT, DPhT, MPhT TBT, DBT, MBT

IDMS calibration blend of 1:1 TB120 SnCl: TB117SnCl used for calibration. The natural reference isotope TBTCl was puriﬁed to .98.5% purity (DSC) and 117 SnTBTCl was synthesized in house. Conﬁrmation of results by external calibration using puriﬁed TB120SnCl Calibration with ethylated standards; veriﬁcation with BCR-477

500

Extraction: sample þ 15 ml methanol/tropolone 0.03% þ 1 ml HCl; soniﬁcation (358C, 15 min), twice; liquid– liquid partitioning methylene chloride (15 ml), twice; solvent exchange to iso-octane; Derivatisation: Grignard alkylation with 1 ml pentylmagnesium bromide (2 M) in ethyl ether. Clean-up: silica-gel column (3 g), hexane/toluene solution (1:1, 10 ml). Separation: Hewlett-Packard HP-5MS capillary column (30 m £ 0.25 mm £ 0.25 mm), methyl-5% phenylsilicone

External calibration with pentylated standards. Pure calibrants TPrT (.99%), MBT (.98%), DBT (.99%), TBT (.98%), all IVM. TPrT as internal standard

R. Morabito

Lab Analytical code technique

TABLE 30.7 (continuation) Species

Sample mass (mg)

17b

GC –FPD

TBT, DBT, MBT

21

GC –ITDMS

TBT, DBT, MBT, TPhT, DPhT, MPhT

24

GC –FPD

TBT, DBT

Extraction: sample þ 15 ml methanol/tropolone (0.03%) þ 1 ml HCl; soniﬁcation (358C, 15 min), twice; liquid– liquid partitioning methylene chloride (15 ml), twice; solvent exchange to iso-octane; Derivatisation: Grignard alkylation with 1 ml pentylmagnesium bromide (2 M) in ethyl ether. Clean-up: silica-gel column (3 g), hexane/toluene solution (1:1, 10 ml). Separation: DB-1 capillary column (J&W) (30 m £ 0.53 mm £ 1.5 mm), 100% methylsilicone 200 Pretreatment: sample þ 5 ml water þ 2 ml HCl; let stand (16 h); þ 0.5 g NaCl; extraction with 2 £ 12 ml diethylether/tropolone (0.20 g/l); Grignard derivatisation (pentylation): þ 1.5 ml 2 M pentylmagnesium bromide in diethyl ether; let stand (5 min); þ 5 ml hexane, þ 5 ml 1 M NH4Cl in water; separate and concentrate hexane fraction to 1 ml. Clean-up: alumina (5 g fully activated), column (11 £ 150 mm) chromatography; elution 16 ml hexane/diethyl ether (4/1 vv); concentrate to 1 ml. Separation: Agilent Model 6890 GC– MSD; column (50 m £ 0.2 mm £ 0.33 mm) 500– 1000 Extraction: sample þ sufﬁcient 0.1% NaOH in methanol (1:1) to achieve a 4:1 MeOH:water (v/v) ratio (min 2 ml of water must be present); shaking (1 h); þ 2 ml hexane þ approx 100 mg sodium borohydride; shaking (20 min); centrifugation (2000 rpm, 5 min); collect hexane phase; no clean-up. Separation: Phenylmethyl silicone column (25 m £ 0.32 mm) 500

Sample pretreatment

Calibration

External calibration with pentylated standards. Pure calibrants TPrT (.99%), MBT (.98%), DBT (.99%), TBT (.98%), all IVM. TPrT as internal standard

External calibration with pentyl derivates of all six organotins: DBT and TBT (.98%); others (.99%), (QUASIMEME). Dibutyldihexyltin and triphenylhexyltin as internal standards added to diethyl/tropolone extract for recovery correction of butyltins and phenyltins, respectively TBT oxide 96%, DBT oxide 98% triphenyltin chloride 95%(TPhT). All Aldrich. Tripropyltin chloride .99%(TPT) as internal standard (CKWitco, Promochem) continued

Sampling and sample treatment in the analysis of organotin compounds

1015

Lab Analytical code technique

1016 TABLE 30.7 (continuation) Species

Sample mass (mg)

Sample pretreatment

Calibration

25

TBT, DBT, MBT, TPhT, DPhT

500

Extraction: sample þ 10 ml TMAH; soniﬁcation (90 min, 508C); after dissolution, þ 5 ml buffer þ 5 ml acetic acid (96% (MERCK) pH 5– 6; derivatisation and extraction (1,000 ml NaBEt4 þ 5 ml n-hexane); shaking for 30 min; centrifugation (5,000 rpm, 15 min, 08C); recovery of organic phase; concentration using N2 stream. Clean-up: SPE cartridges (1 g ﬂorisil, 10 ml n-hexane). Separation: HP-5 (Hewlett-Packard) capillary column (30 m £ 0.25 mm £ 0.25 mm), 5% phenyl methyl silicone

Calibrants: solution of dibutyltin (DBT) dichloride (98%), tributyltin (TBT) chloride (96%), diphenyltin (DPhT) dichloride (98%), triphenyltin (TPhT) chloride (99%) (all Merck) and monobutyltin (MBT) trichloride (95%) (Aldrich) spiked in the ﬁsh tissue IAEA-MA-A-2/OC to matrix match the calibration solutions. Tripropyltin chloride as internal standard (Merck)

GC –FPD

R. Morabito

Lab Analytical code technique

Sampling and sample treatment in the analysis of organotin compounds TABLE 30.8 CRMs available for organotin speciation analysis Material

Producer

Species

Certiﬁed values

RM 424

BCR

BCR 462R

BCR

BCR 477

BCR

BCR 646

BCR

BCR 710b

BCR

PACS-1

NRCC

PACS-2

NRCC

NIES-11

NIES

TBT a DBT a MBT a TBT DBT TBT DBT MBT TBT DBT MBT TPhT DPhT MPhT TBT DBT TBT DBT MBT TBT DBT MBT TBT TPhT a

20 ^ 5 (mg/kg) 53 ^ 19 257 ^ 54 54 ^ 15 (mg/kg) 68 ^ 12 2.20 ^ 0.19 (mg/kg) 1.54 ^ 0.12 1.50 ^ 0.28 480 ^ 80 (mg/kg) 770 ^ 90 610 ^ 120 29 ^ 11 36 ^ 8 69 ^ 18 135 ^ 27 (mg/kg) 82 ^ 19 1.27 ^ 0.22c (mg/kg) 1.16 ^ 0.19c 0.28 ^ 0.17c 0.98 ^ 0.13c (mg/kg) 1.09 ^ 0.15c 0.45 ^ 0.05c 1.3 ^ 0.1c (mg/kg) 6.3c

a

Indicative values. At the certiﬁcation stage. (The material has been proposed also for the certiﬁcation of arsenobetaine and methylmercury.) c As Sn. b

30.4

DETAILED PROCEDURE FOR THE GC –MS DETERMINATION OF ORGANOTIN COMPOUNDS IN ENVIRONMENTAL SAMPLES

As reported above, there is a lot of analytical procedures for organotin determination in environmental samples available in literature. Just as an example, the procedure adopted for solid samples by ENEA (Italy) is reported below. The procedure has been optimised, improved and validate along the almost 15 years of the EU tin speciation project.

1017

R. Morabito

The sample is homogenized and freeze-dried before extraction. TPrT, 50–500 ng, as internal standard, is added to 100 –500 mg of sample as methanolic solution before extraction, allowing 30 min for equilibration. Longer equilibration times, up to 16 h, do not affect absolute recovery of TPrT. The sample is placed in a Pyrex vial and 15 ml of tropolone 0.03% in methanol and 1 ml of concentrated HCl are sequentially added. The vial is put in an ultrasonic bath at a water temperature lower than 408C and left under sonication for 15 min. The vial is then transferred for centrifugation at 3000 rpm for 10 min. The surnatant is transferred in a 250 ml separatory funnel ﬁlled with 100 ml of a 10% NaCl solution and the extraction procedure is repeated. After the second surnatant is transferred to the separatory funnel, liquid– liquid partitioning is performed twice with 15 –20 ml of CH2Cl2. The methylene chloride phases are collected through anhydrous sodium sulphate. After washing the sulphate with 1–2 ml of CH2Cl2, the extract is added of 1 ml of iso-octane and the volume is approximately reduced to 5 ml in a rotary evaporator at a bath temperature lower than 408C and under moderate vacuum. The concentrated extract is transferred to a 15 ml vial and led to almost dryness under moderate ﬂow of nitrogen, operating solvent exchange (methylene chloride to iso-octane). One millilitre of 2 M ethereal solution of pentylmagnesium bromide is added and the mixture is shaken for 5 min. Two millilitre of distilled water is carefully added drop by drop and then 6–7 ml of H2SO4 1 M are added too. Derivatised organotins are extracted with 2–3 ml of hexane; the extraction is repeated twice. The organic phase is put in a vial and concentrated under moderate ﬂow of nitrogen to ca. 0.5 ml. The extract is transferred on top of a silica-gel column (3 g in a glass column 30 cm length and 8 mm as internal diameter) previously wet with 0.5 ml of hexane/toluene 1:1. Hexane/toluene 1:1 mixture is passed through the column until 5 ml is collected in a vial. Finally, the solution is concentrated under moderate ﬂow of nitrogen to ca 0.5 ml. For GC –MS determination, 1 ml is injected. Mass spectrometric detection systems have distinct advantages over the other speciﬁc detectors that have been used for organometallic compounds analysis (like AAS, ICP, AED, etc.): 1. they provide both sensitivity and selectivity together with structural conﬁrmation capabilities; 2. coupling to HRGC is straightforward and there is no need to “adapt” each other different instruments; 3. modern instrumentation is robust, easy to use, relatively inexpensive (benchtop instruments) and is now really at an industry-standard level of reliability; 4. there is no need to be an “MS specialist” to perform analyses. Mass spectrometry is particularly well suited for the analysis of organometallic species (provided they are amenable to GC): the metal atom

1018

Sampling and sample treatment in the analysis of organotin compounds TABLE 30.9 Natural abundances of main isotopes of Sn Mass number

Abundance (%)

116 117 118 119 120 122 124

14.30 7.61 24.03 8.58 32.85 4.72 5.94

(particularly the heaviest ones) gives the compound an “extra mass” with respect to the possible coeluting organic compounds, making detection and conﬁrmation easier and less prone to interferences. Tin has a “rich” isotopic composition as shown in Table 30.9. This isotopic distribution gives a peculiar aspect to the MS spectra of tin compounds: each fragment ion appears as a cluster of m=z values with the above mentioned ratios. From the analytical point of view, this improves identiﬁcation of the analyte but decreases GC/MS sensitivity because the ions produced by a single fragmentation are spread over a range of m=z: The fragmentation pattern under classical EI conditions of fully alkylated organotins is quite simple, consisting of a stepwise loss of the alkyl (aryl) groups. The scan mode detection of quadrupolar detectors cannot cope with the low environmental concentrations of organotins (the ion trap detector could be a valuable alternative for high sensitivity scan) and diagnostic ions have to be chosen for SIM. As a general rule, diagnostic ions for SIM have to fulﬁl some basic requirements: – – – –

they should be among the most abundant ions of the spectrum in order to increase sensitivity; they should be unique and characteristic of the compound in the matrix under analysis; the selected m=z should have the lowest possible instrumental noise in the experimental set-up; m=z values whose ratios are checked for conﬁrmation should be isotopic peaks rather than different fragment ions.

In the case of pentylated organotins, these requirements are easily met thanks to the simple fragmentation and the peculiar isotopic composition.

1019

R. Morabito

As an example, the optimised conditions for pentylated organotin analysis carried out on a Hewlett-Packard HP 5890 GC/ HP 5971 MSD system are as follows: † † † † † † †

electron impact ionisation mode (70 eV); carrier gas: helium, 80 kPa head pressure; column: HP-5 (methyl-5% phenylsilicone, 0.20 mm i.d., 0.11 mm ﬁlm thickness, 25 m length; Hewlett-Packard); temperature program: 808C £ 2 min, then 108C min21 to 2808C; injector: splitless, 2408C; transfer line temperature: 2808C; SIM (selected ions monitoring) operation with the following program (dwell time was 100 ms for all ions):

TPrT TBT DBT MBT Sn(IV) MPhT DPhT TPhT

Start time (min)

m=z

6 8 10 12 14 16 19 21

277, 275, 273 305, 303, 301 319, 317, 315 319, 317, 315 333, 331, 329 339, 337, 335 345, 343, 341 351, 349, 347

The timings reported above are only indicative and should be adapted to the particular instrumental conditions in use. Peak identiﬁcation was based on the matching of retention times (^ 0.5%) and isotopic mass ratios (^ 20%) for the diagnostic ions. The relative response factors were controlled by injecting standard mixtures on a regular basis (one injection every 3–4 samples) to follow the tuning conditions of the MS system. With these chromatographic settings, the limit of detection for TBT, DBT and MBT at a signal-to-noise ratio of 3 is around 1 pg injected. Phenyltins detection is somewhat more sensitive, particularly for TPhT (LOD ¼ 0.3 pg), owing to their peculiar fragmentation pattern. A full description of the advantages of GC –MS analysis for organotin analysis is given elsewhere [98]. ACKNOWLEDGEMENTS Dr Philippe Quevauviller (EU) is gratefully acknowledged for all the discussions had along the 15 years of the tin speciation project that were extremely precious for writing this chapter.

1020

Sampling and sample treatment in the analysis of organotin compounds

Dr Kees Kramer (Mermayde) is gratefully acknowledged for the valuable support in the coordination of the MULSPOT intercomparison and certiﬁcation exercises as well as for all the pleasant and interesting discussions. The ENEA colleagues, Dr Salvatore Chiavarini, Dr Carlo Cremisini, Mr Michele Fantini and Dr Paolo Massanisso, are gratefully acknowledged for the contribution to the development and optimization of the analytical procedure and to the success of the certiﬁcation campaigns described in the text as well as for the 15 years of pleasant working together in the organotin ﬁeld. REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26

S.J. Blunden, L.A. Hobbs and P.J. Smith, The Environmental Chemistry of Organotin Compounds, I.T.R.I. Publ. no. 640, I.T.R.I., London, 1984. S.J. Blunden and C.J. Evans, in: O. Hutzinger (Ed.), The Handbook of Environmental Chemistry, Vol. 3, Part E. 1990, pp. 1 –44. M. Hoch, Appl. Geochem., 16 (2001) 719. A.R.D. Stebbing, Mar. Poll. Bull., 16 (1985) 383. M.A. Champ and F.L. Lowenstein, Oceanus, 30 (1987) 69. S.J. Bushong, M.C. Ziegenfuss, M.A. Unger and L.W. Hall, Environ. Toxicol. Chem., 9 (1990) 359. P.F. Seligman, R.F. Lee, A.O. Valkirs and P.M. Stang, Proceedings of the 3rd International Organotin Symposium—Monaco, 1990, pp. 30–38. A.O. Valkirs, P.F. Seligman and R.F. Lee, Proceedings of the Oceans Organotin Symposium, Washigton DC, IEEE Piscataway NJ, 1986, Vol. 4, pp. 1165– 1170. J.J. Cooney, J. Ind. Microbiol., 3 (1988) 195. G.M. Gadd, FEMS Microbiol. Rev., 11 (1993) 297. S.J. Blunden and A.H. Chapman, Environ. Tech. Lett., 3 (1982) 267. P.M. Stang and P.F. Seligman, Proceedings of the Oceans—An International Workplace, Halifax, 1987, pp. 1386–1391. Ph. Quevauviller and O.F.X. Donard, Fresenius J. Anal. Chem., 339 (1991) 6. S. Chiavarini, C. Cremisini, R. Morabito, MAP Technical Reports Series No. 59, UNEP/FAO/IAEA, Athens, 1991, pp. 179– 187. D.H. Calloway and J.J. McCullen, Am. J. Chem. Nutr., 18 (1966) 1. H.A. Schroeder, J.J. Balassa and I.H. Tipton, J. Chronic Dis., 17 (1964) 483. P.J. Smith and L. Smith, Chem. Br., 11 (1975) 208. C.J. Evans and P.J. Smith, J. Oil Col. Chem. Assoc., 58 (1975) 160. R.B. Laughlin and O. Linden, Ambio, 14 (1985) 88. S.J. Blunden, P.J. Smith and B. Sugavanam, Pest. Sci., 15 (1984) 253. S.J. Blunden and A.H. Chapman, in: P.J. Craig (Ed.), Organometallic Compounds in the Environment. Longman Publ, 1986, pp. 111– 139. U.S. Environmental Protection Agency, U.S. EPA Position Document 2/3, Ofﬁce of Pesticides Programs, Washington, D.C., 1987. S. Chiavarini, C. Cremisini and R. Morabito, in: S. Caroli (Ed.), Element Speciation in Bioinorganic Chemistry. Wiley, New York, 1996, pp. 287– 329, Ch. 9. R.J. Maguire, Appl. Organomet. Chem., 1 (1987) 475. WHO, Environmental Health Criteria 116. World Health Organization, Geneva, 1990. P.E. Gibbs and G.W. Bryan, in: M.A. Champ and P.F. Seligman (Eds.), Organotin— Environmental Fate and Effects. Chapman & Hall, London, 1996, pp. 259– 281.

1021

R. Morabito 27 28 29 30 31

32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

1022

K. Fent, Crit. Rev. Toxicol., 26 (1996) 1. R.E. Anderson, Mar. Environ. Res., 17 (1985) 137. R.F. Lee, Mar. Biol. Lett., 2 (1981) 87. T.L. Wade, B. Garcia-Romero and J.M. Brooks, Environ. Sci. Technol., 22 (1988) 1488. A.M. Caricchia, S. Chiavarini, C. Cremisini, M. Fantini, R. Morabito, R. Scerbo and M. Vitali, Proceedings of Heavy Metals in the Environment, Toronto, 1993, pp. 52– 55. M.J. Waldock and J.E. Thain, Mar. Pollut. Bull., 14 (1983) 411. R.B. Laughlin, W. French and H.E. Guard, Environ. Sci. Technol., 20 (1986) 884. J.W. Short and F.P. Thrower, Proceedings of the Oceans Organotin Symposium, Washington DC, IEEE Piscataway NJ, 1986, Vol. 4, pp. 1202– 1205. C. Alzieu, J. Sanjuan, J.P. Deltreil and M. Borel, Mar. Poll. Bull., 17 (1986) 494. J.M. Ruiz, G. Bachelet, P. Caumette and O.F.X. Donard, Environ. Pollut., 93 (1996) 195. C. Alzieu, Proceedings of the Oceans Organotin Symposium, Washington DC, IEEE Piscataway NJ, 1986, Vol. 4, pp. 1130– 1134. P.E. Gibbs and G.W. Bryan, J. Mar. Biol. Assoc. UK, 66 (1986) 767. P.E. Gibbs, G.W. Bryan, P.L. Pascoe and G.R. Burt, J. Mar. Biol. Assoc. UK, 67 (1987) 507. P.E. Gibbs, P.L. Pascoe and G.R. Burt, J. Mar. Biol. Assoc. UK, 68 (1988) 715. E. Stroben, J. Oehlmann and P. Fioroni, Mar. biol., 113 (1992) 625. P.E. Gibbs, G.W. Bryan, L.G. Hummerstone and G.R. Burt, J. Mar. Biol. Assoc. UK, 66 (1986) 611. P.E. Gibbs and G.W. Bryan, in: K. Kramer (Ed.), Biomonitoring of Coastal Waters and Estuaries. CRC Press, Boca Raton, 1994, pp. 205– 226. C.C. ten Hallers-Tjabbes, J.F. Kemp and J.P. Boon, Mar. Pollut. Bull., 28 (1994) 311. B.P. Mensink, C.C. ten Hallers-Tjabbes, J. Kralt, I.L. Freriks and J.P. Boon, Mar. Environ. Res., 41 (1996) 315. C. Swennen, N. Ruttanadakul, S. Ardseungnern, H.R. Singh, B.P. Mensink and C.C. ten Hallers-Tjabbes, Environ. Technol., 18 (1997) 1245. S.M. Evans and J. Nicholson, Sci. Total Environ., 258 (2000) 73. C.M. Rees, B.A. Brady and G.J. Fabris, Mar. Pollut. Bull., 42 (2001) 873. A.C. Birchenough, N. Barnes, S.M. Evans, H. Hinz, I. Kronke and C. Moss, Mar. Pollut. Bull., 44 (2002) 534. M. Ramon and M.J. Amor, Mar. Environ. Res., 52 (2001) 463. V. Axiak, A.J. Vella, D. Micaleff and P. Chircop, Mar. Biol., 121 (1995) 685. M. Sole´, Y. Morcillo and C. Porte, Environ. Pollut., 99 (1998) 241. A. Terlizzi, S. Geraci and P.E. Gibbs, Ital. J. Zool., 66 (1999) 141. A. Terlizzi, S. Geraci and V. Minganti, Mar. Pollut. Bull., 36 (1998) 749. S. Chiavarini, P. Massanisso, P. Nicolai, C. Nobili and R. Morabito, Chemosphere, 50 (2003) 311. T. Horiguchi, H. Shiraishi, M. Shimizu and M. Morita, Environ. Poll., 95 (1997) 85. K.S. Tan, Mar. Pollut. Bull., 39 (1999) 295. M. Gooding, C. Gallardob and G. LeBlanc, Mar. Pollut. Bull., 38 (1999) 1227. C. Alzieu, Proceedings of the 3rd International Organotin Symposium, Monaco, 1990, pp. 1– 2. M.A. Champ, Proceedings of the Oceans Organotin Symposium, Washington DC, IEEE Piscataway NJ, 1986, Vol. 4, pp. 1– 8.

Sampling and sample treatment in the analysis of organotin compounds 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94

UNEP/FAO/WHO/IAEA, MAP Technical Reports Series No. 33, Athens, 185 p. 1989. H. Vrijhof, Sci. Total Environ., 43 (1985) 221. R.J. Huggett, M.A. Unger, P.F. Seligman and A.O. Valkirs, Environ. Sci. Technol., 26 (1992) 232. M.A. Champ, Sci. Total Environ., 258 (2000) 21. Ph. Quevauviller, Method Performance Studies for Speciation Analysis. The Royal Society of Chemistry, Cambridge, 1998, 271 pp. S. Chiavarini, C. Cremisini, T. Ferri, R. Morabito and A. Perini, Sci. Total Environ., 101 (1991) 217. A.M. Caricchia, S. Chiavarini, C. Cremisini, R. Morabito and R. Scerbo, Anal. Sci., 7 (1991) 1193. A.M. Caricchia, S. Chiavarini, C. Cremisini, M. Fantini and R. Morabito, Sci. Total Environ., 121 (1992) 133. I. Tolosa, L. Merlini, N. de Bertrand, J.M. Bayona and J. Albaiges, J. Environ. Toxicol. Chem., 11 (1992) 145. I. Tolosa, J.W. Readman, A. Blaevoet, S. Ghilini, J. Bartocci and M. Horvat, Mar. Pollut. Bullet., 32 (1996) 335. B.S. Tselentis, M. Maroulakou, J.F. Lascourreges, J. Szpunar, V. Smith and O.F.X. Donard, Mar. Pollut. Bull., 38 (1999) 146. P. Michel, B. Averty, B. Andral, J.F. Chiffoleau and F. Galgani, Mar. Pollut. Bull., 42 (2001) 1128. S.D. Dıez, M.J. Abalos and J.M. Bayona, Water Res., 36 (2002) 905. J.A. Stab, W.P. Coﬁno, B. Van Hattum and U.A.Th. Brinkman, Anal. Chim. Acta, 286 (1994) 335. R. Ritsema, Appl. Organomet. Chem., 8 (1994) 5. S. Biselli, K. Bester, H. Huhnerfuss and K. Fent, Mar. Pollut. Bull., 40 (2000) 233. A. Albalat, J. Potrykus, J. Pempkowiak and C. Porte, Chemosphere, 47 (2002) 165. P.H. Dowson, J.M. Bubb and J.N. Lester, Mar. Pollut. Bull., 26 (1993) 487. Ph. Quevauviller, O.F.X. Donard and H. Etcheber, Environ. Pollut., 84 (1994) 89. J.L. Gomez-Ariza, R. Beltran, E. Morales, I. Giraldez and M. Ruiz-Benitez, Appl. Organomet. Chem., 9 (1995) 51. P. Michel and B. Averty, Mar. Pollut. Bull., 38 (1999) 268. M. Barroso and M.H. Moreira, Mar. Pollut. Bull., 44 (2002) 480. T.L. Wade, B. Garcia Romero and J.M. Brooks, Chemosphere, 20 (1990) 647. B.L. McGee, C.E. Schlekat, D.M. Boward and T.L. Wade, Ecotoxicology, 4 (1995) 39. C.A. Krone, J.E. Stein and U. Varanasi, Environ. Monit. Assess., 40 (1996) 75. R.J. Maguire, Environ. Sci. Technol., 18 (1984) 291. R.J. Maguire, R.J. Tkacz, Y.K. Chau, G.A. Bengert and P.T.S. Wong, Chemosphere, 15 (1986) 253. S.J. de Mora, C. Steward and D. Phillips, Mar. Pollut. Bull., 30 (1995) 50. K. Kannan, S. Tanabe, H. Iwata and R. Tatsukawa, Environ. Pollut., 90 (1995) 279. M.C. Ko, G.C. Bradley, A.H. Neller and M.J. Broom, Mar. Pollut. Bull., 31 (1995) 249. S. Kan-Atireklap, S. Tanabe and J. Sangansin, Mar. Pollut. Bull., 34 (1997) 894. H. Harino, M. Fukushima, Y. Yamamoto, S. Kawai and N. Miyazaki, Arch. Environ. Contam. Toxicol., 35 (1998) 558. H. Harino, M. Fukushima and S. Kaway, Environ. Pollut., 105 (1998) 1. A.A. Ansari, I.B. Singh and H.J. Tobschall, Sci. Total Environ., 223 (1998) 157.

1023

R. Morabito 95 96 97 98

99 100 101 102 103 104

105 106 107 108 109 110 111 112

113 114 115 116 117

118 119 120 121 122 123

1024

J.A. Jacobsen and G. Asmund, Sci. Total Environ., 245 (2000) 131. J.J. Cooney and S. Wuertz, J. Ind. Microbiol., 4 (1989) 375. C. Steward and S.J. de Mora, Environ. Sci. Technol., 11 (1990) 565. R. Morabito, S. Chiavarini and C. Cremisini, in: B. Griepink, E.A. Maier and Ph. Quevauviller (Eds.), Quality Assurance of Environmental Analysis Within the BCR-Programme. Elsevier, New York, 1995, pp. 435–464, Ch. 17. Ph. Quevauviller and O.F.X. Donard, in: S. Caroli (Ed.), Element Speciation in Bioinorganic Chemistry. Wiley, New York, 1996, pp. 331–362, Ch. 10. S.J. de Mora (Ed.), Tributyltin: Case Study of an Environmental Contaminant, Cambridge Environmental Chemistry Series 8, 1996, p. 301. M. Abalos, J.M. Bayona, R. Compano`, M. Granados, C. Leal and M.D. Prat, J. Chromatogr. A, 788 (1997) 1. L. Ebdon, S.J. Hill and C. Rivas, Trends Anal. Chem., 17 (1998) 277. S. Tanabe, Mar. Pollut. Bull, 39 (1999) 62. O.F.X. Donard, G. Lespes, D. Amouroux and R. Morabito, in: L. Ebdon, L. Pitts, H. Crews, O.F.X. Donard and Ph. Quevauviller (Eds.), Trace Element Speciation for Environment, Food and Health. RSC Publ, 2001, pp. 142–175, Ch. 8. R. Morabito, Microchem. J., 51 (1995) 198. R. Morabito, in: N.S. Thomaidis and T.D. Lekkas (Eds.), Proceedings of Metal Speciation in the Environment. Global Nest Publ, 2000, pp. 49–64. R. Scerbo, Thesis, unpublished results. P. Michel, B. Averty, B. Andral, J.F. Chiffoleau and F. Galgani, Mar. Pollut. Bull., 11 (2001) 1128. D.R. Young, P. Schatzberg, F.E. Brinckman, M.A. Champ and S.E. Holm, Proceedings of Ocean 86—Washington D.C., 1986, pp. 1135– 1140. Ph. Quevauviller, A. Bruchet and O.F.X. Donard, Appl. Organomet. Chem., 5 (1991) 125. C.A. Dooley and V. Homer, Naval Ocean System Center Technical Report No. 917, San Diego, CA, 1983. G. Batley, in: S.J. de Mora (Ed.), Tributyltin: Case Study of an Environmental Contaminant, Cambridge Environmental Chemistry Series, 1996, pp. 139– 166, Ch. 5. J.J. Cleary and A.R.D. Stebbing, Proceedings Ocean 87—An International Workplace—Halifax, 1987, pp. 1405– 1410. P.M. Stang, R.F. Lee and P.F. Seligman, Environ. Sci. Technol., 26 (1992) 1382. D. Amouroux, E. Tessier and O.F.X. Donard, Environ. Sci. Technol., 17 (2000) 494. Ph. Quevauviller and O.F.X. Donard, Appl. Organomet. Chem., 4 (1990) 353. S. Chiavarini, C. Cremisini, T. Ferri, M. Fantini, R. Morabito and A. Perini, Environmental Contamination—Proceedings of 4th International Conference— Barcelona, 1990, pp. 514– 516. L. Ebdon, K. Evans and S. Hill, Sci. Total Environ., 68 (1988) 207. C. Clavell, P.F. Seligman and P.M. Stang, Proceedings Ocean 86, Washington D.C., 1986, pp. 1152– 1154. P. Gy, Sampling for Analytical Purposes. C.H.I.P.S. Publ, 1998, 172 pp. Ph. Quevauviller, M.B. de la Calle, E.A. Maier and C. Camara, Mikrokim. Acta, 118 (1995) 131. J.L. Gomez-Ariza, E. Morales, D. Sanchez-Rodas and I. Giraldez, Trends Anal. Chem., 19 (2000) 200. H.A. Meinema, T. Burger-Wiersma, G. Versleris-de Hann and E.C. Gevers, Environ. Sci. Technol., 12 (1978) 288.

Sampling and sample treatment in the analysis of organotin compounds 124 125 126 127 128 129 130 131 132 133 134 135 136 137 138 139 140

141 142 143 144 145 146 147 148 149 150 151 152 153

R.J. Maguire, J.H. Carey and E.J. Hale, J. Agric. Food Chem., 31 (1983) 1060. K. Bergmann, U. Rohr and B. Neidhart, Fresenius J. Anal. Chem., 349 (1994) 815. J.L. Gomez-Ariza, I. Giraldez, E. Morales, F. Ariese, W. Coﬁno and Ph. Quevauviller, J. Environ. Monit., 1 (1999) 197. W.R. Blair, G.H. Olson, F.E. Brinckman, R.C. Paule and D.A. Becker, Natural Bureau of Standards, Gaithersburg, MD, 1986. C. Bancon-Montigny, G. Lespes and M. Potin-Gautier, Water Res., 35 (2001) 224. E.A. Boettner, G.L. Ball, Z. Hollingswort and R. Aquino, U.S. Environ. Protection Agency EPA-600/1-81-062, Cincinnati, 1981. J.L. Gomez-Ariza, E. Morales, R. Beltran, I. Giraldez and M. Ruiz-Benitez, Quım. Anal., 13 (1994) 76. W.C. Li, S. Zhang, Y.K. Chau and A.S.Y. Chau, NWRI Contribution No. 90-139, 1990, pp. 1. W.C. Li, S. Zhang, Y.K. Chau and A.S.Y. Chau, NWRI Contribution No. 90-146, 1990, pp. 1. A. Lamberty, Ph. Quevauviller and R. Morabito, EU Report, EUR 18406 EN, 1998. A.M. Caricchia, S. Chiavarini, C. Cremisini, R. Morabito and R. Scerbo, Anal. Chim. Acta, 286 (1994) 329. Ph. Quevauviller, R. Morabito, L. Ebdon, W. Coﬁno, H. Muntau and M.J. Campbell, EUR Report 17921 EN, Brussels, 1997. R. Morabito, K. Kramer, H. Muntau, Ph. Quevauviller and S. Bowadt, EUR Report, Brussels, in preparation. R. Morabito, Fresenius J. Anal. Chem., 351 (1995) 378. Ph. Quevauviller and R. Morabito, Trends Anal. Chem., 19 (2000) 86. W.M.R. Dirkx, R. Lobinski and F.C. Adams, Anal. Chim. Acta, 286 (1994) 309. W.M.R. Dirkx, R. Lobinski and F.C. Adams, in: Ph. Quevauviller, E.A. Maier and B. Griepink (Eds.), Quality Assurance for Environmental Analysis. Elsevier Science, Amsterdam, 1995, pp. 357–409, Ch. 15. M. Ceulemans and F. Adams, Anal. Chim. Acta, 317 (1995) 161. C. Pellegrino, P. Massanisso and R. Morabito, Trends Anal. Chem., 19 (2000) 97. J.L. Gomez-Ariza, E. Morales, I. Giraldez, D. Sanchez-Rodas and A. Velasco, J. Chromatogr. A, 938 (2001) 211. S. Simon, M. Bueno, G. Lespes, M. Mench and M. Potin-Gautier, Talanta, 57 (2002) 31. M.B. de la Calle-Guntina`s, R. Scerbo, S. Chiavarini, Ph. Quevauviller and R. Morabito, Appl. Organomet. Chem., 11 (1997) 693. A.M. Caricchia, S. Chiavarini, C. Cremisini, R. Morabito and C. Ubaldi, Int. J. Environ. Anal. Chem., 53 (1993) 37. J.L. Gomez-Ariza, E. Morales, R. Beltran, I. Giraldez and M. Ruiz-Benitez, Analyst, 120 (1995) 1171. O.F.X. Donard, B. Lale´re, F. Martin and R. Lobinski, Anal. Chem., 67 (1995) 4250. C.A. Krone, D.W. Brown, D.G. Burrows, R.G. Bogar, S.L. Chan and U. Varanasi, Mar. Environ. Res., 27 (1989). S. Zhang, Y.K. Chau, W.C. Li and A.S.Y. Chau, Appl. Organomet. Chem., 5 (1991) 431. T.J. Gremm and F.H. Frimmel, Water Res., 26 (1993) 1163. M. Ceulemans, C. Witte, R. Lobinski and F.C. Adams, Appl. Organomet. Chem., 8 (1994) 451. G.B. Jiang, P.S. Maxwell, K.W.M. Siu, V.T. Luong and S.S. Berman, Anal. Chem., 63 (1991) 1506.

1025

R. Morabito 154 155 156 157 158 159 160 161 162 163 164 165 166 167

168 169 170 171 172 173 174 175

1026

F. Pannier, A. Astruc and M. Astruc, Anal. Chim. Acta, 287 (1994) 17. U.T. Cumar, J.G. Dorsey and J.A. Caruso, J. Chromatogr. A, 654 (1993) 261. M.J. Waldock and M.E. Waite, Appl. Organomet. Chem., 8 (1994) 649. M. Nagase and K. Hasebe, Anal. Sci., 9 (1993) 517. J. Kuballa and R.D. Wilken, Analyst, 120 (1995) 667. Y.K. Chau, F. Yang and M. Brown, Anal. Chim. Acta, 338 (1997) 51. F. Pannier, A. Astruc, M. Astruc and R. Morabito, Appl. Organomet. Chem., 10 (1996) 471. J. Szpunar, P. Pellerin, A. Makarov, T. Doco, P. Williams and R. Lobinski, J. Anal. At. Spectrom., 14 (1999) 639. C.G. Arnold, M. Berg, S.R. Muller, U. Dommann and R.P. Schwarzenbach, Anal. Chem., 70 (1998) 3094. S. Chiron, S. Roy, R. Cottier and R. Jeannot, J. Chromatogr. A, 879 (2000) 137. Guidelines for the Production of BCR Reference Materials, Doc. BCR/48/93, European Commission, DG XII-C, rue de la Loi 200, 1049 Brussels, Belgium, 1994. R. Morabito, H. Muntau, W. Coﬁno and Ph. Quevauviller, J. Environ. Monit., 1 (1999) 75. P.H. Quevauviller, F. Ariese, W. Coﬁno, G.N. Kramer, T. Linsinger and M.J. Campbell, EUR Report, EUR 19773 EN, European Commission, Brussels, 2001. R. Ritsema, F.M. Martin and Ph. Quevauviller, in: Ph. Quevauviller, E.A. Maier and B. Griepink (Eds.), Quality Assurance for Environmental Analysis. Elsevier Science, Amsterdam, 1995, pp. 490 –500, Ch. 19. R. Morabito, P. Massanisso and Ph. Quevauviller, Trends Anal. Chem., 19 (2000) 113. Ph. Quevauviller, M. Astruc, L. Ebdon, H. Muntau, W. Coﬁno, R. Morabito and B. Griepink, Mikrochim. Acta, 123 (1996) 163. Ph. Quevauviller, M. Astruc, L. Ebdon, S. Sparkes, G.N. Kramer and B. Griepink, EUR Report EN 15901. European Commission, Brussels, 1994. Ph. Quevauviller, M. Astruc, L. Ebdon, G.N. Kramer and B. Griepink, EUR Report EN 15337. European Commission, Brussels, 1994. R. Morabito, P. Soldati, M.B. de la Calle and Ph. Quevauviller, Appl. Organomet. Chem., 12 (1998) 621. Ph. Quevauviller, F. Ariese, W. Coﬁno, G.N. Kramer, T. Linsinger and M.J. Campbell, EUR Report EN 19773. European Commission, Brussels, 2001. Ph. Quevauviller, Method Performance Studies for Speciation Analysis. RSC Publ, 1998, Ch. 5, pp. 69– 103. Ph. Quevauviller, M. Astruc, R. Morabito, F. Ariese and L. Ebdon, Trends Anal. Chem., 19 (2000) 180.

Chapter 31

Sample preparation for arsenic speciation Walter Goessler and Doris Kuehnelt

31.1

INTRODUCTION

For many centuries arsenic was known to be the element with two faces. On the one hand, arsenic, or rather its compounds, were used in special formulations for curing various diseases, as described by Allesch [1]. As an example Salvarsan (arsphenamine) was the only medicine for curing syphilis until the discovery of penicillin [2]. On the other hand, arsenic (arsenic trioxide) was known as the king of poisons because of its lack of smell and taste. Recently, the use of arsenic as poison through the ages was reviewed by Nriagu [3]. The acute toxic properties of arsenic—or rather its compounds—are certainly the reasons for all the research efforts devoted to this element in the past. Nowadays, millions of people, especially in West Bengal and Bangladesh, are exposed long-term to high levels of inorganic arsenic via the drinking water. Such exposure causes an increased risk of skin, bladder, and kidney cancer. In addition to this elevated risk of cancer, cardiovascular and neurological effects have also been attributed to the exposure to inorganic arsenic [4]. Today, this human tragedy is the driving force for many research projects focussing on arsenic, as recently reviewed by Chakraborti et al. [5]. For simpliﬁcation, in this chapter the various ions deriving from arsenous acid [As(III)] and arsenic acid [As(V)] will not be distinguished according to their degree of protonation. For example, the ions deriving from arsenic acid 22 (H3AsO4), dihydrogen arsenate (H2AsO2 4 ), hydrogen arsenate (HAsO4 ), and arsenate (AsO32 ) will always be referred to as arsenate [As(V)]. Methylarsonic 4 acid (MA), methylarsonous acid [MA(III)], dimethylarsinic acid (DMA), and dimethylarsinous acid [DMA(III)] will also not be distinguished according to their degree of protonation but will be, in contrast to As(V) and As(III), referred to as the acids. Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

1027

W. Goessler and D. Kuehnelt

31.2

OCCURRENCE AND DISTRIBUTION OF ARSENIC IN THE ENVIRONMENT

Although arsenic is only ranked 51st in elemental abundance in the earth’s crust, it is ubiquitous in our environment, albeit at low concentrations [6]. About 30 arsenic compounds have been identiﬁed thus far. The chemical structures of the most common are listed in Figs 31.1 and 31.2. Although recent ﬁndings have revealed that there is not much difference in the arsenic speciation of the marine and the terrestrial environment, these two ecosystems are discussed separately. 31.2.1 Marine environment Besides the importance of arsenic in the forensic sciences, due to its abuse as suicidal and homicidal poison, the fact that high arsenic concentrations were detected in seafood forced researchers to look into detail which form of arsenic is present. Although the arsenic concentration in seawater is usually below 2 mg As/l, Jones [7] found high arsenic concentrations in marine algae (5 – 94 mg As/kg dry mass). He speculated that different forms of arsenic must exist in the marine environment, because no toxic effects could be observed after consumption of algae with arsenic concentrations up to , 100 mg As/kg dry mass. Four years later, in 1926, Chapman [8] completed a systematic study of arsenic concentrations in marine algae and ﬁsh and concluded the presence of a more or less complex arsenic containing organic compound of low toxicity. Today the average arsenic intake of a person living in Japan, a country with a diet typically rich in seafood, was estimated by Yamauchi et al. [9] to be 273 mg As/day. In contrast, in the United States the mean total arsenic intake is ,50 mg As/day [10]. It is obvious that, especially in countries with seafood rich diets, high pressure from the public encouraged researchers to identify the arsenic compounds present in seafood because of the bad reputation of arsenic. In 1977, Edmonds et al. [11], an Australian research team, identiﬁed for the ﬁrst time the arsenic compound present in the Western rock lobster to be arsenobetaine (AB). Subsequent work revealed that AB is the dominant arsenic compound in all marine animals. About 4 years later, Edmonds and Francesconi [12] identiﬁed arsenosugars (dimethylarsinoylribosides) as the dominant arsenic compounds in brown kelp. Presently, 16 arsenosugars have been identiﬁed in marine algae, but only four (Fig. 31.2) are usually present at high concentrations. Typical total arsenic concentrations in marine animals and plants including algae, ﬁsh, and crustaceans, range from 0.5 to 50 mg As/ kg (wet mass). Data have been recently compiled by Francesconi and Kuehnelt [13]. With the identiﬁcation and the subsequent synthesis of AB, toxicological studies revealed that AB is not metabolised in the human body and, therefore, does not pose any risk to humans. The acute toxicity (LD50 mouse oral) of AB

1028

Sample preparation for arsenic speciation

Fig. 31.1. Chemical structures of the most common environmental arsenic compounds (abbreviations used in the text in brackets).

1029

W. Goessler and D. Kuehnelt

Fig. 31.2. Chemical structures of the four most common arsenosugars.

was determined by Kaise et al. [14] to be .10 g/kg body mass. This means that AB is about three orders of magnitude less acute toxic than As(III) and As(V). Thus far, only limited toxicity data are available for arsenosugars. Difﬁculties in the synthesis of these compounds on a large scale are certainly to blame for this lack of data. Nevertheless, authorities assume that no risk for humans occurs after consumption of marine algae. In a recent study conducted by Francesconi et al. [15], it was shown that after a single oral dose of arsenosugar 1 (Fig. 31.2), up to 12 metabolites could be detected in the urine of the consumer. About half of the metabolites have thus far not been identiﬁed. Further research is necessary to identify these unknown metabolites to ensure exclusion of any risk to humans. For further, more detailed information about arsenic in the marine environment, two review articles published by Francesconi and Edmonds [16,17] are highly recommended. More recent ﬁndings about arsenic in the marine environment have been summarised by Francesconi and Kuehnelt [13].

1030

Sample preparation for arsenic speciation

31.2.2 Terrestrial environment Whereas the marine environment has been studied intensively, less research effort has been devoted to the terrestrial ecosystem because of the lack of appropriate analytical techniques to handle the low arsenic concentrations. Typical arsenic concentrations in rocks range from 0.5 to 2.5 mg As/kg. In areas with high volcanic activity, such as Japan or Mexico the soil arsenic concentrations can be at least one order of magnitude higher than the average soil arsenic concentration of less than 10 mg As/kg [18]. Apart from the arsenic input into the biosphere through weathering processes and volcanic activities, the human contribution has been estimated to be 40% of the total arsenic input [19]. The use of arsenic containing insecticides and herbicides is forbidden nowadays in many countries. Arsenic compounds are still in use as wood preservatives, in glass the industry, and in farming for protection against parasitic diseases. Additionally, there is a big demand for arsenic in the electronic industries for semiconductor production. Besides the anthropogenic input of arsenic into our environment, microbiological activities are responsible for annually releasing ,2 £ 107 kg of arsenic from land surfaces into the atmosphere, as noted by Frankenberger and Arshad [20]. For a long time it was thought that arsenic compounds in the terrestrial environment were restricted to As(III), As(V), MA, DMA, and trimethylarsine [21]. The more complex arsenic compounds such as AB and arsenosugars, seemed to be a privilege of marine biota. With improvement of analytical methods, especially with the use of ICP-MS as an arsenic-speciﬁc detector, Byrne et al. [22] identiﬁed AB for the ﬁrst time in terrestrial mushrooms in 1995. Two years later, arsenocholine (AC) and the tetramethylarsonium cation (TETRA) were discovered by Kuehnelt et al. [23,24] in the mushroom Amanita muscaria collected at a former arsenic roasting facility. After these reports several research groups devoted their efforts to the analysis of terrestrial samples for arsenic compounds. Arsenoriboses were detected for the ﬁrst time in earthworms by Geiszinger et al. [25]. Nowadays, it is well accepted that identical arsenic compounds are present in marine and in terrestrial biota, albeit at lower concentrations and different concentration ratios. Typical arsenic concentrations in terrestrial plants from uncontaminated sites seldom exceed 20 mg As/kg (wet mass). As an exception, mushrooms have to be mentioned, which may contain several mg As/kg (wet mass) [26]. 31.2.3 Humans Humans are exposed to arsenic via air, food, and drinking water. Except from burning arsenic rich coals and workplace exposure in certain industries, exposure to humans via air is minor. Total arsenic concentrations derived from food as well as drinking water are certainly the most important sources of exposure. In the case of food, one has to keep in mind that only seafood

1031

W. Goessler and D. Kuehnelt

(sometimes mushrooms) contributes substantially to the total arsenic burden. Fortunately, only a few percent of the arsenic present in seafood is inorganic arsenic [27]. The highest health risk for humans is certainly the consumption of arsenic contaminated water, because almost all the arsenic is present either as As(V) or As(III). Although the World Health Organisation [28] has deﬁned a drinking water limit of 10 mg As/l to be safe, there are still some countries in the world having a limit of 50 mg As/l. The usefulness of discussing which limits are safe is questionable, especially when even the higher limits are exceeded in some countries, such as Bangladesh due to the geological situation. Tube wells with arsenic concentrations in the mg As/l range are not uncommon, especially in West Bengal and Bangladesh where millions of people suffer from chronic arsenicism which is manifested in elevated skin and internal cancer rates [5]. Usage of surface water as drinking water is not an alternative because of the bacteriological situation. High temperatures and high humidity in this part of the world are ideal for bacterial growth. Other actions, such as collection of rain water or labelling safe tube wells (arsenic concentration ,50 mg As/l) green and the ones with arsenic concentrations above 50 mg As/l, red have not been very successful until now. Many researchers are still looking for an appropriate solution that is on the one hand affordable and on the other hand acceptable for the people living in these areas. Humans excrete most ingested arsenic via the kidneys. Urinary arsenic concentrations of unexposed people are typically below 10 mg As/l [29]. The ratio of the excreted arsenic compounds is typically 60– 80% DMA, 10 –30% inorganic arsenic [sum of As(III) and As(V)], and 10–20% MA. These numbers can be signiﬁcantly changed by recent (3 –4 days) seafood consumption. Recently, it was shown that there appears to be a polymorphism in arsenic methylation. Whereas Andean people from north-west of Argentina have low percentages of MA (, 2%), in their urine, people from Taiwan have an average relative MA concentration of 27% [29]. The biomethylation of inorganic arsenic is considered to occur via reduction of As(V) to As(III), addition of a methyl group from S-adenosylmethionine, reduction of the formed MA to MA(III), and further transfer of a methyl group to give DMA. Still, little is known about the enzymes involved in this process. The liver seems to be an important site for arsenic methylation, but methylation has been documented in other tissues as well. The organic forms (DMA and MA) are less reactive with tissue constituents, less toxic, and faster excreted in the urine than As(III) and As(V). As(III) was found to be the arsenic compound interacting most with tissue [29]. The methylation of arsenic, and therefore faster excretion from the human body, was always believed to be a detoxiﬁcation process. Recently, MA(III) was discovered by Aposhian et al. [30] to be present in urine of humans exposed to inorganic arsenic. Both MA(III) and DMA(III) were reported by Le et al. [31] in human urine from a person 4 h after administration of 300 mg sodium 2,3-dimercapto-1-propane sulfonate. Subsequent work on cultured cells showed that the two trivalent organoarsenic

1032

Sample preparation for arsenic speciation

compounds are at least as toxic as the inorganic arsenic compounds. Moreover, these two trivalent arsenicals are proposed by Mass et al. [32] to have a very high genotoxicity. These new ﬁndings raise the question of whether the methylation of arsenic in the human body is really a detoxiﬁcation process. There are only a few studies on arsenic speciation in blood. The reasons for this are on the one hand the short half-life of arsenic in blood and on the other hand the difﬁculties in the determination of arsenic compounds in blood or its compartments. Shibata et al. [33] found AB in serum. In a Belgian study, AB and DMA were detected in uremic patients [34]. Due to the high afﬁnity of arsenic to cysteine, which is present in high concentrations in the keratin of the hair, arsenic is accumulated in hair. In contrast to urinary arsenic, which is comparable to a photograph (3 –4 days back), the arsenic concentration in hair is a kind of ﬁlm that, depending on the length of the hair (growth typically 1 cm per month), can provide information about previous exposure. Total arsenic concentration in the hair of unexposed persons seldom exceeds 200 mg As/kg. In hair samples of exposed people 10 times higher values are not unusual [5]. Arsenic speciation in hair is difﬁcult because no appropriate extraction technique is available. The few studies, thus far have revealed that dimethylated species are present. 31.3

STABILITY OF ARSENIC COMPOUNDS

Whereas for total arsenic determinations the sample matrix and often concurrently the arsenic compounds have to be destroyed, a destruction of the sample matrix is not needed for speciation analysis. For speciation analysis, the arsenic compounds must be extracted from the sample. The extraction step should be quantitative but must not destroy any arsenic compounds present. When the question is raised whether the extraction process itself has already altered the species information, one has to reply certainly with “yes”. It is very unlikely that the different arsenic compounds are “swimming” freely in the cytosol without being attached to other cell compartments. As long as different extraction methods (the milder the better) produce similar results with respect to the determined species, we can assume that these compounds were originally present in the sample and are not operationally deﬁned results. When enzymatic digestion is used for destruction of the organic matrix (as often is in case for selenium) and thereafter the fractions (selenoaminoacids) of the proteins are determined, it is difﬁcult to speak about speciation analysis, because the macromolecule (selenium containing compound) is deﬁnitely destroyed [35]. Unfortunately, in situ determination of arsenic compounds at environmental concentrations is not possible with the techniques available at the moment. Xray absorption ﬁne structure spectroscopy has shown to be a possible way of direct speciation analysis, albeit high arsenic concentrations are required as recently described by Langdon et al. [36].

1033

W. Goessler and D. Kuehnelt

Storage of samples in a freeze-dried status over an inert atmosphere, as well as freezing of the samples are often employed. Some detailed information about stabilities of arsenic compounds is given below. 31.3.1 Arsenite and arsenate These two arsenic compounds are commonly found in our environment. As(V) is the dominant arsenic compound in seawater. In marine biota, As(III) as well as As(V) have been reported as minor species, with the exception of the marine alga Hizikia fusiforme, in which As(V) can be found at high concentrations (up to 50%). In the terrestrial ecosystem these two arsenic compounds contribute substantially to the total arsenic concentration. In glass manufacturing, As2O3 is often employed for clearing the glass. This might pose a signiﬁcant problem when glassware (also autosampler vials) are used for speciation analysis of As(III) and As(V) at trace concentrations. As(III) and As(V) are easily interconverted and, therefore, are often found together in real samples. Under oxidising conditions, As(V) is the thermodynamically favoured form and generally found in environmental samples. Under reducing conditions, As(III) is thermodynamically favoured. Arsenites are more soluble and, therefore, more mobile than arsenates. Inskeep et al. [37] recently described the nonequilibrium behaviour of the As(V)/As(III) couple. Arsenites are observed in oxic environments and arsenates persist in anoxic systems. Slow kinetics and/or biological phenomena were invoked to explain the apparent lack of thermodynamic equilibrium. Stock standard solutions of As(V) prepared from Na2HAsO4·7H2O and As(III) prepared from NaAsO2 at a concentration of 1 g As/l can be stored at 48C without any changes. In high alkaline solutions (pH . 12), As(III) is oxidised to As(V) as described by Kuehnelt et al. [38]. At low concentrations (up to 5 mg As/l), we sometimes observe in our laboratory a complete interconversion of As(III) to As(V) or vice versa for aqueous standard solutions. The standard solutions are ﬁlled in closed (crimbed) polypropylene autosampler vials and stored overnight. These phenomena cannot be observed all the time. The samples are always prepared from the same stock solutions with the same water quality. A partial oxidation/ reduction at higher concentrations was not observed thus far. These observations make clear that is very difﬁcult to determine the exact concentrations of As(III) and As(V) at low concentrations. In a recently published article by Bednar et al. [39], it was shown that addition of EDTA for complexation of metals present in natural water can be successfully applied for preserving the species information until analysis in the laboratory. Extraction of spiked As(III) and As(V) from a plant matrix using pressurised liquid extraction (also known as accelerated solvent extraction, ASE) in a temperature range from 60 to 1808C revealed that As(V) was stable up to 1508C whereas at 1208C only ,70% of the As(III) were recovered [40]. Surprisingly, no correlation was found between As(III) diminution and

1034

Sample preparation for arsenic speciation

an increase of As(V). Vela et al. [41] determined the arsenic speciation in carrots using ASE at 1008C and water as a solvent. It was found that the ASE extraction procedure did not cause any redox or interconversion reactions among the arsenic species. Lindemann et al. [42] examined the stability of As(III) and As(V) (in the presence of Se, Sb, and Te species) in water and urine (NIST 2670 normal level) as well as in extracts of ﬁsh (NRCC DORM-2) and soil (NIST 2710) under various storage conditions. Surprisingly, it was found that storage of an aqueous solution at 2 208C for 30 days resulted in a substantial loss of As(III), whereas at 3 and 208C quantitative recovery could be obtained. The recovery of As(V) was not inﬂuenced by the storage temperature. In a urine matrix spiked with As(V) as well as As(III) both were quantitatively recovered after 5 days of storage at 38C. Spiked ﬁsh extracts as well as spiked soil samples could be stored for 3 days without signiﬁcant change of the spiked As(III) and As(V) concentrations. When aqueous arsenic standard solutions were added to the solid ﬁsh and soil material before the extraction procedure, quantitative recovery was obtained for the ﬁsh matrix, whereas only 70% of the spiked As(V) and 40% of the spiked As(III) were recovered from the soil. As no additional arsenic species were detected in the chromatograms, an incomplete extraction was blamed for the ﬁndings, although no arsenic determinations of the extraction residue had been performed. The authors concluded that extraction of species from solid material and species transformation is still the Achilles heel in speciation analysis. In an earlier study, Palacios et al. [43] systematically investigated the stability of several arsenic species in deionised water and urine. The authors did not consider As(III) in their study because “…it is oxidised in almost all conditions to As(V)”. In deionised water and urine samples, As(V) was stable for 67 days at temperatures of 2 20, 48C, and room temperature. In a preceding study, these authors found immediate conversion of As(III) to As(V) when spiked to urine. These ﬁndings are somewhat surprising, as As(III) then should never be detectable in urine samples. Montperrus et al. [44] found that microwave assisted extraction minimises the risk of interconversion of the inorganic arsenic species because of the shorter time needed for complete extraction as compared to conventional shaking. From the above, one can clearly see that no general recipe can be given to preserve the species information. In the case where the As(III) concentration has to be determined in addition to As(V), the stability must be evaluated in the sample matrix to be investigated. For many studies, it is not really very important to distinguish between these two forms as the toxicity of both compounds is very similar and the sum of both is a good measure of exposure. 31.3.2 Methylarsonous acid and dimethylarsinous acid With the more or less clear discovery of the key metabolites of MA(III) and DMA(III) in human urine, interest in studying metabolism and health effects of

1035

W. Goessler and D. Kuehnelt

these two arsenic compounds is increasing. In recent work by Gong et al. [45], the stability of these two compounds in deionised water and urine was systematically investigated. Storage of MA(III) in deionised water at 220 and 48C resulted in a slight oxidation of about 10% to MA after 4 months. At room temperature (258C), 15% of MA(III) were already oxidised after 3 days and 80% after ,20 days. The situation was even worse, when the stability of MA(III) in the urine matrix was investigated. At room temperature, complete conversion of MA(III) to MA occurred within 1 week. At 2 20 and 48C an oxidation of , 30% to MA was evident. After 30 days at 48C, about 98% was converted to MA. The stability of MA(III) was better at 2 208C (time days/% converted: 30/60, 60/80, 110/90). DMA(III) was found to be very unstable in water as well as in the urine matrix. DMA(III) was quantitatively oxidised to DMA in the water matrix after 10 days at 258C, after 13 days at 48C, and after 15 days at 2 208C. In urine, DMA(III) was quantitatively converted to DMA after 90 min at 258C. It took 12 and 17 h to convert DMA(III) quantitatively to DMA at 4 and 2 208C, respectively. This work was performed at high concentrations (100 mg As/l) usually not detectable in urine samples of exposed people. One can easily imagine that at a natural concentration level the situation might be even worse. Thus far, this is the only systematic investigation of the stability of the trivalent arsenicals. Storage at lower temperature, e.g., shock freezing in liquid nitrogen, or storage in an inert atmosphere, e.g., N2, or even freeze-drying might improve the stability, especially that of DMA(III). 31.3.3 Methylarsonic acid and dimethylarsinic acid In contrast to the trivalent forms, MA and DMA are stable arsenic compounds. Storage of stock standard solutions (1 g As/l) over 1 year in polyethylene containers at 48C did not show any changes of these species. A change of the species, even at low concentrations as mentioned for As(III) and As(V), was not detected. Palacios et al. [43] found MA to be stable in deionised water for 67 days at room temperature, 48C, and 2 208C when stored together with As(V), DMA, AB, and AC. In the same study, an increase of the DMA concentration was observed at 48C and room temperature. As a possible explanation, a conversion of AB and AC to DMA was suggested. In a urine matrix, MA as well as DMA were stable at 48C and room temperature for 67 days. Lindemann et al. [42] found good stability of MA and DMA in extracts of four different matrices (water, urine, ﬁsh, and soil) over 30 days at 220, þ 3, and 208C. Schmidt et al. [40] investigated the thermal stability (60 –1808C) of MA and DMA using pressurised solvent extraction with water as the extractant. The recoveries were minus 10% for MA and minus 2 20% for DMA at 1808C as compared to the values obtained at 608C. Detailed information about whether the compounds have been lost or decomposed was not given. Heating MA and DMA in nitric

1036

Sample preparation for arsenic speciation

acid at various temperatures revealed that MA is stable up to 1408C and DMA up to 2008C [46]. MA and DMA are relatively stable arsenic compounds. No special requirements are necessary for extracting them as long as the conditions are not too harsh. 31.3.4 Arsenobetaine, arsenocholine, trimethylarsine oxide, and the tetramethylarsonium ion AB, the dominant arsenic compound in marine animals, is very stable. It can be stored at 48C in a refrigerator for years unless microbiological transformation is excluded. Khokiattiwong et al. [47] found that AB is demethylated to dimethylarsionylacetate by microorganisms. Mu¨rer et al. [48] found that AB as well as AC decompose in the presence of hydrogen peroxide when exposed to daylight, AC being the more unstable arsenic compound. Upon heating in nitric acid, AB and AC are converted quantitatively to trimethylarsine oxide (TMAO), as shown by Goessler and Pavkov [46] which then needs about 1 h heating at 3008C for complete conversion to As(V). Similar results were found by Slejkovec et al. [49] and Wasilewska et al. [50]. When roasting a piece of lobster, AB is converted to TETRA, as shown by Hanaoka et al. [51]. Palacios et al. [43] reported that AC disappears from an aqueous solution after 3 days at room temperature (oxidised to AB). In the same work, AB was found to be converted to DMA after 67 days. These ﬁndings are not in agreement with our experience. As well, Lindemann et al. [42] found AB to be stable for 30 days when spiked to four different matrices (water, urine, ﬁsh, and soil) at 2 20, 3, and 208C. AC is stable up to 1508C when pressurised solvent extraction with water was employed [40]. At 1808C, only 75% recovery was obtained. Kirby and Maher [52] reported that AB, AC, and TETRA were not changed when water– methanol mixtures and microwave assisted heating were used for extracting these arsenicals from ﬁsh matrices. AB, TMAO, and TETRA are stable arsenic compounds when they are not exposed to light in an oxidative environment. AC is rather labile and has to be treated with care. Quaternary arsonium compounds, such as AB and AC, decompose to TMAO when heated in alkaline solutions (2 M NaOH) at , 958C for 3 h [53]. 31.3.5 Arsenosugars Arsenosugars, the dominant arsenic compounds in marine algae, have not been studied too much from their chemical point of view due to a lack of synthetic standards. Aqueous solutions of the four major arsenosugars (Fig. 31.2) tend to be relatively stable, with the exception of arsenosugars 2 and 4, which might be converted to arsenosugar 1 [54]. In early work by Edmonds and Francesconi [12], it was reported that arsenosugars decompose to DMA upon heating with

1037

W. Goessler and D. Kuehnelt

hydrochloric acid. Recently, Gamble et al. [55] found that arsenosugars do not decompose to DMA, but form the base arsenosugar without aglycone when treated in 78 mM hydrochloric acid or nitric acid. This base arsenosugar (5-deoxy-5-dimethylarsinoyl-D -ribose) was already prepared by Edmonds et al. [56] upon treatment of the analogous methylriboside with HCl. Arsenosugars are certainly more labile than AB. The use of harsher (acidic) conditions for higher extraction yields must be avoided in order to determine which arsenosugars are really present in a sample. Because risk assessment should be based on the “real” arsenic compounds present, further research for better characterisation of the arsenosugars is needed.

31.4

EXTRACTION OF ARSENIC COMPOUNDS FROM ENVIRONMENTAL SAMPLES

Thus far, several articles have been published with the goal of ﬁnding the “ideal” method to make the arsenic compounds present in a solid sample available for analysis. Usually, the following points are considered: † † † †

The The The The

extractant extractant extractant extractant

should completely penetrate the sample. should be a solute for the all the arsenic compounds. must not change the speciation. must be compatible with the analytical method chosen.

In the ﬁeld of arsenic speciation in biological samples, water and water/methanol mixtures are certainly the most common extractants. For extraction of nonpolar arsenicals, acetone or chloroform is sometimes employed. Easy handling, good sample penetration, high solubility for the common arsenicals, reasonable stability of the arsenic compounds, and excellent compatibility with the analytical methods are the reasons for using these extractants. In the case of methanol interfering with the chromatographic separation, it can be easily evaporated. To improve extraction yields, mechanical agitation, vortexing, and sonication are often applied. Pressurised liquid extraction (ASE) at elevated temperatures as well as microwave assisted heating were recently shown to increase extraction efﬁciencies. Due to the high arsenic concentrations in marine biota, extraction methods are often compared for ﬁsh or algal reference materials. DORM-1 and DORM-2 (defatted dogﬁsh muscle) from the National Research Council of Canada (NRCC) are well characterised for their arsenic compounds and, therefore, serve as a “model matrix”. Moreover, the compounds present in this CRM are easily extracted, most probably due to the defatted matrix. Extraction with water/methanol (1 þ 9) at a ratio 1:99 (solid material to extractant) and shaking for 14 h extracted 93% of the arsenic present in DORM-2 [57]. A slightly higher extraction yield of 97% was obtained by Mattusch and

1038

Sample preparation for arsenic speciation

Wennrich [58] after threefold extraction of DORM-2 with water/methanol (1 þ 1) and sonication for 30 min. Londesborough et al. [59] released only 82% of the arsenic present in DORM-2 after shaking for 2 h with water. Microwave heating to 70–758C with methanol/water (1 þ 1) for 5 min quantitatively released arsenic (103 ^ 2%) from DORM-2 [52]. McKiernan et al. [60] compared pressurised liquid extraction with sonication and recovered 94 and 92% using acetone and water/methanol (1 þ 1), respectively. Recently, Kuehnelt et al. [61] used 1.5 M phosphoric acid (commonly used as extractant for soil samples), methanol/water (9 þ 1), or water for extracting the arsenicals from DORM-2 at room temperature by shaking for 14 h. The recoveries obtained were 94, 92, and 87%, respectively. The results indicate that methanol improves the extraction yields of arsenic compounds, especially AB, from marine animals. Although there is no clear proof for the role of AB in marine animals, it has been suggested that it is utilised in a manner similar to gylcine betaine, an important osmolyte, as speculated by Gailer et al. [62]. The usually high extraction yields for AB imply that it is not bound to any cell compartment and support the speculations that it functions as an osmolyte and is therefore free in the cytosol. When liver tissues of marine mammals were extracted with methanol/water (9 þ 1) and mechanical agitation for 14 h, the extraction yields ranged from 44 to 77% [63]. The fact that a procedure extracting practically 100% from a muscle tissue is only capable of extracting up to 77% from a liver tissue clearly shows that no common extraction procedure is available thus far. Besides extraction of arsenicals from marine animals, much effort has been devoted to the extraction of the arsenic compounds present in marine algae. Kuehnelt et al. [61] used 1.5 M phosphoric acid, methanol/water (9 þ 1), or water for extracting the arsenicals from the brown alga Hijiki fuziforme at room temperature by shaking for 14 h. The methanol/water mixture extracted only 33% of the total arsenic, whereas the same procedure extracted 92% from DORM-2. With pure water, 62% and with 1.5 M phosphoric acid, 76% were extractable. The concentrations of the arsenic compounds present in the different extracts were about the same with the exception of arsenosugar 1, which was best extracted with phosphoric acid, and of As(III) of which only ,10% were extractable with methanol/water. Yoshinaga et al. [64] also found water to be more effective for removing As(V) from algal material. When investigating mushrooms with high concentrations of inorganic arsenic, Byrne et al. [22] obtained better extraction yields with water as compared to water/methanol mixtures. Extraction yields above 85% were obtained by McSheehy et al. [65] when seaweed samples were sonicated for 3 h with water/methanol (1 þ 1) and then sonicated with water/methanol (9 þ 1) for 3 h and 20 min. Only a few systematic investigations have been conducted for the extraction of arsenic species from terrestrial plants. Vela et al. [41] successfully employed pressurised liquid extraction to release arsenic from freeze-dried carrots. For most of the analysed samples, extraction yields above 80% were obtained. Moreover, it is

1039

W. Goessler and D. Kuehnelt

important to mention that the authors did not observe any species transformation. Helgesen and Larsen [66] used methanol/water (1 þ 9) and microwave assisted heating for 8 min at 708C for extracting arsenic compounds from carrots. Extraction yields obtained ranged from 46 to 69%. In a recent publication by Heitkemper et al. [67] quantitative extraction of arsenic compounds from SRM 1568a rice ﬂour (NIST) was reported using pressurised solvent extraction at room temperature or 1008C with water or water/methanol mixtures, but only 36% was extracted from a natural rice sample. The extraction efﬁciency for this rice sample was improved to 92%, only when hydrolysing the sample with 2 M triﬂuoroacetic acid at 1008C for 6 h. Arsenic compounds present in apples were extracted with water/ methanol (1 þ 1) and 6 h sonication. The extraction yield was ,70% [68]. A further improvement in the extraction yield to 80–90% was obtained when an enzymatic treatment with amylase was performed prior to the solvent extraction with water/methanol. The amylase treatment successfully broke up the starch structure of the apple matrix. Extraction of arsenic and its compounds from soil and sediments usually gives low yields. As extractants, mineral acids are commonly employed because with water or water/methanol mixtures, less than 10% are extractable. Demesmay and Olle [69] used hydrochloric acid/nitric acid mixtures (with magnetic stirring or microwave solubilisation) for the extraction of arsenic compounds from a lake sediment sample. Quantitative extraction yields were obtained but As(III) was quantitatively oxidised to As(V). MA and DMA were stable under these conditions. The oxidation of As(III) was avoided when the arsenic compounds were extracted with 0.3 M orthophosphoric acid and magnetic stirring. With microwave assisted solubilisation, about 10–30% of the As(III) was oxidised to As(V). Quantitative recovery of the arsenic compounds was also obtained with 0.3 M ammonium oxalate at pH 3. Vergara Gallardo et al. [70] extracted a soil (NIST SRM 2709, NIST), a river sediment (CRM 320, BCR), and a sewage sludge (CRM 007-040, RT) with orthophosphoric acid at three concentrations (0.3, 1, 3 M) using microwave solubilisation. Quantitative extraction yields were obtained for the sediment and the sludge sample, irrespective of the orthophosphoric acid concentration used, whereas the yield for the soil sample did not exceed 62%. A partial conversion of As(III) to As(V) was observed with increasing orthophosphoric acid concentration for the sediment sample, whereas the opposite was observed for the sludge sample. Moreover, it was reported for the sludge sample that 90% of the As(III) was converted to As(V) after 4 h when the 3 M extract was stored. Neutralisation or dilution of the extract ensures stability of As(III) for several hours. For the soil sample, an improvement in the extraction yield was obtained with increasing orthophosphoric acid concentration. A 0.2 M ammonium oxalate solution was reported by Montperrus et al. [44] to extract .80% of the total arsenic present in this soil sample. In classical extraction schemes, ammonium oxalate was used to dissolve crystalline iron oxides.

1040

Sample preparation for arsenic speciation

31.5

CONCLUSIONS

Frequently scientiﬁc publications imply that speciation analysis is nowadays already routinely performed. This might be true for the determination step (chromatographic separation and detection) of the analysis, although there are only a few round robin exercises that have generated good agreement between different laboratories. The sample storage and sample preparation step is often completely neglected in these publications. It is well known that the sampling process, together with sample storage and preparation accounts for more than 80% of the total error of an analytical result. In our opinion, there much research remains to be done to improve the extraction efﬁciencies without changing the compounds. The use of microwave assisted extraction at low power settings, or the employment of pressurised liquid extraction for releasing arsenic compounds from a solid matrix, can certainly improve the situation with respect to extraction efﬁciencies, less time consumption, and species preservation. Nevertheless, the extraction procedures have to be optimised with respect to the sample matrix and the arsenic compounds present in the sample (unfortunately there is no ultimate extraction procedure available thus far). For the routine determination of arsenic compounds in solid samples of any matrix, much research has to be done to obtain accurate and comparable results, necessary for legislators to set appropriate limits for human health and to save our environment. The “lipid soluble” arsenic (the fraction of arsenic that is not extracted with polar solvents), in particular, has been treated as a stepchild until now and will need a lot more attention in the future.

REFERENCES 1 2

3 4 5 6 7

8

R.M. Allesch, Arsenik. Ferd. Kleinmayr, Klagenfurt, 1959, pp. 239 –250. J.M. Azcue and J.O. Nriagu, Arsenic: historical perspectives. In: J.O. Nriagu (Ed.), Arsenic in the Environment. Part I: Cycling and Characterisation. Wiley, New York, 1994, pp. 1–15. J.O. Nriagu, Arsenic poisoning through the ages. In: W.T. Frankenberger (Ed.), Environmental Chemistry of Arsenic. Marcel Dekker, New York, 2002, pp. 1 –26. Arsenic in drinking water 2001 update. National Academy Press, Washington, DC, 2001. D. Chakraborti, M.M. Rahman, K. Paul, U.K. Chowdhury, M.K. Sengupta, D. Lodh, C.R. Chanda, K.C. Saha and S.C. Mukherjee, Talanta, 58 (2002) 3. Handbook of Chemistry and Physics 67th edn. 1986– 1987. CRC Press, Boca Raton, 1986. A.J. Jones, The arsenic content of some of the marine algae. In: C.H. Hamshire (Ed.), Yearbook of Pharmacy, Transactions of the British Pharmaceutical Conference. Churchill, London, 1922, pp. 388–395. A.C. Chapman, Analyst, 51 (1926) 548.

1041

W. Goessler and D. Kuehnelt 9 10 11 12 13

14 15 16 17 18 19

20

21 22 23 24 25 26 27

28 29

30

31 32 33 34

1042

H. Yamauchi, K. Takahashi, M. Mashiko, J. Saitoh and Y. Yamamura, Appl. Organomet. Chem., 6 (1992) 383. S.S. Tao and P.M. Bolger, Food Addit. Contam., 16 (1999) 465. J.S. Edmonds, K.A. Francesconi, J.R. Canon, C.L. Raston, B.W. Skelton and A.H. White, Tetrahedron Lett., 18 (1977) 1543. J.S. Edmonds and K.A. Francesconi, Nature, 289 (1981) 602. K.A. Francesconi and D. Kuehnelt, Arsenic compounds in the environment. In: W.T. Frankenberger (Ed.), Environmental Chemistry of Arsenic. Marcel Dekker, New York, 2002, pp. 51–94. T. Kaise, S. Watanabe and K. Itoh, Chemosphere, 14 (1985) 1327. K.A. Francesconi, R. Tanggaar, C.J. McKenzie and W. Goessler, Clin. Chem., 48 (2002) 92. K.A. Francesconi and J.S. Edmonds, Oceanogr. Mar. Biol. Annu. Rev., 31 (1993) 111. K.A. Francesconi and J.S. Edmonds, Adv. Inorg. Chem., 44 (1997) 147. P. O’Neill, Arsenic. In: B.J. Alloway (Ed.), Heavy Metals in Soils, 2nd edn. Blackie Academic & Professional, London, 1995, pp. 105 –121 (and references therein). D.C. Chilvers and P.J. Peterson, Global cycling of arsenic. In: T.C. Hutchinson and K.M. Meema (Eds.), Lead, Mercury, Cadmium and Arsenic in the Environment. Wiley, London, 1987, pp. 279– 301. W.T. Frankenberger and M. Arshad, Volatilization of arsenic. In: W.T. Frankenberger (Ed.), Environmental Chemistry of Arsenic. Marcel Dekker, New York, 2002, pp. 360–380, (and references therein). W.R. Cullen and K.J. Reimer, Chem. Rev., 89 (1989) 713. A.R. Byrne, Z. Slejkovec, T. Stijve, L. Fay, W. Goessler, J. Gailer and K.J. Irgolic, Appl. Organomet. Chem., 9 (1995) 305. D. Kuehnelt, W. Goessler and K.J. Irgolic, Appl. Organomet. Chem., 11 (1997) 289. D. Kuehnelt, W. Goessler and K.J. Irgolic, Appl. Organomet. Chem., 11 (1997) 459. A. Geiszinger, W. Goessler, D. Kuehnelt, K.A. Francesconi and W. Kosmus, Environ. Sci. Technol., 32 (1998) 2238. Z. Slejkovec, A.R. Byrne, T. Stijve, W. Goessler and K.J. Irgolic, Appl. Organomet. Chem., 11 (1997) 673. E.H. Larsen and T. Berg, Trace element speciation and international food legislation—a codex alimentarius position paper on arsenic as contaminant. In: L. Ebdon, L. Pitts, R. Cornelis, H. Crews, O.F.X. Donard, P. Quevauviller (Eds.), Trace element speciation for environment, food and health. Cambridge: RSC, 2001, pp. 251–260 (and references therein). Environmental Health Criteria 224. Arsenic and Arsenic Compounds. 2nd edn. World Health Organisation, Geneva, 2001. M. Vahter, Variation in human metabolism of arsenic. In: W.R. Chappell, C.O. Abernathy, R.L. Calderon (Eds.), Arsenic exposure and health effects. Amsterdam: Elsevier, 1999, pp. 267–279 (and references therein). V.H. Aposhian, E.S. Gurzau, X.C. Le, A. Gurzau, S.M. Healy, X. Lu, M. Ma, L. Yip, R.A. Zakharyan, R.M. Maiorino, R.C. Dart, M.G. Tirus, D. Gonzalez-Ramirez, D.L. Morgan, D. Avram and M.M. Aposhian, Chem. Res. Toxicol., 13 (2000) 693. X.C. Le, X. Lu, W.R. Cullen, V. Aposhian and B. Zheng, Anal. Chem., 72 (2000) 5172. M.J. Mass, A. Tennant, R.C. Roop, W.R. Cullen, M. Styblo and D.J. Thomas, Chem. Res. Toxicol., 14 (2001) 305. Y. Shibata, J. Yoshinaga and M. Morita, Appl. Organomet. Chem., 8 (1994) 249. X. Zhang, R. Cornelis, J. De Kimpe, L. Mees, V. Vanderbiesen, A. De Cubber and R. Vanholder, Clin. Chem., 42 (1996) 249.

Sample preparation for arsenic speciation 35 36

37

38

39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59

D.M. Templeton, F. Ariese, R. Cornelis, L.-G. Danielsson, H. Muntau, H.P. Van Leeuwen and R. Lobinsky, Pure Appl. Chem., 72 (2000) 1453. C.J. Langdon, A.A. Meharg, J. Feldmann, T. Balgar, J. Charnock, M. Farquhar, T.G. Piearce, K.T. Semple and J. Cotter-Howells, J. Environ. Monit., 4 (2002) 603. W.P. Inskeep, T.R. McDermott and S. Fendorf, Arsenic (V)/(III) cycling in soils and natural waters: chemical and microbial processes. In: W.T. Frankenberger (Ed.), Environmental Chemistry of Arsenic. Marcel Dekker, New York, 2002, pp. 183–215 (and references therein). D. Kuehnelt, W. Goessler and K.J. Irgolic, The oxidation of arsenite in aqueous solutions. In: C.O. Abernathy, R.L. Calderon and W.R. Chappell (Eds.), Arsenic: Exposure and Health Effects. Chapman & Hall, London, 1997, pp. 45– 54. A.J. Bednar, J.R. Garbarino, J.F. Ranville and T.R. Wildeman, Environ. Sci. Technol., 36 (2002) 2213. A.-C. Schmidt, W. Reisser, J. Mattusch, P. Popp and R. Wennrich, J. Chromatogr. A, 889 (2000) 83. N.P. Vela, D.T. Heitkemper and K.R. Stewart, Analyst, 126 (2001) 1011. T. Lindemann, A. Prange, W. Dannecker and B. Neidhard, Fresenius J. Anal. Chem., 368 (2000) 214. M.A. Palacios, M. Gomez, C. Camara and M.A. Lopez, Anal. Chim. Acta, 340 (1997) 209. M. Montperrus, Y. Bohari, M. Bueno, A. Astruc and M. Astruc, Appl. Organomet. Chem., 16 (2002) 347. Z. Gong, Y. Lu, W.R. Cullen and X.-C. Le, J. Anal. At. Spectrom., 16 (2001) 1409. W. Goessler and M. Pavkov, Analyst, 128 (2003), 796. S. Khokiattiwong, W. Goessler, S.N.R. Cox and K.A. Francesconi, Appl. Organomet. Chem., 15 (2001) 481. A.J.L. Mu¨rer, A. Abildtrup, O.M. Poulsen and J.M. Christensen, Analyst, 117 (1992) 677. Z. Slejkovec, J.T. van Elteren and U.D. Woroniecka, Anal. Chim. Acta, 443 (2001) 277. M. Wasilewska, W. Goessler, M. Zischka, B. Maichin and G. Knapp, J. Anal. At. Spectrom., 17 (2002) 1121. K. Hanaoka, W. Goessler, H. Ohnu, K.J. Irgolic and T. Kaise, Appl. Organomet. Chem., 15 (2001) 61. J. Kirby and W. Maher, J. Anal. At. Spectrom., 17 (2002) 838. S. Maeda, H. Wada, K. Kumeda, M. Onue, A. Okhi, S. Higashi and T. Takeshita, Appl. Organomet. Chem., 1 (1987) 456. A.D. Madsen, W. Goessler, S.N. Pedersen and K.A. Francesconi, J. Anal. At. Spectrom., 15 (2000) 657. B. Gamble, P.A. Gallagher, J.A. Shoemaker, X. Wei, C.A. Schwegel and J.T. Creed, Analyst, 127 (2002) 781. J.S. Edmonds, Y. Shibata, K.A. Francesconi, J. Yoshinaga and M. Morita, Sci. Total Environ., 12 (1992) 321. W. Goessler, D. Kuehnelt, C. Schlagenhaufen, Z. Slekovec and K.J. Irgolic, J. Anal. At. Spectrom., 13 (1998) 183. J. Mattusch and R. Wennrich, Anal. Chem., 70 (1998) 3649. S. Londesborough, J. Mattusch and R. Wennrich, Fresenius J. Anal. Chem., 363 (1999) 577.

1043

W. Goessler and D. Kuehnelt 60 61 62 63 64 65 66 67 68 69 70

1044

J.W. McKiernan, J.T. Creed, C.A. Brockhoff, J.A. Caruso and J.M. Lorenzana, J. Anal. At. Spectrom., 14 (1999) 607. D. Kuehnelt, K.J. Irgolic and W. Goessler, Appl. Organomet. Chem., 15 (2001) 445. J. Gailer, K.A. Francesconi, J.S. Edmonds and K.J. Irgolic, Appl. Organomet. Chem., 9 (1995) 341. W. Goessler, A. Rudorfer, E.A. Mackey, P.R. Becker and K.J. Irgolic, Appl. Organomet. Chem., 12 (1998) 491. J. Yoshinaga, Y. Shibata, T. Horiguchi and M. Morita, Accred. Qual. Assur., 2 (1997) 154. S. McSheehy, M. Marcinek, H. Chassaigne and J. Szpunar, Anal. Chim. Acta, 410 (2000) 71. H. Helgesen and E.H. Larsen, Analyst, 123 (1998) 791. D.T. Heitkemper, N.P. Vela, K.R. Stewart and C. Westphal, J. Anal. At. Spectrom., 16 (2001) 299. J.A. Caruso, D.T. Heitkemper and C. B’Hymer, Analyst, 126 (2001) 136. C. Demesmay and M. Olle, Fresenius J. Anal. Chem., 357 (1997) 116. M. Vergara Gallardo, Y. Bohari, A. Astruc, M. Potin-Gautier and M. Astruc, Anal. Chim. Acta, 441 (2001) 257.

Chapter 32

Sample preparation for speciation of selenium Claudia Ponce de Leon, Anne P. Vonderheide and Joseph A. Caruso

32.1

WHY SELENIUM SPECIATION?

Selenium is an essential element, yet it has a rather narrow window between essentiality and toxicity. Selenium can exist in several oxidation states (þ 4 or þ6), as elemental selenium [Se(0)] and in a variety of organic compounds (2 2), including naturally occurring amino acids, such as selenomethionine, selenocystine, methylselenocysteine and selenocysteine [1]. Research has shown that its bioavailability, as well as how it is stored in the body, depends on its chemical form [2]. Currently, the two primary areas in which speciation information is critical include the elucidation of selenium biological mechanisms and environmental risk assessment as a result of the differing selenium toxicities of various species found in the environment [3]. For these reasons, determination of the different forms of selenium has become important [4]. Speciation of different forms of selenium commonly employs a separation technique coupled to a selective and sensitive detector. High performance liquid chromatography (HPLC), gas chromatography (GC) and capillary electrophoresis (CE) are ﬂexible methods, which have proven to separate a wide variety of selenium species. One of the most common detectors to which these separation techniques are interfaced is the inductively coupled plasma mass spectrometer (ICP-MS); this detector has shown to be both selective and sensitive in the analysis of selenium-containing compounds [5]. 32.2

GENERAL SAMPLE PREPARATION

Sample preparation for speciation studies is a complex task as it is imperative to preserve the nature of the species. Digestion methods commonly employed for total element determinations are not adequate as they usually change the original species. Therefore, milder procedures are required and, in some cases, Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

1045

C.P. de Leon et al.

concentration techniques may be necessary due to the low levels of some species. An additional factor that must be taken into account in the design of a sample preparation method for speciation concerns the instrumental set-up chosen for analysis. Some instruments are intolerant to particular matrices or extraction media. For example, although the ICP-MS is a commonly used detector for elemental speciation studies [6], extraction phases high in organic solvent may destabilize or even extinguish the plasma. In general, qualitative separation schemes need to be coordinated with selective detection. Another issue that must be considered relates to lack of commercially available selenium standards. For those that are available, identiﬁcation based upon retention times matched to the known standards is common protocol. However, unidentiﬁed peaks must be elucidated by other means. This is often accomplished by mass spectrometry coupled with ionization sources that are gentler than the inductively coupled plasma. Electrospray mass spectrometry (ES-MS) is often used in this capacity and it must be recognized that extracts need to be prepared with special precautions when analyzed with this type of instrumentation [7]. Generally, sample preparation techniques for speciation analyses consist of several steps [8]. The necessary steps depend on the matrix to be analyzed and the analytes to be determined. For example, in the speciﬁc case of selenium, it may be wise to employ enzymatic treatments for biological matrices should the analytes of interest include selenoamino acids, because these may be incorporated into the protein structure of the matrix. Conversely, inorganic selenium may be best determined following aqueous extraction, and volatile organic selenium compounds might be easily sampled from the headspace of a particular material. In the following sections, various matrices are considered for the purpose of selenium speciation analysis. Matrix-speciﬁc extraction techniques are discussed with the goal of extraction efﬁciency and species preservation. 32.3

MAMMALS

32.3.1 Body ﬂuid analysis Considerable interest has emerged for the analysis of selenium in body ﬂuids since the discovery of its potential therapeutic properties [9,10]. These have triggered the study of the bioselenium compounds in the body and, as a result, a series of selenium proteins with still unknown functions has been discovered. The enzyme, glutathione peroxidase, is probably the most well known selenium biocompound and it is involved in antioxidative processes. The most common body ﬂuids analyzed for selenium include blood and its fractions [plasma (serum) and red cells (erythrocytes)], breast milk and urine. Analysis of biocompounds is a challenge as difﬁculties have been experienced

1046

Sample preparation for speciation of selenium

with sample preservation and preparation techniques adequate to preserve the original species. Detailed descriptions of these procedures can be found in the literature [11–13]. Plasma and urine have the advantage of possessing soluble compounds; red cells have to be lysed to liberate Se species. When undertaking speciation in clinical samples, a distinction between low molecular mass (LMM) and high molecular mass (HMM) should be made. The separation of the two can be obtained through centrifugal ultraﬁltration. When only the LMM is to be studied, ultraﬁltration can be done using a semi-permeable membrane. This membrane can be purchased with different cut-off molecular weight ﬁlters (e.g. 25, 50 kDa). The ultraﬁltrate is collected and subsequently analyzed by the method of choice. Desalting can be another necessary step in the analysis of clinical samples. It is usually used when the ionic strength of the solution does not comply with the conditions of the chromatographic technique. For instance, urine samples have a variable electrolyte composition; supernatant tissue homogenates have a very high saline content and therefore have to be desalted. Commercial desalting columns, such as HR 10/10 (FDC) from Pharmacia, can be used for this purpose. 32.3.1.1 LMM analysis Although urine is a complex matrix, it possesses the advantage that all its normal constituents are water-soluble and this permits direct chromatography of the sample. However, it contains a high salt concentration (approximately 1% NaCl). Furthermore, the complex matrix may overload the column and produce peak broadening or split peaks. Gonzalez-La Fuente et al. [14] analyzed urine for selenium species with HPLC –ICP-MS by diluting 1:1 with ultrapure Milli-Q water and ﬁltering the sample through a Millipore 20 mm membrane; 50 ml was injected into the chromatographic system. All samples were kept in the dark at 48C before analysis. Quijano et al. [15] processed human urine in a vacuum manifold system by passage through Bond-Elut C18 cartridges previously conditioned with 5 ml of methanol followed by 5 ml of Milli-Q water. The cartridge was washed with 5 ml of 3.5 mM phosphate buffer at pH 6.0. The eluate was diluted to 10 ml with Milli-Q water. This process improved the stability of urine samples by removing the organic matrix and thus avoiding bacterial growth. A similar process was used by Gomez et al. [16]. To avoid bacterial growth, other authors have suggested the use of toluene, formaldehyde, hydrochloric acid and nitric acid [17] but the risk of species transformation should be noted. A novel approach for selenium analysis of urine has been taken by Gammelgard et al. [18] in which they preconcentrated the selenium compounds with a crown ether extraction. Four milliliters of urine and 100 mg of picric acid were mixed with 4 ml 50 mM benzo-15-crown-5-ether solution for 3 min.

1047

C.P. de Leon et al.

The mixture was centrifuged for 5 min. The extraction was repeated on the supernatant four times. Anion-exchange solid-phase extractions of samples were performed on Bond-Elut SAX 500 mg cartridges. These were conditioned with 3 ml methanol, followed by 3 ml of 10 mM ammonium formate at pH 3. One milliliter of extracted sample was applied followed by 1 ml of 10 mM ammonium formate, pH 3. The eluate from the last 2 ml was collected and an aliquot was injected without dilution; cation and anion-exchange chromatography were utilized. This method signiﬁcantly reduced sodium and potassium interferences. 32.3.1.2 HMM analysis In principle, no preliminary ultraﬁltration is needed for the analysis of HMM, but rather a separation by a suitable chromatography, such as fast liquid chromatography. Koyama et al. [19] analyzed Se-proteins in human and mouse plasma. For the separation of HMM, the samples were centrifuged at 1500 g for 10 min and the supernatant was stored at 2808C prior to analysis. The separation was carried out with a prepacked heparin afﬁnity column and a sizeexclusion column. The mobile phases were: (A) 0.02 M sodium phosphate buffer pH 7.5 in 100 ppm Na2-EDTA and (B) 500 units ml21 in mobile phase A, respectively. Similar methods have been used by Sasakura and Susuki [20] for the analysis of selenoprotein P in the blood stream and its interaction with transition metals. The selenoprotein P in serum was adsorbed on heparinsepharose resin and then separated from extracellular glutathionine peroxidase and other plasma proteins. The separation of proteins with different trace metals was done in a GS 520 column (7.5 £ 500 mm). Susuki et al. [21] worked with Se-containing biological constituents in rats fed with different selenium diets. A size-exclusion column with an exclusion limit of Mr 300,000 and a mobile phase of 50 mM Tris –HCl (pH 7.4) were used. As mentioned earlier, the analysis of erythrocytes requires lysing. Bergdahl et al. [22] and Gercken and Barnes [23] used three freeze-thaw cycles to lyse the cells followed by a 10-fold dilution with buffer and centrifugation to remove fragments of membranes. An alternative approach is suggested by Cornelis et al. [11], where lysis is obtained when the packed cells are mixed with cold toluene and ice-cold water. The lysate is centrifuged at 15; 000g at 48C and then ﬁltered through a 0.45 mm ﬁlter. Analysis of selenium compounds on human erythrocytes lysate was done by Laborda et al. [24]. Blood samples with EDTA as anticoagulant were centrifuged at 200g for 10 min at room temperature and the supernatant was discarded. The erythrocytes were washed twice with 0.15 M NaCl. The ﬁnal pellet of erythrocytes was treated with another volume of deionized water and frozen. The frozen preparation was centrifuged at 10; 000g for 10 min before injection onto the column. The separation was obtained using a size-exclusion column and Tris –HCl buffer ðpH ¼ 7:0Þ with 0.1 M of NH4NO3 for ionic strength. Detection was made online with graphite furnace atomic absorption.

1048

Sample preparation for speciation of selenium

Breast milk is a pool of nutrients in which casein is the major protein. Casein is a 20 kDa protein that exists in micellar form. The micelles are spherical polyspread colloidal aggregates with molecular weights exceeding 1000 kDa. Infant formulas mimic these characteristics of breast milk. The analysis of milk requires the elimination of lipids and this can be done by centrifugation at 3000 rpm for 30 min at 48C; precipitation of casein can be performed with 1 M acetate and the resulting whey used for analysis. If further separation is needed, dialysis and SEC separation [25] can be executed. Michalke [26] fractionated defatted human milk in a metal-free SEC (with Toyo Pearl TSK HW 40) utilizing double distilled water at a ﬂow rate of 3 ml min21 at 208C. The separation was monitored at 232 nm with UV detection. Fractions were collected every 5 min with a fraction collector. The fractions were frozen at 2208C and subsequently freeze dried. The dry powder was re-suspended with doubly distilled water (500 ml) and used for the determination of selenium and CE analysis. Bermejo et al. [27] separated selenium proteins in infant formula in an SEC column (TSK gel G2000 glass 30 cm £ 8 mm, Japan) utilizing a mobile phase composition of 0.2 M NH4NO3 adjusted to pH 6.7 with NH4OH. An alternative to SEC is ultraﬁltration using 10 kDa cut-off ﬁlters [28]. 32.3.2 Tissue sample analysis Solubilization of the species from tissue samples begins with subjecting the sample to a very intense ultrasonic homogenization in an isotonic phosphate buffer (pH 7.3) saline solution. This can be done by applying short bursts (approximately 10 s) in an ice-water bath. The homogenate is then centrifuged at 48C at 15; 000g for 1 h. The supernatant is removed and the precipitate is washed and centrifuged 3 times with equal volumes of phosphate buffer, adding the washings to the supernatant. This ensures the solubilization of the compounds trapped inside the precipitate. The supernatant can be further treated as described in Section 32.3.1. Gu et al. [29] studied selenoprotein W in monkey muscle. About 180 g of the frozen skeletal muscle was chopped with scissors and 4 volumes (w/v) of 50 mM sodium phosphate buffer, pH 6.5 containing 0.1 mM phenylmethyl sulforyl ﬂuoride (PMSF) and 0.02% sodium azide (buffer A) added. This mixture was homogenized with an Omnimixerw for 3 min at maximum speed. The homogenate was centrifuged at 10; 000g for 30 min and the supernatant ﬁltered through two layers of cheese cloth and recentrifuged at 100; 000g for 90 min using an ultracentrifuge. The supernatant was concentrated with slow stirring under nitrogen using an Amicon cell. The concentrated cytosols were chromatographed in a large Sephadex G-50 column with buffer A as the mobile phase. Detection for protein was made at 280 nm and for selenoprotein W by slot blots. A pool of low molecular weight immunoreactive column eluant was concentrated to 10 ml. The concentrate was re-chromatographed on a smaller Sephadex G-50. The immunoreactive fractions were pooled to exclude signiﬁcant contamination from pink color.

1049

C.P. de Leon et al.

The G-50 pool was adjusted to 100 ml with buffer A and chromatographed on a CM-Sephadex cation-exchange column. The selenoprotein was eluted with 400 ml linear gradient of 0–30 M NaCl in buffer A. The immunoreactive fractions were pooled and concentrated to 250 ml using a centrifugable microporous concentrator. This preparation was chromatographed on a C-18 reverse phase with 30 – 60% acetonitrile gradient in water with 0.1 M triﬂuoroacetic acid. The protein was monitored at 230 nm. Fractions of the column eluant were again assayed by slot blots for selenoprotein W. Rat liver homogenate for the speciation of metabolites of selenate has been analyzed by Shiobara et al. [30]; selenate was incubated in a sample of 4 mM of liver homogenate for 0, 30 min and 4 h. The incubated solution was heat treated at 758C for 5 min and then centrifuged at 8000g for 10 s to remove heat-unstable proteins. The supernatant was analyzed by SEC with 50 mM Tris– HCl buffer at pH 7.4. LMM can also be determined in tissue samples. Quijano et al. [31] speciated selenocysteine, trimethylselenonium, selenomethionine, selenoethionine, selenite and selenate in tuna and mussel samples. Fats were extracted from marine materials in two steps with 6 ml of 2:1 and 1:2 CH3OH–CHCl3, respectively. The samples were hydrolyzed enzymatically in two steps, ﬁrst with a nonspeciﬁc protease (pronase E) and second with another protease. The enzymatic digest was passed through C18 cartridges previously washed with 3.5 mM phosphate buffer at pH 6.0. These extracts were then injected on a C18 ODS Spherisorb column utilizing two different mobile phases: 5 mM phosphate buffer at pH 2.8 and 3.5 –7.0 mM phosphate buffer at pH 6.0. 32.4

FISH/BIRDS

Herring gull eggs are among a set of environmental biomarkers and are hence used to determine levels of selenium in the environment. In the work performed by Jakubowski et al. [32] for the analysis of the selenoamino acids, samples were frozen with liquid nitrogen upon collection. An aliquot was extracted by shaking with a 9:1 methanol:water mixture overnight and then centrifuged. Methanol was removed from the decanted solution by evaporation at reduced pressure and the resulting residue redissolved in DDI water. The aqueous extract was ﬁltered through cellulose acetate ﬁlters with a pore diameter of 0.45 mm prior to HPLC analysis. Lindemann et al. [33] were interested in the analysis of ﬁsh for Se(IV), Se(VI) and selenomethionine. A 1 g dry mass of ﬁsh material was extracted with a 50:50 mixture of methanol:water in an ultrasonic bath for 20 min. This was followed by centrifugation and decanting. The resulting supernatant was taken to dryness with a rotary evaporator and the residue redissolved in DDI water. The extracts were ﬁltered with a 0.45 mm PTFE membrane ﬁlter prior to injection for HPLC analysis.

1050

Sample preparation for speciation of selenium

Crews et al. [34] investigated the bioavailability of selenium in cod to humans. Cooking was therefore part of the sample preparation procedure; skin and bones were removed after cooking and the remaining ﬂesh was homogenized. The cooked cod sample was incubated at 378C for 4 h with 10 ml of gastric juice (1% m/v pepsin in 0.15 M NaCl, acidiﬁed with HCl to pH 2.0). Once complete, the sample was adjusted to pH 4.8 with saturated NaHCO3 and 10 ml of gastrointestinal juice (1.5% pancreatin, 0.5% amylase and 0.15% bile salts, m/v, in 0.15 M NaCl) was added. The pH was then adjusted to 6.9 and the samples incubated for a further 4 h at 378C. The samples were then centrifuged and the supernatant used for analysis of selenomethionine, selenocystine, sodium selenite and sodium selenate by HPLC. Investigation of enzymatic treatments as a means to extract selenium species from ﬁsh samples was undertaken by Quijano et al. [31] First, the fats were extracted from the marine materials with mixtures of methanol and chloroform. The selenium compounds were extracted by a two-step enzymatic hydrolysis. First, a non-speciﬁc protease (pronase E, subtilisin or a mixture of both) and 5 ml of 0.1 M phosphate buffer at pH 7.5 were added to the sample. Incubation was performed for 24 h at 378C. This enzymatic treatment was repeated and the resulting extracts were passed through a C18 cartridge. Selenium compounds of interest were eluted with 3.5 mM phosphate buffer at pH 6.0 and the resulting extracts were analyzed using HPLC. The enzymatic procedure was applied to both tuna and mussel samples. Gomez-Ariza et al. [35] also used enzymatic treatments in their investigation of selenium species in shellﬁsh. To freeze-dried samples, a mixture of protease XIV, lipase VII and protease VIII was added and the sample shaken at 300 rpm in the dark for 24 h using a mechanical shaker at 378C. The supernatant was then separated from the sample by centrifugation and ﬁltered with 0.45 mm ﬁlters to eliminate suspended solids. Additionally, the solution was passed through a 10,000 Da molecular weight cut-off ﬁlter. Because of the complexity of the matrix, clean-up procedures were required to remove interferents that affected the chromatographic separation. The extracts were ﬁrst partitioned in dichloromethane and then passed through a column with aminopropylsilane. The selenium compounds were eluted with DDI water that had been adjusted to pH 9.8. 32.5

PLANTS

Selenium levels in grains and vegetables depend on the selenium content of the soil where the food is raised. [36] The selenium in the soil is taken up by plants (it is unclear as to whether it is absorbed or transported), and subsequently, to humans and animals that eat them [37]. Natural selenium levels in the soil are highly variable throughout the world [38]. In plants, selenium may be associated with a complex matrix and some of the selenium accumulated or absorbed is built into proteins or exists as free

1051

C.P. de Leon et al.

selenoamino acids. Acquisition of the particular tissues of interest and homogenization are then the ﬁrst tasks to be accomplished. Immersion of the sample of interest in liquid nitrogen has been successfully used to break up the plant matrix [39]. Once the possession of a representative sample is achieved, the extraction of free selenoamino acids in plants can be accomplished with mild acid solution and further analyzed with the techniques of choice. Alternatively, an appropriate hydrolysis procedure may be applied that will allow for complete destruction of the matrix while simultaneously preserving the selenium species of interest. Generally, the requirement for a nondestructive sample procedure can be fulﬁlled with the use of enzymes for the protein-incorporated selenium. This is due to the inherent properties of enzymes; they break in a speciﬁc and well-known process under moderate pH and temperature conditions [40]. Finally, in some instances, it is necessary to clean-up the extract, particularly if a selenium-selective detector is not to be used. 32.5.1 Leafy plants Montes-Bayon et al. studied the selenium species of Brassica juncea, a selenium hyperaccumulating plant that may be used in phytoremediation processes [39]. The authors evaluated selenium extraction using several digestion/extraction procedures, including the use of HCl, Tris–HCl buffer and enzymatic hydrolysis (using proteinase K and protease XIV). It was found that the best extraction was obtained with proteinase K, which extracted about 75% of the total selenium present in the plant. Meija et al. analyzed the volatile selenium species of the same plant [41]. The plants of interest were grown in a selenium-rich medium and subsequently examined for the elucidation of the biochemical steps of this process, known as phytovolatilization. After the seedlings were 2–3 days old, they were placed in 10 ml glass vials sealed with caps containing rubber septa. The volatile selenium compounds were extracted in the headspace by solid-phase microextraction, a relatively new extraction technique whereby the analytes of interest are absorbed into a polymeric substance coated on a fused silica ﬁber. Once equilibrium was reached, the volatile selenium species were desorbed in the injection port of a GC, and subsequent separation and ICP-MS detection were performed. Other sample preparation techniques were investigated by Kotrebai et al. for the selenium speciation studies of such hyperaccumulating plants, Astragalus praleongus and B. juncea [42]. In this work, hot water and enzymatic extractions (protease XIV) were utilized for sample preparation. Results indicated that the two extraction procedures were equally effective with the exception of their ability to extract selenomethionine; the hot water procedure did not extract this selenium species.

1052

Sample preparation for speciation of selenium

Zhang and Frankenberger undertook a different approach to the task of sample preparation for selenium speciation in plant matrices [43]. In their work, they extracted the selenium species from a freeze-dried plant and then used a sample preparatory column containing an anion-exchange resin to separate the selenium compounds in the aqueous extract. Detection was thus performed on the fraction containing each individual selenium compound. 32.5.2 Broccoli Broccoli samples, in addition to other foods, were prepared for selenium speciation by Cai et al. [44] A ﬁnely powdered, freeze-dried sample was suspended in 0.1 N HCl using a vortex mixer. The sample was then centrifuged; speciﬁc volumes of ethanol, pyridine and ClCO2Et were sequentially added to the supernatant to effect derivatization of the selenoamino acids. Once the reaction was complete, the derivatives were extracted into chloroform and injected into GC. 32.5.3 Spices (garlic, onion, white clover) McSheehy et al. investigated speciation of selenium in garlic harvested in naturally seleniferous soil [7]. Freeze-dried samples were leached with water and the aqueous extract was fractionated by preparative size-exclusion chromatography to separate selenium species by molecular weight. Each fraction was then individually analyzed by reversed-phase liquid chromatography with ICP-MS detection. In the selenium speciation of three members of the Allium family, garlic, onions and ramp, Kotrebai et al. utilized the same hot water and enzymatic procedures that had previously been applied to plants [42]. Again, it was found that the enzymatic treatment allowed the recovery of various selenoamino acids that were not liberated from the matrix with the hot water extraction. Bird et al. utilized speciﬁc extraction procedures that correlated with the intended liquid chromatography technique to be utilized for separation of selenium species in garlic [45]. For ion-exchange HPLC, samples were sonicated with a mixture of 10% methanol in 0.2 M HCl. In the application of reversed-phase HPLC, the samples were extracted in the same manner, however, the selenoamino acids were then converted to N-2,4-dinitrophenyl (DNP) derivatives prior to chromatography so as to allow for their detection. Four extraction techniques were examined for the liberation of Se species from freeze-dried white clover by Emteborg et al. [46] These included mechanical shaking with a 2:3:5 mixture of water:chloroform:methanol, sonication with a 1:1 methanol:water mixture, sonication with a 1:1 methanol: water mixture (0.28 M HCl added) and sonication with a 1:1 methanol:water mixture (4% ammonia). The extracts were then ﬁltered and the pH was adjusted to 9.5. The extracts were cleaned by passage through end-capped C18

1053

C.P. de Leon et al.

cartridges. Evaluation of the four extracts showed the best extraction efﬁciency for the selenium analytes was obtained with 1:1 methanol:water (4% ammonia) and it was speculated that this extraction mixture most effectively liberated the selenoamino acids from possible protein-binding sites. 32.5.4 Grains Calle-Guntinas et al. [47] developed a method for the analysis of selenomethionine in wheat. First, a volume of tetramethylammonium hydroxide solution was added to the wheat sample to effect hydrolysis of the proteins. This mixture was shaken for 4 h at 608C. The resulting extract was cleaned on an aluminum oxide column. Finally, a derivatization reaction was performed to affect the suitability of the selenomethionine for GC analysis. The carboxylic acid was esteriﬁed with propan-2-ol in a 4 M HCl medium and the amino group was acylated with heptaﬂuorobutyric anhydride. 32.5.5 Nuts Several sample preparation strategies were examined by Vonderheide et al. with the goal of selenium speciation in Brazil nuts [48]. These included mild microwave-assisted extractions and an enzymatic digestion. The enzymatic digestion proved to be the most effective; it was concluded that selenium was in the form of selenoamino acids that were incorporated into the plant proteins. Interestingly, Brazil nuts are high in fat and removal of this lipid fraction was deemed necessary to obtain accurate analytical analysis. Removal of the lipids was accomplished by the addition of an equal volume of 1:2 methanol:chloroform mixture to the nut samples. The solvent/nut mixture was sonicated and the lipid layer removed with vacuum ﬁltration. Other subsequent work done by the same authors presented a second means of extracting the selenium species from the nut matrix. In this procedure, the proteins were isolated from the nut samples by dissolution in 0.1 M sodium hydroxide and subsequent precipitation with acetone. The protein fraction was then treated with the same nonspeciﬁc protease (proteinase K). This sample preparation procedure was also applied to other types of nuts in the second manuscript, including walnuts, black walnuts, cashews and pecans [49]. 32.5.6 Mushrooms A sequential sample preparation process was developed by Stefanka et al. for the speciation of Se-enriched edible mushrooms [40]. Five different extraction methods were compared, each using a different combination of three enzymes: pepsin, trypsin and pronase. The most efﬁcient was found to be a three-step process involving the use of water extraction and two proteolytic enzymes, pepsin and trypsin. Also worthy of note is the observation that sufﬁcient

1054

Sample preparation for speciation of selenium

extraction efﬁciency could not be achieved without proteolytic enzymes, as neither water nor buffer extractions was capable of recovering more than 40% of the selenium present in Se-enriched mushrooms. This further illustrates that selenium is partially bound to proteins. 32.6

MICROORGANISMS

Microorganisms can incorporate selenium in a non-speciﬁc way (by sulfur replacement) or as part of a protein (with selenocysteine as the only naturally occurring selenoamino acid). The isolation of selenoproteins has been done for over 20 years and although it was left for some years, interest is now reemerging. Many years ago, Jones et al. [50] utilized a formate enzyme after suspending the cells in 50 mM phosphate (pH 7) containing 2 mM mercaptoethanol. Treatment with deoxyribonuclease and sonication followed. Cell debris was separated by centrifugation; the extract was heated at 608C for 10 min. This procedure denatured up to 40% of the total proteins, which were then sedimented by centrifugation, while the enzyme remained in the supernatant. Separation of the selenoproteins was achieved utilizing sizeexclusion chromatography with the appropriate detection system. The elucidation of the selenoamino acids occurring in the proteins was ﬁrst done by Cone et al. [51] in clostridial glycine reductase utilizing a trypsin enzymatic digestion. Since then, different enzymes, such as protease XIV [52], pronase E [53] and proteinase K [54], have been widely used in the enzymatic hydrolysis when analyzing selenium compounds. In general, enzymatic hydrolysis involves the addition of the chosen enzyme to the sample (e.g. 1:10 ratio, respectively) in a neutral aqueous media and subsequent incubation at 378C. The fractions of selenoamino acids obtained can then be separated with liquid chromatography and detected with a suitable detector (e.g. UV, ICP). It is worthy of note that when hydrolyzing Se proteins, care has to be taken to minimize the oxidation of the labile selenium compounds. Procedures can be used to prevent degradation. These include the use of anaerobic conditions or the preparation of carboxymethyl derivatives of the selenoamino acids. 32.7

ENVIRONMENTAL

32.7.1 Air Volatile selenium species may enter the atmosphere because of the natural metabolic processes of plants. Additionally, selenium species are introduced into the air as trace by-products of the coal combustion process. The formation of various species depends on the temperature, atmosphere (reducing or oxidizing conditions) and chlorine content of the combustion process [55]. The most common method for the determination of volatile species involves sampling a speciﬁc volume of air through an inert container enclosing a solid

1055

C.P. de Leon et al.

sorbent. It is intended that species of interest are absorbed onto the sorbent and then subsequently desorbed by either liquid or gas and then separated by a GC column. Pecheyran et al. developed a method which utilized cryofocusing for sample collection and preparation [56]. Volatile analytes were collected by cryofocusing on a small glass wool packed column at 2 1758C; samples were then ﬂash desorbed and analysis was performed by GC –ICP-MS. Profumo et al. developed a sequential extraction procedure for the speciation of inorganic selenium in air samples [57]. The procedure entailed the selective sequential solubilization of the inorganic selenium compounds and allowed the determination of Se(0), Se(IV) and Se(VI). As is a known fact, organoselenium compounds are present in the headspace of cut or crushed garlic. Cai et al. investigated the natural abundance of these compounds in human breath after the ingestion of garlic. Breath samples were collected by expiration into two 1.5 l Tedlar air-sampling bags. The contents of the bags were then suctioned through a tenax trap and the trap inserted into the injection port of a GC. Resulting data showed the major organoselenium compound to be dimethyl selenide with lesser amounts of several other volatile selenium species [58]. 32.7.2 Water Generally, water samples may ﬁrst need to be ﬁltered before analysis of selenium species. Filtering the samples ensures the necessary removal of the particulates that may impede analysis, though it may lead to losses of volatile selenium species. Not all aqueous samples require ﬁltration, however, deletion of this step may result in changes to the distribution of the selenium species over time. This phenomenon would be a result of adsorption and desorption processes occurring at the particles’ surfaces. The selenium analytes can be analyzed within the aqueous sample or the analytes can be extracted into an organic solvent and this phase then analyzed. The choice between the two methods depends on the analytes of interest. Polar analytes might best be separated in the aqueous sample by HPLC; polar analytes could also be derivatized to reduce their polarity and separated by GC. Non-polar analytes may be extracted into an organic media, potentially derivatized, and subsequently separated by GC. Several derivatization methods have been investigated, including hydride generation, reaction with tetraalkylborates and reaction with alkylchloroformates. Selenium readily forms a hydride, but only in its lower oxidation state. Therefore, Se(VI) needs to reduced prior to formation of the hydride. Se(IV) will also readily react with sodium tetraethylborate for a selective analysis of this ion. Selenoamino acids were esteriﬁed and acylated with ethylchloroformate by Pelaez et al. [59] to ensure their amenability to GC analysis. A relatively new extraction technique, solid-phase microextraction, was applied in the analysis of Se(IV) by Guidotti [60]. This procedure entails the use

1056

Sample preparation for speciation of selenium

of a polymeric-coated fused silica ﬁber as described above. Once the extraction is complete, the ﬁber is placed in the injection port of GC and the analytes are thermally desorbed. In this work, due to the low volatility of Se(IV), it was selectively derivatized to ethane, 1,1’-selenobis by reaction with sodium tetraethylborate. Both headspace and direct immersion procedures were studied. In most work performed to date, the inorganic species of selenium have been of greatest interest in the analysis of water samples. Subsequently, many researchers have worked in different manners to separate Se(IV) and Se(VI). In method development conducted by Raessler et al. [61], HPLC was used as the means of separating Se(IV) from Se(VI) in order to determine individual species’ concentrations. Two fractions were collected from the HPLC efﬂuent in time frames corresponding to the elution of each. The fractions were then analyzed by HG-AAS to obtain concentrations of the two selenium species. Methods for the analysis of both Se(IV) and Se(VI) may entail determination of Se(IV) and total inorganic selenium [after reduction of Se(VI)] and the difference calculated as the concentration of Se(VI) [62]. In the work conducted by Wang et al. [63], various reagents were combined with microwave digestion techniques in stepwise procedures designed to obtain concentrations of Se(0), Se(2 II), Se(IV) and Se(VI). Each procedure employed reagents that had different effects on the various oxidation states. Concentrations of each of the species were determined by subtraction and process of elimination calculations. In a similar manner of calculation, Gregori et al. [64] divided each aqueous sample into two aliquots and, in the ﬁrst, determined total selenium by acid microwave digestion. The second aliquot was used to selectively determine Se(IV) by both electrochemical and spectroscopic methods. Once accomplished, Se(VI) in this second aliquot was chemically reduced to Se(IV) by addition of HCl and heating, and the concentration of Se(IV) subsequently measured. The concentration of Se(VI) was then calculated as the difference between these two determinations. Se(2II) was calculated as the difference between total Se and the sum of Se(IV) and Se(VI). Others have separated the inorganic species chromatographically with the employment of online detection schemes. Wallschlager et al. [65] used ion chromatography –ICP-MS to sequentially separate and detect Se(IV), Se(VI) and SeCN2. It may also be in the interest of the analyst to concentrate the selenium species prior to analysis to obtain lower limits of detection. GomezAriza et al. [66] utilized an anion-exchanger phase (SAX) inline with a hydrophobic phase (C18) for the preconcentration of selenite [Se(IV)], selenate [Se(VI)], dimethylselenide, dimethyldiselenide, diethylselenide and diethyldiselenide. Preconcentration of the inorganic species was based on their retention in the SAX cartridge; Se(IV) was eluted with 1 M formic acid followed by elution of Se(VI) with 1 M HCl. Organic species were retained in the C18 phase and subsequently eluted with carbon disulﬁde.

1057

C.P. de Leon et al.

32.7.3 Soil and sediments (solid matrices) Jackson and Miller investigated the extraction of Se(IV) and Se(VI) from coal ﬂy ash. [67] This group used both deionized water and an aqueous buffer solution ðpH ¼ 5Þ as extraction solvents. Once this aqueous volume was added to the solid samples, they were shaken for 16 h and then ﬁltered prior to analysis. After analysis of 24 samples, they concluded that the total extractable selenium concentration measured in the two extracts was substantially different. The buffered extract yielded much lower Se(VI) concentrations than the deionized water extracts while neither extraction method produced signiﬁcant concentrations of Se(VI). The analysis of wetlands is of great interest as they have been used to remove trace elements, including selenium, from contaminated wastewaters, and several fractionation procedures have been developed for the speciation of selenium. These types of samples are generally dried prior to analysis. Zhang and Moore [68] point out the possible inconvenience of the procedure, e.g. increasing the selenium concentration in the samples or selenium oxidation to selenate. These changes are especially notorious at higher temperatures. Gao et al. [69] analyzed soluble, adsorbed (or ligand exchangeable) and carbonate-associated Se, in a wetland system in the Tulare Lake using, respectively, 0.25 M KCl, 0.1 M K2HPO4 (pH 8) and 1.0 M sodium acetate (pH 5). The organic selenium was determined using 0.1 M NaOH after extraction with KCl and KH2PO4. Elemental selenium was extracted by formation of the sulﬁte complex with 1 M Na2SO3. Similarly, a sequential extraction scheme was developed by Martens and Suarez [70] to identify selenium oxidation states in several fractions. The ﬁrst step utilized a 0.1 M K2HPO4 –KH2PO4 buffer at pH 7.0 to release soluble Se(IV), Se(VI) and Se(2 II). The second step involved oxidation of organic materials with 0.1 M K2S2O8 at 908C to release Se(2 II) and Se(IV) associated or occluded with organic matter. The ﬁnal step used nitric acid (908C) to solubilize insoluble selenium remaining in the sample. These selenium fractions were then speciated by a selective hydride generation atomic absorption spectrophotometry technique to determine the selenium concentration in each individual oxidation state. A volume of 50 ml of extraction solvent [0.3 M ammonium acetate (pH 4.6), 0.3 M ammonium oxalate (pH 3.0), Milli-Q water (pH 5.8), 0.3 M sodium hydrogen carbonate (pH 8.0) and 0.3 M sodium carbonate (pH 11.0)] was used by Vassileva et al. for the extraction of 5 g of soil sample [71]. The samples were extracted by agitating for 4 h at room temperature followed by ﬁltration through membrane ﬁlters with a pore size of 45 mm. The extracts were analyzed for both Se(IV) and Se(VI) using ion chromatography coupled with ICP-MS detection. Zhang et al. considered soluble selenium in both agricultural drainage waters and aqueous soil-sediment extracts using HG-AAS [72]. The soluble

1058

Sample preparation for speciation of selenium

selenium was ﬁrst removed from the soil sample by extraction with deionized water followed by centrifugation and ﬁltration through a 0.45 mm membrane. Se(IV) concentrations were then determined by HG-AAS. Persulfate was added to oxidize organic Se(2II), and manganese oxide was used as an indicator for oxidation completion. The samples were placed in a hot water bath (808C) for 20 min and then transferred to a room temperature water bath for another 20 min. The organic selenium was calculated as the difference between the selenium in this solution and the Se(IV) concentration as determined previously. Gomez-Ariza et al. [73] investigated volatile selenium in sediment samples. Species considered included dimethylselenide, dimethyldiselenide and diethylselenide. Instrumentation was based on a coupling between a prevaporation module, a preconcentration sorptive trap and a GC –MS. Volatile selenium compounds were purged from the sediment sample, trapped on a tenax column and thermally desorbed into the GC.

REFERENCES 1 2 3 4 5 6 7 8 9

10

11 12 13 14

K. Pyrzynska, Anal. Sci., 14 (1998) 479 –483. C.D. Thomson, Analyst, 123 (1998) 827–831. J. Szpunar and R. Lobinski, Trends Anal. Chem., 363 (1999) 550– 557. S. Fairweather-Tait and R.F. Hurrell, Nutr. Res. Rev., 9 (1996) 295 –324. M. Kotrebai, J.F. Tyson, E. Block and P.C. Uden, J. Chromatogr. A, 866 (2000) 51–63. R. Lobinski, J.S. Edmonds, K.T. Suzuki and P.C. Uden, Pure Appl. Chem., 72 (2000) 447– 461. S. McSheehy, W. Yang, F. Pannier, J. Szpunar, R. Lobinski, J. Auger and M. PotinGautier, Anal. Chim. Acta, 421 (2000) 147– 153. J.A. Caruso, K.L. Sutton and K.L. Ackley, Elemental Speciation: New Approaches for Trace Element Analysis. Elsevier, New York, 2000. L.C. Clark, B.W. Turnball, E.H. Slate, D.K. Chalker, J. Chow, L.S. Davis, R.A. Glover, D.K. Graham, E.G. Gross, A. Krongrad, J.L. Lesher, H.K. Park, B.B. Sanders, C.L. Smith, J.R. Taylor, D.S. Alberts, R.J. Allison, J.C. Bradshaw, D. Curtus, D.R. Deal, M. Dellasega, J.D. Hendrix, J.H. Herlong, L.J. Hixon, F. Knight, J. Moore, J.S. Rice, A.I. Rogers, B. Schuman, E.H. Smith and J.C. Woodard, J. Am. Med. Assoc., 276 (1996) 1957. L.C. Clark, B. Dalkin, A. Krongrad, G.F.J. Combs, B.W. Turnbull, E.H. Slate, R. Witherington, J.H. Herlong, E. Janosko and D. Carpenter, Br. J. Urol., 81 (1998) 730 –734. R. Cornelis, J.D. Kimpe and X. Shang, Spectrochim. Acta B, 53 (1998) 187–196. A. Sanz-Medel, Spectrochim. Acta B, 53 (1998) 197–211. P.D. Whanger, Y. Xia and C.D. Thompson, J. Trace Elem. Electrolytes Health Dis., 8 (1994) 1– 7. J.M. Gonzalez-La Fuente, J.M. Marchante-Gayon, M.L. Fernandez-Sanchez and A.S. Medel, Talanta, 50 (1999) 207– 217.

1059

C.P. de Leon et al. 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36

37 38 39 40 41 42 43 44 45

1060

M.A. Quijano, A.M. Gutierrez, M.C. Perez-Conde and C. Camara, Talanta, 50 (1999) 165–173. M.M. Gomez, T. Gasparic, M.A. Palcios and C. Camara, Anal. Chim. Acta, 374 (1998) 241–251. M. Sanz-Alejo and C. Diaz-Romero, Chem. Rev., 95 (1995) 227. B. Gammelgaard, O. Jons and L. Bendahl, J. Anal. At. Spectrom., 16 (2001) 339–344. H. Koyama, K. Omura, A. Ejima, Y. Kasanuma, C. Watanabe and H. Satoh, Anal. Biochem., 267 (1999) 84– 91. C. Sasakura and K.T. Susuki, J. Inorg. Biochem., 71 (1998) 159– 162. K.T. Susuki, M. Itoh and M. Ohmichi, J. Chromatogr. B, 666 (1995) 13–19. I.A. Bergdahl, A. Schuetz and A. Grubb, J. Anal. At. Spectrom., 11 (1996) 735. B. Gercken and R.M. Barnes, Anal. Chem., 63 (1991) 283 –287. F. Laborda, M.V. Vicente, J.M. Mir and J.R. Castillo, Fresenius J. Anal. Chem., 357 (1997) 837–843. Y. Makino and S. Nishimura, J. Chromatogr., 579 (1992) 346– 349. B. Michalke, J. Chromatogr. A, 716 (1995) 323 –329. P. Bermejo, J. Barciela, E.M. Pena, A. Bermejo, J.M. Fraga and J.A. Cocho, J. Anal. At. Spectrom., 16 (2001) 188– 193. A. Theobald and L. Dunemann, J. High Resolut. Chromatogr., 19 (1996) 608– 612. Q.-P. Gu, M.A. Beilstein, E. Barofsky, W. Ream and P.D. Whanger, Arch. Biochem. Biophys., 361 (1999) 25–33. Y. Shiobara, Y. Ogra and K.T. Suzuki, Analyst, 124 (1999) 1237–1241. M.A. Quijano, P. Moreno, A.M. Gutierrez, M.C. Perez-Conde and C. Camara, J. Mass Spectrom., 35 (2000) 878– 884. N. Jakubowski, D. Stuewer, D. Klockow, C. Thomas and H. Emons, J. Anal. At. Spectrom., 16 (2001) 135 –139. T. Lindemann, A. Prange, W. Dannecker and B. Neidhart, Anal. Bioanal. Chem., 368 (2000) 214– 220. H.M. Crews, P.A. Clarke, D.J. Lewis, L.M. Owen, P.R. Strutt and A. Izquierdo, J. Anal. At. Spectrom., 11 (1996) 1177–1182. J.L. Gomez-Ariza, M.A.C.d.l. Torre, I. Giraldez, D. Sanchez-Rodas, A. Velasco and E. Morales, Appl. Organomet. Chem., 16 (2002) 265–270. M.P. Longnecker, P.R. Taylor, O.A. Levander, M. Howe, C. Weillon, P.A. McAdam, K.Y. Patterson, J.M. Holden, M.J. Stampfer, J.S. Morris and W.C. Willett, Am. J. Clin. Nutr., 53 (1991) 1288–1294. M. Stadlober, M. Sager and K.J. Irgolic, Food Chem., 73 (2001) 357– 366. J.C. Chang, W.H. Gutenmann, C.M. Reid and D.J. Lisk, Chemosphere, 30 (1995) 801 –802. M. Montes-Bayon, E.G. Yanes, C.P.d. Leon, K. Jayasimhulu, A. Stalcup, J. Shann and J.A. Caruso, Anal. Chem., 74 (2002) 107–113. Z. Stefanka, I. Ipolyi, M. Dernovis and P. Fodor, Talanta, 55 (2001) 437–447. J. Meija, M. Montes-Bayon and J.A. Caruso, Analytical Chemistry, 74 (2002) 5837– 5844. M. Kotrebai, M. Birringer, J.F. Tyson, E. Block and P.C. Uden, Analyst, 125 (1999) 71– 78. Y. Zhang and W.T. Frankenberger Jr., Sci. Total Environ., 269 (2001) 39–47. X.J. Cai, E. Block, P.C. Uden, X. Zhang, B.D. Quimby and J.J. Sullivan, J. Agric. Food Chem., 43 (1995) 1754–1757. S.M. Bird, H. Ge, P.C. Uden, J.F. Tyson, E. Block and E. Denoyer, J. Chromatogr. A, 789 (1997) 349– 359.

Sample preparation for speciation of selenium 46 47 48

49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73

H. Emteborg, G. Bordin and A.R. Rodriguez, Analyst, 123 (1998) 245– 253. M.B.D.L. Calle-Guntinas, C. Brunori, R. Scerbo, S. Chiavarini, P. Quevauviller, F. Adams and R. Morabito, J. Anal. At. Spectrom., 12 (1997) 1041– 1046. A.P. Vonderheide, K. Wrobel, S. Kannamkumarath, C. B’Hymer, M. Montes-Bayo´n, C. Ponce de Leon and J.A. Caruso, Journal of Agricultural and Food Chemistry, 50 (2002) 5722– 5728. S.S. Kannamkumarath, K. Wrobel, K. Wrobel, A.P. Vonderheide and J.A. Caruso, Analytical and Bioanalytical Chemistry, 373 (2002) 454– 460. J.B. Jones, G.L. Dilworth and T.C. Stadtman, Arch. BioChem. Biophys., 195 (1979) 255 –260. J.E. Cone, R.M.d. Rio, J.N. Davis and T.C. Stadtman, Proc. Natl Acad. Sci., 73 (1976) 2659– 2670. C. Casiot, J. Szpunar, R. Lobinski and M. Potin-Gautier, J. Anal. At. Spectrom., 14 (1999) 645–650. B. Michalke, H. Witte and P. Schramel, J. Anal. At. Spectrom., 16 (2001) 593– 597. C.A. Ponce de Leon, M. Montes-Bayon and J.A. Caruso, J. Appl. Microbiol., 92 (2002) 602–610. R. Yan, D. Gauthier and G. Flamant, Combustion Flame, 120 (2000) 49– 60. C. Pecheyran, C.R. Quetel, F.M.M. Lecuyer and O.F.X. Donard, Anal. Chem., 70 (1998) 2639– 2645. A. Profumo, G. Spini, L. Cucca and B. Mannucci, Talanta, 55 (2001) 153–161. X.J. Cai, E. Block, P.C. Uden, B.D. Quimby and J.J. Sullivan, J. Agric. Food Chem., 43 (1995) 1751– 1753. M.V. Pelaez, M. Montes-Bayon, J.I.G. Alonso and A. Sanz-Medel, J. Anal. At. Spectrom., 15 (2000) 1217– 1222. M. Guidotti, J. AOAC Int., 83 (2000) 1082–1085. M. Raessler, B. Michalke, S. Schulte-Hostede and A. Kettrup, Sci. Total Environ., 258 (2000) 171– 181. C. Camera, R. Cornelis and P. Quevauviller, Trends Anal. Chem., 19 (2000) 189 –194. Z. Wang, Y.X. Gao and N. Belzile, Anal. Chem., 73 (2001) 4711 –4716. I.D. Gregori, M.G. Lobos and H. Pinochet, Water Res., 36 (2002) 115–122. D. Wallschlager and R. Roehl, J. Anal. At. Spectrom., 16 (2001) 922 –925. J.L. Gomez-Ariza, J.A. Pozas, I. Giraldez and E. Morales, Analyst, 124 (1999) 75– 78. B.P. Jackson and W.P. Miller, J. Anal. At. Spectrom., 13 (1998) 1107 –1112. Y.Q. Zhang and J.N. Moore, Commun. Soil Sci. Plants Anal., 28 (1997) 341– 350. S. Gao, K.K. Tanji, D.W. Peters and M.J. Herbel, J. Environ. Qual., 29 (2000) 1275– 1283. D.A. Martens and D.L. Suarez, Environ. Sci. Technol., 31 (1997) 133–139. E. Vassileva, A. Becker and J.A.C. Broekaert, Anal. Chim. Acta, 441 (2001) 135 –146. Y. Zhang, J.N. Moore and W.T. Frankenberger, Environ. Sci. Technol., 33 (1999) 1652– 1656. J.L. Gomez-Ariza, A. Velasco-Arjona, I. Giraldez, D. Sanchez-Rodas and E. Morales, Int. J. Environ. Anal. Chem., 78 (2000) 427– 440.

1061

This page intentionally left blank

Chapter 33

Sample preparation for mercury speciation Holger Hintelmann

33.1

INTRODUCTION

Mercury is an ubiquitous element and of great concern as an environmental pollutant. A remarkable number of aquatic organisms in many ecosystems all over the world show elevated Hg levels. Freshwater ﬁsh communities are known to be contaminated with toxic and bioavailable methylmercury (MeHg) [1]. Consequently, many countries have issued advisories to manage the consumption of ﬁsh, representing the main entry of methylmercury into the human diet and thus, being the main concern from a human health point of view. Numerous studies have been conducted to elucidate the fate of Hg species in the environment and to understand the factors controlling methylmercury formation and bioaccumulation. Similarly, many different methods have been developed to accurately determine the concentrations of Hg species in various environmental matrices [2]. Despite those efforts the accurate determination of methylmercury remains a challenge for analytical chemists. Natural levels of methylmercury span almost eight orders of magnitude, from low pg/l in surface waters to mg/kg in ﬁsh. An overview of commonly found concentrations in a variety matrices is given in Table 33.1. Clearly, to accommodate such an enormous range of concentrations, sample preparation techniques must be carefully selected considering the ﬁnal analytical technique. Numerous variations of a few standard procedures have been developed during the past 40 years. This chapter focuses on a few selected methods that have been established in the authors’ laboratory and successfully used for the routine analysis of a large number of samples encompassing the full range of environmental matrices.

33.2

AQUEOUS SOLUTION CHEMISTRY OF METHYLMERCURY

The methylmercuric ion CH3Hgþ shows a very rich, yet fascinatingly simple coordination chemistry in aqueous solution. CH3Hgþ has a strong preference Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

1063

H. Hintelmann TABLE 33.1 Range of methylmercury concentrations commonly found in environmental samples Matrix Air Water Freshwater Oceans Rain Snow Solids Forest soils Lake sediments Vegetation Aquatic wildlife Fish Zooplankton Zoobenthos Phytoplankton Mammals Whales Otter Birds

Concentration rangea

Reference

0– 20

[3]

0.02– 1.5 0.01– 0.1 0.06– 0.4 0.01– 0.1

[4] [5] [6] [7]

0.05– 0.5 0.05– 5 0.005– 1

[8] [9] [6]

100– 2000 10– 500 30– 800 100– 300

[10] [11] [12] [13]

100– 1500 100– 1500 30– 500

[14] [15] [16]

a

Air concentrations in pg/m3, water concentrations in ng/l, all others in ng/g.

for the co-ordination number of one in its aqueous complexes and as such shows many similarities with the proton Hþ. An example is the ability of CH3Hgþ to form multinuclear complexes analogue to H3Oþ, where it can readily substitute the water protons: CH3HgOH2 þ, (CH3Hg)2OHþ and (CH3Hg)3Oþ have all been reported to exist in aqueous solution [17,18]. However, the stability constants of corresponding complexes differ widely as a result of the different character of the two cations. Hþ is the simplest “hard” Lewis acid, whereas CH3Hgþ is the simplest “soft” Lewis acid preferring coordination with “soft” Lewis bases. Schwarzenbach and Schellenberg [19] have determined a series of stability constants for the CH3HgL system. As expected of a soft Lewis acid, CH3Hgþ forms extremely strong complexes with sulﬁde ligands, which are preferred over N and O derivatives. Halogens form stable complexes in the order F , Cl , Br , I. Compared to the mercuric ion Hg2þ, CH3Hgþ stability constants are slightly smaller, the difference increasing as the “softness” of the ligand increases. This may be attributed to the soft CH3-ligand, which decreases the afﬁnity of Hg for other soft bases. In contrast to Hg2þ, CH3Hgþ forms only few multiligand complexes of the nature CH3HgLn (12n).

1064

Sample preparation for mercury speciation

Only the existence of CH3HgI22 has been suggested [19]. Multinuclear complexes, like (CH3Hg)nLþ are formed with O (e.g. (CH3Hg)2OHþ) and S (e.g. (CH3Hg)2S) ligands. Although formation constants are generally large, actual complex concentrations can still be small due to competing reactions. At the one end of the pH spectrum, protons compete with CH3Hgþ for ligands and at the other end, hydroxide ions compete with the ligand for CH3Hgþ. Nevertheless, under environmental and physiological conditions CH3Hgþ complexes will still be dominated by sulfhydryl and sulﬁde interactions (the only exception is in the stomach, where a small amount of CH3HgCl should be formed). Owing to the strong afﬁnity of CH3Hgþ towards sulﬁdic ligands, it can be predicted that virtually all of the methylmercury in natural waters and biological tissue will be bound to such groups (e.g. free sulﬁde in water, cysteine groups in biota). All CH3Hgþ complexes are surprisingly labile. Ligand exchange reactions of CH3Hgþ in aqueous solution are extremely fast, often only limited by the rate of diffusion. Even the thermodynamically very stable glutathione complex easily dissociates and exchanges ligands, particularly with other sulfhydryl groups. Methylmercury species are very stable against chemical attack. Even semiconcentrated acids do not lead to decomposition, as the half-life of CH3HgCl in 1 M H2SO4 is 300 days [20]. Even in hot conc. HNO3 CH3HgCl degrades very slowly. Only strong oxidizing reagents such as dichromate, permanganate and halogens (Cl2, I2 or BrCl) quickly cleave the Hg– C bond. While CH3Hgþ is mostly stable to photo degradation at wavelength . 290 nm, UVa and UVb radiations quickly decompose methylmercury [21]. Microbial degradation of methylmercury is widespread in the environment and is accomplished by the enzyme organomercurial lyase [22], which facilitates oxidative and reductive reaction pathways. CH3HgCl and other uncharged methylmercury species easily penetrate biological membranes and are highly soluble in lipophilic environments. Octanol –water distribution coefﬁcients ðDOW Þ depend greatly on the equilibrium distribution of CH3Hgþ species. However, the calculated partitioning coefﬁcients, KOW ; for chlorides and hydroxides are very small (1.7 and 0.07, respectively) [23]. Those complexes are of little relevance in predicting the fate of methylmercury in the environment as they fail to explain the tremendous biomagniﬁcation potential of methylmercury.

33.3

SAMPLE COLLECTION, PRESERVATION AND STORAGE

This important aspect of every analytical procedure is routinely neglected in method development. While much effort went into reﬁning existing sample preparation techniques or designing new and more sensitive detection systems, very few activities were concerned with the aspects of adequate sample preservation and long-term storage of samples for methylmercury

1065

H. Hintelmann

determination. Our current strategies are mostly based on anecdotal reports or ad hoc investigations prompted by the necessities of an ongoing project rather than the result of systematic studies. Nevertheless, there appears to be general consensus about successful sample handling techniques minimizing losses of individual species concentrations. 33.3.1 Cleaning of sampling and laboratory equipment Glass (quartz or pyrex) and ﬂuoropolymers (“Teﬂon”) are the most appropriate materials for sample collection and storage. Glassware is preferred in our laboratory since Teﬂon is more difﬁcult to clean for repeated use. Species like methylmercury chloride easily diffuse into any polymer material, and will eventually also creep into Teﬂon. However, for ﬁeld work, the lighter Teﬂon has still an edge over breakable glassware. Rigorous cleaning protocols must be followed for sampling and laboratory equipment coming in contact with samples. Procedures used for glass and Teﬂon are similar, except that Teﬂon is not subjected to a BrCl soaking step. Teﬂon bottles designated for methylmercury analysis must never be in contact with such solutions, since BrCl diffuses into Teﬂon and could be released into the next sample causing decomposition of methylmercury. Teﬂon bottles, vials and tubing are rinsed ﬁrst with tap water, then with Milli-Q water and are immersed in (or ﬁlled with) a 50% (v/v) HNO3 solution and heated to 658C for 2 days. This is followed by at least four rinses with MilliQ water and soaking in 10% (v/v) conc. HCl at room temperature for another 3 days. The Teﬂon is rinsed four times with Milli-Q water and all bottles are ﬁlled with 1% low Hg-grade HCl, capped and double bagged in new polyethylene bags for storage. For new bottles and equipment, the initial nitric acid soaking may be omitted. Glassware is ﬁrst rinsed four times with Milli-Q water and then transferred into a container ﬁlled with 10% (v/v) HCl. After overnight soaking at room temperature, the equipment is rinsed four times with Milli-Q water and soaked for another night at room temperature in a BrCl solution (16 l water, 160 ml HCl conc. and 80 ml 0.2N BrCl). Finally, the equipment is rinsed four times with Milli-Q water. Distillation bridges are stored in sealed plastic tubs, while distillation tubes are ﬁlled with 1% low Hg-grade HCl for storage. Quartz ﬁber ﬁlters (QF/F, 47 mm diameter) used to collect particles from water samples are cleaned prior to use by heating overnight in a mufﬂer furnace at 5008C. Individual ﬁlters are stored in precleaned (procedure as for new Teﬂon equipment) polyethylene petri-dishes. 33.3.2 Water sampling Owing to the ultra-low levels of methylmercury in most water bodies, the use of the “clean hand/dirty hand” protocol is essential for contamination-free

1066

Sample preparation for mercury speciation

sampling. It requires two persons, the clean hands conducting the immediate sample collection, whereas the dirty hands assists, i.e., they handle the sample containers up to the inner plastic storage bag. Both the hands must wear protective gloves. All sample containers are stored before and after sampling double-bagged using zip-lock type plastic bags. Surface water samples are collected by submersing the sample bottle below the water surface using armlength gloves. Ideally the bottle is opened below the water surface to avoid sampling of the surface microlayer. Water column proﬁles are obtained by pumping the water through Teﬂon tubing using a peristaltic pump or even better, an impeller driven all-Teﬂon micropump. The inlet line is connected to a solid Teﬂon cylinder, which serves as a sinking weight. Particulate methylmercury can be determined by two ways. First, by measuring the concentration difference between an unﬁltered and ﬁltered water sample. However, this method leads to greater uncertainty, when low concentrations of particulate methylmercury are present. We usually collect particulate methylmercury on a quartz ﬁber ﬁlter by in-line ﬁltration of the water sample using an all-Teﬂon ﬁlter cartridge. This method is beneﬁcial for more accurate measurements. Large volumes of water can be processed to collect a sufﬁcient amount of particles, even when particulate concentrations are low. Before and after ﬁltration, ﬁlters are stored individually in petri dishes and analyzed for methylmercury using the same procedure as for sediments. 33.3.3 Preservation and storage of water samples Earlier work by Leermakers et al. [24] has investigated the storage of methylmercury amended water samples and concluded that both Teﬂon and glass are appropriate materials, as long as storage occurs in the dark at low temperatures. The main route for methylmercury losses was adsorption to the container walls. Plastics such as polypropylene or polyethylene were not suitable leading to quick disappearance of methylmercury from the aqueous solution. However, this investigation was conducted with methylmercury levels 2 – 3 orders of magnitude above typical background concentrations of uncontaminated waters. Extrapolation of conclusions to environmental levels is uncertain, since adsorptive losses are possibly more severe at ultra-low concentrations. There is only anecdotal evidence on the various effects, advantages and disadvantages of adding different mineral acids for preservation of different types of water samples (rain, sea, surface, pore water). Evidently, oxidizing reagents popular in preserving water samples for total mercury analysis are ruled out, since they would quickly decompose methylmercury. Exposure to UVa and UVb must be avoided as well. Although pyrex glass ﬁlters out these wavelengths, photodegradable (.290 nm) CH3HgI and CH3HgSR are potentially present in the sample. Hence storage of water samples in the dark is absolutely necessary.

1067

H. Hintelmann

Common preservation procedures for water samples to be analyzed for methylmercury are acidiﬁcation and storage in the dark at low temperatures (0 –58C) or freezing without adding any preservative. Both methods gave consistent results in our laboratory with storage periods of at least 6 months. However, since it is cumbersome to freeze large numbers of water samples in the ﬁeld followed by shipping to the laboratory on dry ice, we routinely acidify water samples by adding 1% HCl (v/v; i.e., add 1 ml of conc. HCl per 100 ml of sample). The disadvantage of acidiﬁcation lies in the potential coagulation of dissolved organic material (DOM), especially in unﬁltered water samples. Even ﬁltered water with high DOM levels (. 20 mg/l) may be suspected to precipitate DOM, which will cause larger measurement uncertainties. 33.3.4 Preservation and storage of tissue and vegetation samples Whole ﬁsh and other tissue samples are stored frozen to prevent decomposition of methylmercury. No special containers are necessary. Zooplankton and zoobenthos have extremely variable water contents and thus are analyzed best after freeze drying. Vegetation samples are also freeze dried and subsequently homogenized using a coffee grinder that is meticulously cleaned between samples. 33.3.5 Preservation and storage of soil and sediment samples These samples should be frozen immediately after sampling, especially sediments. Mercury methylating and methylmercury demethylating bacteria are ubiquitously present in any sediment. A change in external environmental conditions such as temperature increase and sample mixing will readily change the activity of the microbial communities in each sample, affecting the overall methylation and demethylation rates and consequently the steady state concentration of methylmercury. Only freezing will potentially achieve preservation of the original content. There is no systematic investigation of the effect of drying on resulting methylmercury levels. The outcome may depend on the type of sediments, its texture, organic carbon and total mercury content. Contaminated samples may generate additional methylmercury during drying due to artiﬁcial methylation of inorganic mercury and are better analyzed wet. Wet/dry ratios are determined separately and ﬁnal results are expressed on a dry weight basis. Own experiments with lake sediments have shown no statistical difference between wet, air dried and oven dried (608C) samples, where concentrations of 2.48 ^ 0.39, 2.32 ^ 0.06, and 2.29 ^ 0.01 were determined ðn ¼ 3Þ: If anything, measurements using dried sediments were more precise owing to the difﬁculty of obtaining a representative sample from a wet sediment slurry with high water content.

1068

Sample preparation for mercury speciation

33.4

SAMPLE PREPARATION

Figure 33.1 outlines schematically the main approaches for isolating methylmercury from different matrices. Biological tissues include hair, blood and urine samples, which are frequently analyzed to estimate exposure of humans or wildlife to mercury and its compounds. Depending on the analytical system used for ﬁnal analysis modiﬁcations to the general scheme may be necessary. Solutions obtained by distillation are virtually matrix free and well suited for method involving subsequent aqueous phase derivatization (ethylation [25,26] as well as hydride formation [27]) or SPME sampling [28] techniques. The following procedures are concerned with “true” mercury speciation, focusing on the species speciﬁc determination of methylmercury, which is by far the most dominant and important environmental organic mercury species. The term “methylmercury” is in itself a misnomer, since such a molecule does not exist. Methylmercury refers to all species containing the CH3Hgþ cation. The monovalent ion is usually complexed by a variety of anionic ligands, which change throughout the analytical process. Useful in this context is the parent/matrix/analytical species concept as suggested by Bernhard et al. [29]. Here, the parent species would be CH3Hgþ, which may be present in sea water as the matrix species CH3HgCl and is converted through aqueous phase ethylation into the analytical species CH3HgC2H5. In contrast, the occasionally used term “organic mercury” is only operationally deﬁned and summarizes either all mercury species containing a mercury– carbon bond or refers to inorganic mercury complexed by organic ligands such as humic or fulvic acids. Other operationally deﬁned techniques are designed to extract methylmercury selectively from the sample matrix [30]. The extract is mineralized (digested) and analyzed for total mercury. Assuming that only methylmercury was extracted in the ﬁrst place, this measurement is assumed to be equivalent to the methylmercury concentration in the sample. However, complex matrices such as pore waters, soils or sediments are often too inhomogeneous to ensure 100% discrimination of inorganic mercury (Hg2þ) [31]. A small co-extraction of inorganic Hg2þ into the organic solvent in the form of neutral Hg2þ-complexes such as HgCl2 or Hg-fulvates can greatly change the perceived methylmercury content, particularly in samples having a small percentage of the total Hg present as methylmercury. Hence, such techniques are too uncertain and not even recommended for screening purposes. A review of the chemical/physical characteristics of methylmercury reveals that CH3Hgþ does not exist as the free cation in any sample matrix. Almost all sample preparation methods, therefore, involve acidiﬁcation of the sample to release CH3Hgþ from its binding sites into the dissolved phase. To enhance complete release, another thiophilic metal cation (usually Cu2þ) can be added to outcompete methylmercury in binding to available ligands in the matrix.

1069

H. Hintelmann

Fig. 33.1. General scheme for extracting methylmercury from environmental samples.

This dissolution step is usually accompanied by a derivatization/complexation step to convert CH3Hgþ into a suitable analytical species. Often, halides are used to generate uncharged chloride or bromides, which are easily extracted into an organic solvent or distilled from the sample leachate.

1070

Sample preparation for mercury speciation

33.4.1 Extraction of methylmercury from water Levels of methylmercury in pristine surface waters are usually very low (,100 pg/l) often requiring the processing of large volumes of water and use of preconcentration steps. Classical liquid/liquid extractions into dichloromethane or toluene after acidiﬁcation have been developed [25,32]. Typically, this method is used in conjunction with the aqueous phase ethylation technique [25], requiring the transfer of methylmercury from the organic phase into aqueous solution, which is accomplished by adding the organic solvent to an equal volume of water and subsequent evaporation of the organic solvent. Other techniques have tried some form of solid phase extraction using custommade adsorbents employing sulﬁde functional groups or chelating reagents. Most popular for the extraction of methylmercury from water is modiﬁed sulfhydryl cotton ﬁber [33,34]. However, recent reports suggest that this material may generate artiﬁcial methylmercury during the preconcentration/ elution steps [35]. An extremely sensitive technique to isolate methyl mercury was developed around the observation that methylmercury halides are volatile and can be distilled from almost any sample using vacuum or steam distillation [27,36]. The landmark papers by Horvat [4,9] discuss the nitrogen assisted distillation technique in detail. The procedure separates methylmercury from its matrix very efﬁciently resulting in a clean distillate containing only methylmercury. The method is particularly suitable for organic rich water samples, which are difﬁcult to analyze with other techniques. We have modiﬁed the original procedure by using an all-glass distillation apparatus shown in Fig. 33.2. It consists of two 50 ml glass tubes connected via an air cooled glass distillation bridge (i.d. 8 mm). Fifty grams of water is weighed into each distillation vial and 500 ml of H2SO4 (9 M) and 200 ml of KCl (20%, w/v) are added to each sample. At this stage, isotope-enriched methylmercury chloride may be added for isotope dilution quantiﬁcation. 5 ml of Milli-Q water are placed into the receiving vial. Both vials are connected to the distillation bridge and the set-up is placed into an aluminum heating block set to 1408C. Hg-free N2 gas (60–80 ml/min, controlled by individual ﬂow meters) is bubbled through each sample to hasten the distillation process. Approximately 85% (42.5 ml) of the sample are distilled within 3 h. Samples can be stored in the dark at 48C in the receiving vials without additional preserving agents for up to 1 week. Compared with Teﬂon, the use of glass equipment has several advantages. It allows the cleaning of all distillation ware with BrCl, which is very efﬁcient in removing methylmercury from surfaces. This resulted in markedly lower absolute procedural blanks of routinely less than 5 pg (often below 1 pg). The use of BrCl is not recommended for cleaning Teﬂon ware that is used for methylmercury determinations. Secondly, glass conducts heat much better than plastic resulting in more than halving the time required for distilling a 50 ml water sample.

1071

H. Hintelmann

Fig. 33.2. Glass distillation apparatus used for distilling methylmercury from water samples.

33.4.2 Extraction of methylmercury from soils, sediments and particles The isolation of methylmercury from soils and sediments is still a formidable challenge in analytical chemistry. Methylmercury usually does not exceed ,1% of the total Hg in these samples and it is very difﬁcult to validate that the measured concentration is identical to the natural in situ level. Sample treatment typically involves the addition of acid or base, which potentially disturbs the established steady state concentration and may change the methylmercury level. Unfortunately, current reference materials cannot aid in this aspect of method validation, since CRMs are typically dried, sterilized materials, and are therefore, only an approximation for real sediments. The acidic leaching procedure was originally developed by Japanese and Scandinavian researchers [37,38]. Numerous variations of the principal method have been reported since. In general, a sulfuric acid/copper bromide mixture is added to the sample, followed by extraction of the formed methylmercury halide into an organic solvent such as toluene or dichlorobenzene. A further clean-up of the extract is achieved by backextracting methylmercury into an aqueous cysteine or thiosulfate solution. These ligands form more stable complexes with CH3Hgþ than the chloride or bromide

1072

Sample preparation for mercury speciation

complex established during the acid treatment. CH3HgX are non-polar molecules and prefer the organic phase. The cysteine and thiosulfate complexes are either zwitterionic or anionic species in the pH range from 5 to 7 and partition readily into the aqueous phase. In case, gas chromatography is used as the analytical system, ﬁnal transfer into an organic solvent is accomplished by acidiﬁcation with a halogenic acid, which protonates the cysteine (or thiosulfate) ligand. The liberated MeHgX is extracted into an organic solvent, which can be injected onto the GC column. This double back-extraction technique is essential to remove many interferences (e.g. thiols), which may either interfere with unspeciﬁc detection (e.g. electron capture) or otherwise lead to rapid deterioration of the analytical column. Recently, acidic extractions have been improved by developing microwave assisted methods, which generally shortens the required extraction time [39]. The general difﬁculty with liquid/liquid extractions are variable extraction yields and frequent formation of persistent emulsions, which are a challenge to separate. Hence, a distillation procedure similar to the one described for water samples is employed and results in cleaner, matrix-free sample solutions for further analysis. To speed up the distillation step, we use a smaller Teﬂon distillation apparatus for these samples. Blanks obtained with Teﬂon equipment are often slightly larger and more variable compared to the ones obtained with glass stills. However, this is usually acceptable, since methylmercury levels in sediment and soil samples are much higher than in water. Approximately 1 g of wet sediment (soil) or a whole ﬁlter paper with collected particles is placed into a 25 ml Teﬂon vial. Ten milliliters of Milli-Q water, 500 ml of H2SO4 (9 M), 200 ml of KCl (20%, w/v) and isotopically enriched methylmercury are added to each sample. Subsequent steps are identical to distilling water samples. Again, 85 –90% of the sample is distilled, which should take about 90 min. 33.4.3 Extraction of methylmercury from biological tissue A popular technique to extract methylmercury from biological samples involves alkaline digestion of the matrix. A solution of 20 % KOH in methanol (w/v) is used to solubilize the tissue sample. After neutralization with halogenic acid, mercury halides are extracted into an organic solvent and processing continues as described for sediment samples. However, this variation often results in severe emulsion formation. Alternatively, an aliquot of the alkaline solution can be processed for aqueous phase ethylation. This works well for ﬁsh and similar samples having high methylmercury levels requiring only small sample aliquots (#50 ml), which can be added directly to the reaction vessel. When the available sample mass is small or the methylmercury concentration is low, larger volumes of digest must be processed. Prime examples are the measurement of methylmercury in zooplankton and -benthos, which are more

1073

H. Hintelmann

challenging. Own investigations have recently shown that methylmercury in larger aliquots of the alkaline digest (. 100 ml) is not quantitatively derivatized. Apparently, the organic matrix is not completely decomposed and interferes with the ethylation step, which must be conducted at pH 4.5– 5. This is similar to the observation made by other researchers processing directly untreated water and sediment samples [40]. We recently developed an acid digestion using HNO3 followed by aqueous phase ethylation, overcoming any matrix effect [41]. A 1 –10 mg subsample of dried biological tissue material are weighed into a 5 ml conical Teﬂon vial. Diluted HNO3 (5 ml; 4 M) and isotopically enriched methylmercury are added to the sample. The vial is tightly capped and heated at 508C for 24 h. The resulting digest is neutralized using KOH (20%, w/v) and further analyzed by aqueous phase ethylation. Using this method, it was possible to process the whole 5 ml digest without any loss in derivatization efﬁciency. 33.4.4 Direct techniques involving no sample preparation The ideal analytical method would employ no sample preparation at all and transfer the analyte from the sample directly into the analytical system. Unfortunately, such direct methods are rare. Attempts have been made to derivatize methylmercury directly in water samples and even sediments. However, careful evaluation has shown that such techniques are prone to incomplete conversion owing to matrix interferences, particularly in sediments [40]. It is well accepted now that a separation of methylmercury from complex matrices is essential. A potential exception is homogenous sample matrices such as ﬁsh muscle tissues. A technique developed by Lansens and Baeyens [42] used iodoacetic acid to form MeHgI, which was collected from the sample headspace. Another technique employed an electrothermal vaporization of MeHgCl from ﬁsh samples [43]. However, both the methods are limited so far to samples having high concentrations of methylmercury and are not applicable to more complex matrices such as sediments. 33.4.5 Extraction of mercury species other than methylmercury As mentioned before, methylmercury is, by far, the most important mercury species in the environment. It is rarely released into the environment but generated in situ by biotic and abiotic methylation reactions. The only other organic mercury compound that occurs naturally is dimethylmercury. It should be noted that the commonly used procedures for methylmercury analysis involving acidiﬁcation of the sample would almost certainly destroy dimethylmercury very quickly. As well, distillation techniques are not able to capture dimethylmercury. If this species is to be analyzed, it must be collected ﬁrst by purging it from the original untreated sample and trapped on a solid adsorbent.

1074

Sample preparation for mercury speciation

Sporadic screenings never revealed the presence of dimethylmercury in ordinary samples of any kind. This species was detected occasionally in deep sea water [5], and in the air above highly contaminated ﬂood plains [44]. It probably originates from bacterial processes or dismutation of (CH3Hg)2S [45]. No natural formation pathways have been suggested for other organomercurials and it must be concluded that the occurrence of those species is related to anthropogenic release into the environment. Ethylmercury had been detected as a result of industrial spills [46,47]. The observation of ethylmercury in the Florida Everglades [48] is controversial since no explanation for the occurrence could be found. Alkylethoxy- and phenylmercury species have been used as fungicides in the past, but are highly unstable under environmental conditions. If emitted, concentrations will quickly decrease below detectable levels. A mild extraction method employing solubilization with citrate at pH 4 and extraction with dithizone in chloroform was developed [49] to isolate exotic organomercury species from soils. A series of unusual species was later detected in a contaminated industrial setting [50]. 33.5

QUALITY CONTROL

It is most difﬁcult to control the accuracy of results for methylmercury in water samples at pg/l levels. Stringent QA/QC protocols must be implemented to ensure consistent results in routine analysis. Modern detection techniques are usually sensitive enough to detect even smallest amounts of methylmercury and limits of detection often depend on the magnitude and variability of the procedural blank. Since reagents and laboratory environments are normally not contaminated with methylmercury, the largest contribution to the overall blank originates from the sampling devices, and other equipment used during sample manipulation and preparation. This is why rigorous cleaning of all sampling and laboratory ware is paramount. It is also important to note that at least 20% of all methylmercury determinations are QA/QC samples including blanks and certiﬁed reference materials (for samples other than water). 33.5.1 Artifactual formation of methylmercury Recent discoveries have indicated that artifact methylmercury may be generated during sample preparation procedures [51,52]. A detailed discussion on sources of error in methylmercury determination during sample preparation, derivatization and detection is reported in a special journal issue summarizing the ﬁndings of an expert’s workshop on this topic [53]. Contaminated soils and sediments with high levels of total mercury and very small proportion of methylmercury (, 0.1% of HgT) are especially prone to artifactual formation of MeHg using distillation and other techniques. If even a

1075

H. Hintelmann

small amount of the inorganic mercury was converted to methylmercury during analysis, the measured methylmercury would be signiﬁcantly affected because of the large ratio of inorganic to methylmercury in those samples. A comparison of different sample preparation procedures concluded that extraction techniques rarely form artifacts in high level Hg samples [32]. Pristine samples with background concentrations of total mercury usually do not show detectable levels of artifactual methylmercury formation, even if distillation techniques are applied. Very recent discoveries have shown evidence of methylmercury generation when extracting Hg species from large volumes of water using sulfhydryl-cotton ﬁber [35]. The work concluded that reagents used to derivatize the cotton may be responsible for methylating inorganic mercury during the preconcentration process. Clearly, these ﬁndings require immediate attention. In general, it may be possible to use stable isotopes of mercury to quantify the extent of artifactual mercury formation during sample preparation. The technique proposed by Hintelmann et al. [51] add a known amount of enriched Hg2þ to the sample prior to sample preparation and measures the fraction of methylmercury formed from this stable isotope. The observed artifactual conversion rate is applied to the native mercury to correct the apparent concentration of methylmercury. The procedure is valid as long as the added inorganic isotope tracer fully equilibrates with the native inorganic mercury in the sample, a prerequisite similar to standard addition and isotope dilution techniques, which are widely used to quantify total metal and element species concentrations. 33.5.2 Spike recoveries Incomplete and variable extraction/distillation yields are the perennial problems with all methods developed for methylmercury isolation. Consequently, isolation yields must be determined for each individual sample to obtain the accurate methylmercury values of the original sample. Since standard addition techniques are time consuming, particularly in combination with tedious methylmercury determinations, recoveries were often only determined once for a single sample and the value obtained was applied to all samples of the same batch. Inevitably, this approach invites considerable uncertainty, particularly with inhomogeneous samples, where yields can vary signiﬁcantly from sample to sample. The use of enriched stable isotopes allows a convenient assessment of individual recoveries without any additional sample preparation. However, by its very nature this procedure requires the use of ICP/MS detection rather than the more common and less expensive atomic ﬂuorescence detection. A known amount of CH201 3 HgCl is added to the sample prior to the isolation step as an internal standard. Using the concepts described in Hintelmann and Ogrinc þ [54], we can determine the amount of CH201 recovered and correct the 3 Hg

1076

Sample preparation for mercury speciation

Fig. 33.3. Distillation recoveries for methylmercury from different lake sediments. The ﬁrst two series are replicate distillations from sediment slurries with low and high organic carbon content, respectively. The following three series represent individual samples from lake sediment cores.

concentration of all other isotopes in the sample accordingly. Of course, this approach assumes that the added spike equilibrates during the sample preparation procedure rendering its behavior indistinguishable from the native methylmercury originally present in the sample. However, conventional standard addition methods are based on the same assumption. Fig. 33.3 shows results of individual recoveries for a series of distillations from sediment cores and replicate distillations of the same sediment slurry. Although average recoveries (85 ^ 10%) are similar to recoveries typically reported by other researchers, it is clear that individual yields can vary greatly. In particular, single outliers in a batch may easily go undetected, if no individual distillation yields are determined. 33.5.3 Reference materials Certiﬁed reference materials (CRM) are an indispensable tool for method development, method validation and ongoing quality control programs. Table 33.2 lists currently available reference materials that are certiﬁed for methylmercury. At least one CRM should be prepared and analyzed for every batch of 10 samples. Ideally a reference material having similar matrix and comparable methylmercury concentration should be used. A look at Table 33.2 shows that a variety of biological tissues are readily available. However, a number of important matrices are still missing in this list, most

1077

H. Hintelmann TABLE 33.2 Available reference materials certiﬁed for methylmercury Supplier

Name

Description

NRCa

DORM-2 DOLT-3 LUTS-1

Fish muscle tissue Fish liver tissue Lobster tissue

TORT-2

Lobster tissue

SRM 2977 SRM 1974a

Mussel tissue Mussel tissue

SRM 2976 SRM 1566b

Mussel tissue Oyster tissue

IAEAf

IAEA-140/TM IAEA-085 IAEA-086 IAEA-405

IRMMg

NIESh

NIST

e

Methymercury (ng/g)

Total mercury (ng/g)

4470 ^ 320 1700b 9.4 ^ 0.6c 63 ^ 4d 152 ^ 13

4640 ^ 260 3700 ^ 140 16.7 ^ 2.23 112 ^ 1.5d 270 ^ 60

36.2 ^ 1.7 8.80 ^ 0.35c 77.3 ^ 3.1d 27.8 ^ 1.1 13.2 ^ 0.7

101 ^ 4b 20.1 ^ 1.5c 176 ^ 13d 61.0 ^ 3.6 37.1 ^ 1.3

Seaweed Human hair (spiked) Human hair Estuarine sediment

0.626 ^ 0.107 22,900 ^ 1000 258 ^ 22 5.49 ^ 0.55

38 ^ 6 23,200 ^ 800 573 ^ 39 810 ^ 40

CRM-463 CRM-464 CRM-580 CRM-422

Tuna ﬁsh Tuna ﬁsh Estuarine sediment Mussel tissue

3040 ^ 160 5500 ^ 1700 75.5 ^ 3.7 0.43b

2850 ^ 160 5240 ^ 100 13,2000 ^ 3000

CRM-13

Human hair

3800 ^ 400

4420 ^ 220

a

National Research Council, Canada—Institute for National Measurement Standards. Informational value only. Based on wet weight. d Based on dry weight. e National Institute of Standards & Technology, USA. f International Atomic Energy Agency—Analytical Quality Control Services (AQCS). g Institute for Reference Materials and Measurements, European Community. h National Institute of Environmental Studies, Japan. b c

notable—water, vegetation and uncontaminated sediments. As a substitute, inhouse prepared samples can be used as a laboratory reference material for training purposes and control charts to ensure that methods are under control. Additionally, interlaboratory comparisons are a valuable quality control instrument, particularly for matrices that are not available as CRM or RM. In fact, interlaboratory comparisons are the only independent avenue to ensure the comparability and hopefully validity of values obtained by different research groups.

1078

Sample preparation for mercury speciation

ACKNOWLEDGEMENT The author wishes to thank Brina Dimock for his sketch of the distillation apparatus. REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29

30 31

Mercury report to congress, ed. EPA. Vol. 452/R-97-0003, 1997. W.L. Clevenger, B.W. Smith and J.D. Winefordner, Crit. Rev. Anal. Chem., 27 (1997) 1. C. Brosset and E. Lord, Water Air Soil Pollut., 82 (1995) 739. M. Horvat, L. Liang and N.S. Bloom, Anal. Chim. Acta, 282 (1993) 153. R.P. Mason and W.F. Fitzgerald, Nature, 347 (1990) 457. V. St. Louis, J. Rudd, C. Kelly, B. Hall, K. Rolfhus, K. Scott, S. Lindberg and W. Dong, Environ. Sci. Technol., 35 (2001) 3089. M. Abbott, D. Susong, D. Krabbenhoft and A. Rood, Water Air Soil Pollut., 139 (2002) 95. D. Schwesig and E. Matzner, Sci. Total Environ., 260 (2000) 213. M. Horvat, N.S. Bloom and L. Liang, Anal. Chim. Acta, 281 (1993) 135. Ontario Ministry of the Environment, Guide to Eating Ontario Sport Fish, 19th edn. Ontario Ministry of the Environment, 1997. M.J. Paterson, J.W.M. Rudd and V. St. Louis, Environ. Sci. Technol., 32 (1998) 3868. A. Tremblay, L. Cloutier and M. Lucotte, Sci. Total Environ., 219 (1998) 209. A. Kirkwood, P. Chow-Fraser and G. Mierle, Environ. Toxicol. Chem., 18 (1999) 523. R. Wagemann, E. Trebacz, G. Boila and W.L. Lockhart, Sci. Tot. Environ., 218 (1998) 19. R.D. Evans, E.M. Addison, J.Y. Villeneuve, K.S. MacDonald and D.G. Joachim, Environ. Res., 84 (2000) 133. R. Dietz, C. Nielsen, M. Hansen and C. Hansen, Sci. Tot. Environ., 95 (1990) 41. J.H.R. Clarke and L.A. Woodward, Trans. Faraday Soc., 64 (1968) 1041. P.L. Goggin and L.A. Woodward, Trans. Faraday Soc., 58 (1962) 1495. G. Schwarzenbach and M. Schellenberg, Helv. Chim. Acta, 48 (1965) 28. M.M. Kreevoy, J. Am. Chem. Soc., 79 (1957) 5927. J.A. Tossel, J. Phys. Chem., 102 (1998) 3587. J.B. Robinson and O.H. Tuovinen, Microbiol. Rev., 48 (1984) 95. C. Faust Bruce, Environ. Toxicol. Chem., 11 (1992) 1373. M. Leermakers, P. Lansens and W. Baeyens, Fresenius J. Anal. Chem., 336 (1990) 655. N. Bloom, Can. J. Fish. Aquat. Sci., 46 (1989) 1131. S. Rapsomanikis, O.F.X. Donard and J.H. Weber, Anal. Chem., 58 (1986) 35. M.A. Morrison and J.H. Weber, Appl. Organomet. Chem., 11 (1997) 791. Z.N. Mester, J. Lam, R. Sturgeon and J. Pawliszyn, J. Anal. At. Spectrom., 15 (2000) 837. M. Bernhard, F.E. Brinckman and K.J. Irgolic, Why ‘Speciation’. In: M. Bernhard, F.E. Brinckman and P.J. Sadler (Eds.), The Importance of Chemical ‘Speciation’ in Environmental Processes. Springer, Berlin, 1986, p. 7. K. May, M. Stoeppler and K. Reisinger, Toxicol. Environ. Chem., 13 (1987) 153. M. Horvat, K. May, M. Stoeppler and A.R. Byrne, Appl. Organomet. Chem., 2 (1988) 515.

1079

H. Hintelmann 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46

47 48 49 50 51 52 53 54

1080

H. Hintelmann, Chemosphere, 39 (1999) 1093. Y. Cai, R. Jaffe, A. Alli and R.D. Jones, Anal. Chim. Acta, 334 (1996) 251. Y.H. Lee, Int. J. Environ. Anal. Chem., 29 (1987) 263. V. Celo, R.V. Annath, D.R.S. Lean and S.L. Scott. Third general Meeting of COMERN. Elmhirst Resort, Keene, Ont., Canada, 2002. M. Floyd and L.E. Sommers, Anal. Lett., 8 (1975) 525. K. Sumino, Kobe J. Med. Sci., 14 (1968) 115. G. Westoeoe, Acta Chem. Scand., 20 (1966) 2131. C.M. Tseng, A.d. Diego, F. Martin and O. Donard, J. Anal. At. Spectrom., 12 (1997) 629. R. Reuther, L. Jaeger and B. Allard, Anal. Chim. Acta, 394 (1999) 259. H.T. Nguyen and H. Hintelmann, Anal. Chim. Acta, (2003) in press. P. Lansens and W. Baeyens, Anal. Chim. Acta, 228 (1990) 93. I. Gelaude, R. Dams, M. Resano, F. Vanhaecke and L. Moens, Anal. Chem., 74 (2002) 3833. D. Wallschla¨ger, H. Hintelmann, R.D. Evans and R.-D. Wilken, Water Air Soil Pollut., 80 (1995) 1325. P.J. Craig and P.A. Moreton, Mar. Pollut. Bull., 15 (1984) 406. A. Jerneloev and G. Wennergren, Studies of Concentrations of Methyl Mercury in Sediments from the St. Clair System and Rate of Biological Methylation in Incubated Samples of Sediments. Swedish Water and Air Pollution Research Institute, Stockholm, 1980, p. 1. H. Hintelmann and R.-D. Wilken, Appl. Organomet. Chem., 7 (1992) 1. Y. Cai, R. Jaffe and R. Jones, Environ. Sci. Technol., 31 (1997) 302. M. Hempel, H. Hintelmann and R.-D. Wilken, Analyst, 117 (1992) 669. H. Hintelmann, M. Hempel and R.-D. Wilken, Environ. Sci. Technol., 29 (1995) 1845. H. Hintelmann, R. Falter, G. Ilgen and R.D. Evans, Fresenius J. Anal. Chem., 358 (1997) 363. N.S. Bloom, J.A. Colman and L. Barber, Fresenius J. Anal. Chem., 358 (1997) 371. R. Falter, H. Hintelmann and P. Quevauviller, Chemosphere, 39 (1999) 1039. H. Hintelmann and N. Ogrinc, Determination of stable mercury isotopes by ICP/MS and their application in environmental studies. In: Y. Cai (Ed.), Biogeochemistry of Environmentally Important Trace Elements. American Chemical Society, 2002.

Chapter 34

Sample preparation for speciation of lead Freddy C. Adams and Monika Heisterkamp

34.1

INTRODUCTION

Tetraalkyllead (TeAL) compounds are used worldwide as anti-knock agents in leaded fuel to prevent premature ignition of the compressed air–gasoline mixture, which is detected as knocking of the motor. During the ﬁring process, the majority of the lead–carbon bonds are broken and the organolead is converted into inorganic lead halides that leave the engine via the exhaust pipe. Depending on the motor type and the velocity, a small fraction of organolead (,2%) enters the atmosphere because of incomplete combustion. In view of the enormous consumption of leaded gasoline, this is by far the most important source of organolead in the atmosphere. TeAL is also released into the environment by evaporation during production, transport and storage of both anti-knock agents and leaded gasoline. Distribution of the latter at petrol stations is also a considerable source due to liberation of the volatile organolead compounds during the tanking process. Once released into the atmosphere, these TeAL compounds can be transported over great distances as free molecules or adsorbed on particles, so that they can even be detected in remote areas, such as Greenland or Antarctica [1]. These species (tetramethyllead [TeML], tetraethyllead [TeEL] or mixed tetra-ethylmethyllead) decompose via their ionic trialkyllead (TAL: trimethyllead [TML] and triethyllead [TEL]), dialkyllead (DAL: dimethyllead [DML] and diethyllead [DEL]) into inorganic lead (Pbi). The breakdown reaction is initiated by reactive species such as hydroxyl radicals generated by photochemical activity or by direct photolysis. The ionic organolead is washed out of the atmosphere by precipitation and transported to the hydrosphere. Sunlight and the components present in the aqueous medium determine the rate of the decomposition, as many metals catalyze this process. Table 34.1 presents a survey of the compounds on which this section is mainly focused. The question of whether methyllead is also formed by biological methylation of inorganic lead is still controversial. At this time, it is not clear Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

1081

1082 Environmentally important organolead compounds TeAL TAL DAL Monoalkyl lead ? Inorganic lead

(CH3)4Pb tetramethyllead (CH3)3Pbþ trimethyllead, TML (CH3)2Pbþ þ dimethyllead, DML CH3Pbþ þ þ monomethyllead

(CH3)3C2H5Pb trimethylethyllead CH3)2C2H5Pbþ dimethylethyllead CH3C2H5Pbþ þ methylethyllead C2H5Pbþ þ þ monoethyllead

(CH3)2C2H5)2Pb dimethyldiethyllead (CH3)(C2H5)2Pbþ methyldiethyllead (C2H5)2Pbþ þ diethyllead, DEL

(CH3)(C2H5)3Pb methyltriethyllead (C2H5)3Pbþ triethyllead, TEL

(CH3)4Pb tetraethyllead

F. C. Adams and M. Heisterkamp

TABLE 34.1

Sample preparation for speciation of lead

whether methylation of lead in the aquatic environment is a biological process or just an abiotic reaction or a combination of both. A two-stage mechanism, starting with oxidative addition of a carbonium ion to Pbi, primarily via a biotic process, was proposed by Walton et al. [2], but such a methylation can also be performed via an abiotic route [3]. TML, DML and TeML are generated in a second step by abiotic reductive disproportionation and dismutation reactions. Because of the low concentration of methylating agents, such as methyliodide, which seems to be too low to produce any detectable amount of methyllead under ﬁeld conditions, methylation via a slow biotic process seems to be favored. This assumption is underpinned by results reported by Pongratz and Heumann, who proved the production of methyllead species in arctic regions by macroalgae [4] and marine bacteria [5]. Only TML and no TeML or DML was found. The macroalgae were collected in the Kongsford in Spitsbergen (Norway) and cultivated in incubation vessels under polar conditions. Growth, as well as the irradiation conditions inﬂuence the release rates for the different macroalgae. Depth proﬁles of the TML concentration in the polar seawater show that, at least at deeper water levels, bacteria must be the main species contributing to biological methylation [5]. Nevertheless, due to the enormous release of organolead into the atmosphere caused by automobile emissions, the fraction of naturally formed organolead is considered negligible. A schematic illustration of the sources and environmental pathways for organolead is given in Fig. 34.1 [6]. 34.2

TOXICITY OF ORGANOLEAD COMPOUNDS

Lead is toxic in all forms, but to varying degrees, depending on the species in which it occurs. The toxicity of organolead compounds increases with the number of alkyl groups bound to one lead atom and differs considerably from those of elemental or inorganic lead. Due to their higher liposolubility, organolead compounds are able to penetrate biological membranes, which makes them generally more toxic than inorganic lead compounds. Pathways for TeAL uptake include inhalation, due to the high volatility, or adsorption through the intact skin, due to the capability of organolead species to penetrate membranes, with TeML to a lesser extent than TeEL. The rate of uptake depends on skin thickness, vascularity and temperature for dermal adsorption and on the dose and time of exposure for pulmonary absorption. Another uptake path for TAL and other ionic organolead in the body is via the food chain because of bioaccumulation of ionic organolead in marine organisms or birds, for example. Concentrations up to 25 ng g21 TEL were found in kidneys of urban pigeons [7]. Inside the body, TeAL is cleared within a few minutes from the blood by degradation to TAL, DAL and subsequently inorganic lead. While TAL is quite stable and can remain in the body for extended time, DAL is rapidly excreted via the feces.

1083

1084 F. C. Adams and M. Heisterkamp

Fig. 34.1. Sources and pathways for entry of organolead into the environment [6].

Sample preparation for speciation of lead

The toxic potential of organolead compounds is generally high, but varies according to the chemical structure and differs among the various target organisms. To a large extent, these differences are caused by different rates of absorption, transformation, distribution in, and elimination from, the body. Normally, DAL is less toxic than TAL and TeAL due to its shorter lifetime in the body. Whereas for ﬁsh, TeAL compounds show a higher toxicity due to the easy uptake via the gills, for most mammals TAL compounds are more toxic. The dealkylation process for methyl- and ethyllead seems to be speciﬁc for different target species; dogs and mice are more sensitive to methyllead, while ethyllead is more toxic for rats and humans. The latter is indicated by the estimated LD50 value for humans, which is, extrapolated from data for rats; 250 mg kg21 body weight for TeEL and more than 1000 mg kg21 for TeML [8]. Acute exposure to high doses of TeAL does not immediately result in the development of symptoms. Animal experiments with TeAL and TAL suggest that TAL formed by dealkylation processes in the body, results in the toxicity of TeAL. The necessary metabolism for the degradation, which mainly takes place in the liver, and the following distribution of TAL to target sites, predominantly the brain, may explain the relatively long latency period. The symptoms of acute organolead poisoning can arise with a delay of up to 3 days and the toxic effects are insomnia, depression and peculiar hallucinations. Furthermore, tremor and vegetative disturbances, such as hypotension can occur leading to paranoid states, unconsciousness and ﬁnally death. Under chronic exposure conditions, recognizable symptoms may not appear for a long time, up to several years, but symptoms are similar to those following acute exposure. TeAL can also initiate cancer and neurological diseases [9]. The brain is considered the critical organ in organolead intoxication, but most lead accumulates in the liver and kidneys and scarcely in the bones. TAL can pass the blood– brain barrier and exert neurotoxic effects. It is supposed that TAL increases the anion permeability across cell and mitochondrial membranes. The hydroxide and chloride forms of TAL are sufﬁciently lipophilic to pass through biological membranes. The organolead cations are capable of acting as “ferries” for these anions, disturbing the concentration gradient of chloride anions across the membrane. As a result, certain neurons show an enhanced excitability, leading to an increased activity of these cells. This triggers an enhancement in glucose utilization, exceeding the maximum capacities of adenosine triphosphate, which is generated during the oxidative phosphorylation in mitochondria. Furthermore, TAL compounds also restrain the phosphorylation process so that an insufﬁcient supply of the cells together with their elevated activity causes localized cell death [10]. 34.3

THE HISTORY OF LEADED GASOLINE

For a long time, the composition of fuel was the bottleneck in the development of high efﬁciency internal combustion engines due to its insufﬁcient

1085

F. C. Adams and M. Heisterkamp

compression ratio. The fuel furthest from the spark plug, known as the end gas, is subject to compression and heating for a longer time than in any other part of the engine. This mixture is brought to an abnormal state of partial oxidation, which tends to give rise to a spontaneous, untimely detonation. This knocking in the motor leads to loss of power, overheating and subsequently to damage of the pistons. Much research effort was spent on compounds able to suppress this knocking of the fuel. General Motors checked more than 33,000 compounds over a period of 6 years, before stumbling over the anti-knocking properties of TeEL in 1921 [11,12]. This invention was the origin of the growth in the automotive industry, because using leaded gasoline permitting a higher compression ratio allowed much more effective engines. The industrial production of TeEL started using the Kraus-Callis process in 1923 and can be described by the following reaction: 4PbNa þ 4Ch3 CH2 Cl ! 3Pb þ 4NaCl þ ðCH3 CH2 Þ4Pb

ð34:1Þ

By melting lead and sodium, a highly reactive lead –sodium alloy is produced which is then autoclaved with ethyl chloride. The Ethyl Corporation was established in 1924 to produce and market leaded gasoline on an industrial scale. In 1928, the drawback of deposition of lead oxides on the exhaust valves, spark plugs and combustion chamber could be overcome by adding scavengers, such as dichloroethane, to the fuel to convert the oxides into more volatile halides. With these improvements, the way was paved for automobile mobility all over the world, making organolead one of the organic compounds produced in the highest bulk quantities. An average addition of 0.52 g l21 (as Pb) of TeAL led to a yearly consumption of these compounds exceeding 375,000 t (as Pb) worldwide in 1970, corresponding to more than 720 billion liters of leaded gasoline [13]. The suspicion that the use of organolead as a gasoline additive might be a potential threat to public health arose immediately after the anti-knocking properties were discovered. Five laboratory personnel of an oil company died in 1924 and 35 others were seriously injured within 4 days and some US states banned the production of leaded gasoline after this tragedy [14]. The prohibition was only temporary, because the industrial lobby convinced the authorities by giving reassuring statements that the use of organolead as anti-knocking agent was a milestone in industrial progress. In 1925, a conference was held in the Unites States to assess the toxicity of organolead. Industry won the ﬁght [16] and it was concluded that it was not necessary to ban organolead in gasoline provided that production, distribution and sales were properly regulated [15]. The menace of organolead in gasoline causing severe problems for public health remained fundamentally overlooked and unappreciated for more than 30 years. In the 1960s, overwhelming evidence appeared that the average concentration of lead in both the environment and human bodies had increased

1086

Sample preparation for speciation of lead

signiﬁcantly. The determination of the isotopic composition of lead additives and aerosols showed similar patterns, proving leaded gasoline to be the largest source of atmospheric lead pollution. Not only urban inhabitants, but also those in rural areas, showed increased lead concentrations in blood due to long-term exposure to environmental lead. A screening program in the United States made clear that the content of lead in children’s blood tended to increase gradually with the level of occupational exposure to automobile exhaust. Hence, it was realized that lead additives have become a health peril affecting the average citizen nationwide. As a consequence, the debate on the widespread use of leaded fuel arose again with the same fervor as in the 1920s, but the overall impact on both the environment and human health could no longer be glossed over or denied. Ironically, the ﬁrst legislative restrictions on the use of leaded fuel in the United States were not due to environmental lead pollution but to smog problems in Los Angeles. The Clean Air Act was enacted in 1970 and enabled the authorities to control or prohibit any fuel or fuel additives that contribute to air pollution leading to severe problems for public health. Since 1975, all new cars in the United States had to be equipped with catalytic converters and fuel stations had to distribute at least one grade of unleaded gasoline due to these lead-intolerant converters. Furthermore, the average lead level of fuel was reduced from 0.52 g l21 (as Pb) to 0.28 g l21 in 1982 and to a “lead free” level of 0.026 g l21 (as Pb) in 1989. The substitution of lead additives in Europe started in 1986 and the maximum content of alkyllead in fuel has been restricted to 0.15 g l21 since 1989 [13]. From January 2000, leaded gasoline was phased out in the countries of the EU and only unleaded fuel containing less than 0.026 g l21 (as Pb) is available at petrol stations [15]. Approximately 20 years after the recognition of the menace of organolead in fuel on the environment and human health, adequate legislation concerning automobile emissions was enacted. Due to these regulations, the concentrations of lead in the environment decreased with the decline in the production of lead additives. Although the distribution of leaded gasoline is now banned, or at least restricted, in more or less all countries of the Western World, only few attempts were made to curtail the use of lead additives in developing countries where the consumption of organolead is still expanding. The world’s largest users of leaded gasoline nowadays are countries where both the gasoline consumption and the concentration of TeAL as anti-knock agent in the fuel are high. In the late 1990s, these countries were, for example, Nigeria, having an average lead level in gasoline of 0.6 g l21, Indonesia with 0.44 g l21 and Saudi Arabia with 0.40 g l21 (all concentrations expressed as Pb) [16]. A recent study clearly correlated the use of TeAL as anti-knock agents in leaded gasoline and blood lead levels of the populations of various countries on different continents [17]. The blood lead concentrations decreased in Belgium, for example, from 17 mg l21 in 1979 to 15 mg l21 in 1985 to 9 mg l21 in 1987. The lead concentration in gasoline decreased over this same period from 0.45 to

1087

F. C. Adams and M. Heisterkamp

0.4 to 0.15 g l21 (as Pb) [18]. The most remarkable point of this study was the convergence of blood lead levels across locations and time as gasoline lead concentrations were progressively banned. This convergence indicated that the remaining sources of lead exposure at these locations contribute no more than about 3 mg l21 to the average lead exposure of the general population with the phase-out of leaded gasoline being the key contributor to decreased concentrations in human blood. Starting from 1970s, considerable work has been done to identify and quantify alkyllead compounds in environmental matrices in an effort to establish a quantitative model of their environmental pathways. Many of the earlier methods suffered from two general drawbacks: a rather poor speciﬁcity, largely due to an inability to discriminate unambiguously between inorganic lead and (some of the) organic lead species, and a limited sensitivity, allowing measurements only in highly contaminated areas. Evaluation of accidental pollution, in particular, such as that of the Cavtat shipwreck [19], the urgent need for reliable detection of the different TeAL, DAL and TAL species and their degradation products arose. 34.4

PROPERTIES OF ORGANOLEAD COMPOUNDS

Organometallic compounds are fundamentally different in both chemical and biological properties from ionic compounds of the same metal. The availability to organisms and the toxicity of a heavy metal differs considerably from one chemical species of the metal to another. Despite the huge diversity of synthetic species, the environmental chemistry of organolead is dominated by a relatively small number of products. It is determined almost exclusively by 5 TeAL, these being TeML, TeEL and the mixed methyl –ethyl species, and further the 7 TAL and DAL salts which can possibly result from these ﬁve species through successive dealkylations (see Table 34.1). Organolead derivatives can be divided into two main groups. A ﬁrst class consists of those compounds in which the lead atom is bonded exclusively to carbon or to another lead atom. With tetraorganolead species being the most typical representatives, this class further includes hexaorganodilead and diorganolead species, which are at present merely interesting on theoretical grounds. The low molecular weight TeAL compounds are clear, colorless liquids, whereas the high molecular weight derivatives are solids. Taking into account the small difference in the electronegativities of lead and carbon, the lead-to-carbon bond in tetravalent lead compounds, to which the lead atom contributes a sp3 hybrid orbital, exhibits a high degree of covalent character. This is reﬂected by the physico-chemical properties that characterize the compounds [20–25]. TeALs are soluble in the common organic solvents, such as ether, benzene, chloroform or absolute ethanol. The symmetrical tetra-aryllead compounds,

1088

Sample preparation for speciation of lead

however, appear to be almost insoluble in alcohol or ethers. Generally, R4Pb compounds are quite insoluble in water and, chemically, fairly stable to water and air. Tetraorganolead species, being relatively unreactive compared to other organolead derivatives, will resist organometallic reactions, such as addition to a carbonyl group, as well as reactions with aqueous bases. With aqueous strong acids, a reaction takes place at moderate rate, while towards anhydrous acids or halogens the carbon– lead bond is quite sensitive and cleaved readily and completely. Under more drastic conditions (at elevated temperatures, often in solution in organic solvents) the compounds undergo a wide variety of reactions, both electrophilic and nucleophilic. To produce selective cleavage of one or two leadto-carbon bonds, a strict control of the reaction stoichiometry and temperature is then usually a prerequisite. Pyrolysis of organolead compounds gives free organic radicals [26]. Although PbR4 compounds (R ¼ organic radical) are generally fairly stable to light and can be stored for extended periods in brown glass bottles without excessive decomposition, if air is rigorously excluded, irradiation with ultraviolet light causes decomposition of tetraorganolead compounds to lead metal and free organic radicals. Photolysis tends to produce results similar to pyrolysis. A second class of organolead derivatives contains those compounds built up by one or more lead–carbon bonds, but in addition by at least one bond to an atom other than carbon or lead. Most of the known compounds forming the second class are of the types R3PbX, R2PbX2 and RPbX3, where X is an anionic group, usually a halogen or a carboxylate group. Through dipole moment measurements on several selected tri- and diorganolead halides, it has been determined that the lead–halogen bond in these compounds has a high degree of ionic character, approaching that of the lead –halogen bonds in the divalent lead halides [27]. Although the lead compounds R3PbX, R2PbX and RPbX3 are commonly called “organolead salts”, it should be emphasized that they do not have an ionic structure, but are in fact coordination compounds in which the metal–halogen bond is a more or less polarized covalent bond. Consequently, the solubility of R3PbX species, for example, depends both on the radical R and the anionic group X. The solubility in water diminishes with increasing size of the radical R, undoubtedly due to decreasing solvatation, whereas the solubility will also vary considerably between halides and compounds containing an organic radical X. The tri- and diorganolead compounds are clearly less stable than tetraorganolead species. They are assumed to undergo a slow decomposition, even at room temperature, via a disproportionation reaction [24]: 2Rb3 PbX ! R2 PbX2 þ R4 Pb

ð34:2Þ

2R2 PbX2 ! R3 PbX þ ½RPbX3 ! RX þ PbX2

ð34:3Þ

1089

F. C. Adams and M. Heisterkamp TABLE 34.2 Physical properties of selected organolead compounds [24,25] Compound

Boiling point (8C)

Vapor pressure 208C (mmHg)

Melting point (8C)

Dipole moment (Debye)

Water solubility (mg Pb l21)

Tetramethyllead Trimethylethyllead Tetraethyllead Trimethyllead chloride Triethyllead chloride Diethyllead chloride

110 130 202a

26 7.3 0.26

227 2137

0 0 0 4.47 4.39 4.70

15

190a 166a

,0.1 20,000 50,000

a

Decomposes.

The ﬁrst reaction is reversible; the second is irreversible because of the instability of most RPbX3 species. Because of the relative lability of alkyllead halides, any interpretation of the mechanism of reactions involving these salts should consider the possibility of a simple thermal disproportionation. The high degree of ionic character of the organolead halides is clearly reﬂected in the large number of reactions they undergo that are analogous to those of the divalent lead salts. Among these, the redistribution reactions between organolead compounds, discovered by Calingaert [28,29], should explicitly be mentioned because of their remarkable nature: ﬁrstly an ionizing solvent is required, but only the use of a catalyst, and secondly, the randomness of the reactions leads to product composition obeying the laws of probability. Compounds of the types R2PbX2, and particularly RPbX3 [30], generally tend to be less stable than the R3PbX species. Although the majority of organolead salts are stable to hydrolysis, as evidenced by their facile preparation in aqueous media, this resistance appears to become weaker as the number of alkyl groups decreases. Practically, the organolead salts undergo similar reactions as the tetraorganolead products. Because a detailed survey of the physico-chemical properties would be beyond the scope of this chapter, we refer to the outstanding compilation by Shapiro and Frey [24] for more details. Some important physical properties of those organolead species that are frequently detected in the environment are summarized in Table 34.2. 34.5

SYNTHESIS OF ORGANOLEAD COMPOUNDS

The general synthetic chemistry of organolead has not proceeded markedly and has been extensively reviewed elsewhere [24]. Only a brief summary of frequently applied strategies for the preparation of organolead compounds

1090

Sample preparation for speciation of lead

will be presented here. The synthesis of symmetrical tetraorganolead products is relatively straightforward. On a laboratory scale, reactions of lead halides with organolithium and Grignard reagents are generally preferred, proceeding through intermediate formation of R2Pb and R6Pb2 species. Other organometallic reagents, such as R2Zn, R3Al and R3B, have also been used, but appear to be less versatile or efﬁcient for lead alkylation. A second useful route is the reaction of lead alloys with an organic halide, which enjoys commercial signiﬁcance, being the major process for the manufacture of TeAL anti-knock agents. Industrially, the reaction is carried out with sodium–lead alloy and an equimolar amount of ethyl or methyl chloride in a pressure autoclave, in the presence of a catalyst (see Section 34.2). Three-fourths of the lead is converted back to lead metal and must be recycled; the ﬁnal yield amounts to about 85%. Additionally, reactions for the synthesis of tetraorganolead starting from lead metal have been optimized, involving addition of an alkyl halide, occasionally in combination with another reactive metal or organometallic reagent. Finally, much effort has been devoted to the industrial realization of an electrolytic synthesis of the anti-knock agents. The reaction of a lead anode with a complex organometallic electrolyte has been extensively studied and has led to several interesting laboratory applications [24]. The electrochemical reduction of an alkyl halide at a sacriﬁcial lead cathode enjoyed attention for the preparation of radiolabeled alkyllead standards [31]. For the preparation of unsymmetrical tetraorganolead compounds, only one method is generally suitable; it is the reaction of a triorganolead halide or diorganolead dihalide with a Grignard or organolithium reagent. By treating the resultant reaction mixture with halogen at 2 788C, an unsymmetrical triorganolead halide is produced in situ which, in turn, can be treated with a different Grignard or organolithium reagent. Repetition of this cycle ﬁnally yields a tetraorganolead derivative containing four distinct alkyl groups. The cleavage of the organic group by halogen is not random, shorter chain alkyl groups being cleaved more easily than longer chain groups and aryl preferentially to alkyl groups. Organometallic reagents other than Grignard or organolithium may catalyze a redistribution reaction in which a mixture of all possible compounds is formed. Numerous methods of synthesis are known for R3PbX and R2PbX2 compounds, most of them involving the use of another organolead compound, such as R4Pb as a starting material, and a halogen or halogen acid as reagent. The selectivity largely depends on the choice of the reaction parameters. Presumably, due to their instability, the preparation of monoorganolead salts is more challenging, particularly that of monoalkyllead halides. To our knowledge, only one report described the synthesis of monoalkyllead triiodides [32]; surprisingly, it was found that the compounds were stable at room temperature towards water, acids and bases.

1091

F. C. Adams and M. Heisterkamp

34.6

THE BIOGEOCHEMICAL CYCLE OF LEAD

To summarize the knowledge about sources, occurrence and sinks of organolead in the biosphere, geochemical cycles have been proposed, outlining the fate of the pollutants [6]. Figure 34.1 presents such a schematic overview. Man-made alkyllead enters the atmosphere in the form of TeAL and probably also as TAL, emitted into the atmosphere or spilled in the aqueous or terrestrial environment. In both cases, the original pollutants have a lifetime of a few days at most. Conversion, primarily by reaction with hydroxyl radicals, to TAL and DAL species extends the atmospheric lifetime and allows transport, mainly in the gaseous phase, to more remote areas before complete breakdown to inorganic lead occurs. Aqueous TeAL degrades rapidly to TAL, especially under the inﬂuence of light. Wet and dry deposition processes transfer the species, as TAL, into the hydrosphere and deposit them on soils. From natural water and soil, the products may get accumulated by plants and animals, in concentrations closely following the environmental exposure, and pass on to the food chain. Apart from the most volatile species that may evaporate, the lead species eventually degrade or deposit with excreta, dead organisms or as particulate-bound lead. Sediments and/or biota may, in addition, regenerate alkyllead through a possible environmental alkylation. These last stages, in particular, remain as yet obscure, as do the mechanisms of the various transformations. The determination of the species is necessary in the air (the apolar TeAL) and in water and in various solid phases, such as biological materials, sediments, air particulate matter, road and other dust (the polar TAL salts). It is the determination of the TAL and DAL species in solids, especially in biological materials, that is most demanding.

34.7

ANALYTICAL TECHNIQUES FOR SPECIATION ANALYSIS OF ORGANOLEAD COMPOUNDS

Organolead compounds comprise only a small fraction of the total lead content of a sample. Determination of the different organolead species demands not only separation of the compounds, but also highly sensitive detection techniques able to perform analyses of samples containing the analytes in the ng l21 range for waters and ng g21 for solid matrices. Hyphenated techniques combining powerful separation methods with sensitive and element speciﬁc detection systems are most commonly applied for speciation analysis of lead and other organometallic compounds. Generally, gas chromatography (GC) is superior to liquid chromatography (LC) with regards to resolution, separation time and sensitivity. The main drawback of GC is the necessity to derivatize the ionic, non-volatile organometallic species (and thus transform them to fully coordinated alkyllead species) prior to analysis to obtain volatile, thermally stable compounds still containing the species-speciﬁc information.

1092

Sample preparation for speciation of lead

Originally, total volatile organolead was determined by converting the volatile TeAL compounds emitted to the atmosphere into non-volatile lead salts through a series of reactions with iodine: PbR4 ! PbR3 þ ! PbR2 2þ ! Pb þ2 : The reaction between TeAL and iodine proceeds sufﬁciently fast to allow quantitative collection by passing the air through an aqueous iodine solution. These early procedures for the determination of the environmental impact were required to follow the huge emissions into the atmosphere and culminated in a procedure developed by Hancock and Slater [33] in which organic lead (the sum of TeML, TeEL and mixed methyl/ethyl species) was extracted into carbon tetrachloride with dithizone. Any inorganic lead remained in the aqueous phase by masking with EDTA. The organic lead was then stripped from the carbon tetrachloride extract with nitric acid/hydrogen peroxide solution, the lead solution then being determined by atomic absorption spectrometry (AAS). This procedure was successfully adopted for determinations in environmental samples but suffered from the drawback that it allowed only the determination of total volatile organolead. Species-speciﬁc measurements of airborne organolead implied the use of chromatographic separation techniques by GC –LC, originally with electron capture detection (ECD) [34]. The most important breakthrough occurred with the introduction of hyphenated methods combining GC and AAS for the determination of all ﬁve TAL compounds by Chau et al. [35]. These measurements were highly speciﬁc: only lead containing components in the GC efﬂuent were detected. Moreover, if the AAS measurement is started simultaneously with the sample introduction, the retention time can be used for identifying the compounds. In later modiﬁcations of the procedure, especially by using an electrically heated silica furnace in the AAS unit, the detection limits were decreased to less than 0.1 ng for individual species. Applications to organolead speciation and, soon afterwards, to other organometal species, were the ﬁrst to demonstrate the advantages in selectivity and sensitivity of hyphenated techniques which combined chromatographic separation with sensitive atomic detectors. The most important method for the determination of various organic components, conventional GC –MS, could not reach similar performance characteristics. 34.7.1 Hyphenated techniques for organometal determinations It is appropriate to review at this time what the ideal requirements are of hyphenated methods of analysis and how they can be most conveniently achieved. A number of combinations of separation and detection systems are summarized in Table 34.3. Any of the various combinations can be exploited for the determination of species of varying complexity: simple redox systems (e.g., As5þ and As3þ), structurally simple organometal species (such as the TeAL in this particular section) and, ﬁnally, high molecular mass compounds such as metalloporphyrins, metalloproteins, metalloenzymes and metallodrugs.

1093

F. C. Adams and M. Heisterkamp TABLE 34.3 Frequently coupled separation and detection techniques for speciation analysis Separation techniques Gas chromatography (GC) Packed column Capillary column Multi-capillary column High performance liquid chromatography (HPLC) Reversed phase Ion exchange Ion pair Supercritical ﬂuid chromatography (SFC) Capillary electrophoresis (CE)

Detection techniques Atomic absorption spectrometry (AAS) Flame Electrothermal (graphite/quartz furnace) Optical emission spectrometry (OES) Flame Plasma (MIP) Atomic ﬂuorescence spectrometry (AFS) Mass spectrometry (MS) Electron impact (EI-MS) Plasma (ICP) Flame photometric detection (FPD) Electrochemical detection

The choice of a particular hyphenated technique depends primarily on the problem at hand. The separation component is of special concern when different target species have close physico-chemical properties, while the detector component is important when low limits of detection are required. The interface between both parts of the system is often not problematic and connections and interfaces are often directly constructed in the laboratory. With few exceptions, the analytical instrumentation market has been reluctant to introduce any speciﬁc instrumentation for this purpose. The main requirements for the development of a speciation technique are: (1) selectivity sufﬁcient to determine the chemical forms of the element of interest and (2) high sensitivity for ultratrace measurement [36]. Atomic spectroscopic techniques, including atomic absorption spectroscopy (AAS), electrothermal atomic absorption spectroscopy (ET-AAS), plasma source atomic emission spectroscopy (PS-AES) and plasma source mass spectrometry (PS-MS) offer the high sensitivity required for ultratrace measurements but lack the selectivity necessary to identify different species of the same element. However, the combined selectivity of a separation technique such as GC and capillary electrophoresis (CE), coupled with the sensitivity of atomic spectroscopy, has generated a variety of hyphenated techniques shown to have great promise for speciation analysis [37]. Of these atomic spectroscopic techniques, PS-MS (mainly in the form of microwave induced plasma (MIP)-MS and inductively coupled plasma (ICP)-MS has been particularly effective when applied to speciation analysis [38,39]. ICP-MS is especially attractive and offers

1094

Sample preparation for speciation of lead

a robust system with extremely low detection limits for nearly all elements in the periodic system, and a broad dynamic range. Separation techniques generally produce a time-dependent transient signal. Such transient signals can cause several difﬁculties when measured by scanning mass analyzers, such as the quadrupole mass ﬁlter (QMS) or a sequentially scanned double-focussing mass spectrometer. This scanning process usually requires a trade-off between sensitivity and precision on the one hand, or restricted mass coverage on the other, particularly when the analysis time is limited. Moreover, quantiﬁcation errors known as spectral skew [40] can arise during the measurement of adjacent mass-spectral peaks at different times along a transient signal. Also, non-simultaneous ion extraction limits the ability of scanning mass analyzers to exploit ratioing techniques to reduce multiplicative noise associated with sample introduction and plasma ﬂuctuations [41]. To analyze fast transient signals and to minimize multiplicative noise, selected ion monitoring (SIM) has been utilized with scanning mass spectrometers [42]. SIM increases the effective scan speed of the instrument by measuring only selected mass-spectral peaks instead of the entire mass spectrum. However, SIM restricts the maximum number of elements or isotopes that can be monitored. To overcome such complications, several research groups have successfully modiﬁed mass spectrometers based upon scanning analyzers, to impart to them the ability to measure multiple massspectral peaks simultaneously. The quadrupole ion trap (IT) has recently proven to be an attractive alternative mass analyzer for coupling with an ICP [43]. The combination offers good sensitivity, the ability to selectively reject unwanted ions and the capability to perform tandem mass spectrometric experiments by collisionally dissociating trapped polyatomic ions [44]. Although the IT eliminates spectral skew by sampling all m/z values from the ICP simultaneously, individual m/z values are measured sequentially, resulting in a loss of sampling speed and duty cycle. Additionally, the restricted capacity of an ion trap mass spectrometer (104 –106 ions) can seriously limit its dynamic range. Fourier transform ion cyclotron resonance mass spectrometry (FT-ICR-MS) has also been coupled with the ICP for elemental analysis [45]. The highresolution of FT-ICR-MS can eliminate most isobaric interferences that occur in ICP-MS and the method offers simultaneous multi-elemental measurements. However, cost, ultra-high vacuum requirements, restricted ion-storage capability and lengthy measurement time have limited its routine application. In contrast to the systems described above, time-of-ﬂight mass spectrometry (TOF-MS) is almost ideally suited for the measurement of transient signals. Mahoney and co-workers have recently demonstrated the use of ICPTOF-MS for the detection of transient signals produced by laser ablation [46] and electrothermal vaporization [47]. TOF mass analysis is accomplished by isoenergetically accelerating an ion packet into a ﬁeld-free region (ﬂight tube), in which ions of different masses are separated based on their m/z-dependent

1095

F. C. Adams and M. Heisterkamp

velocities. When given the same kinetic energy, light ions will travel at greater velocities than heavier ions, resulting in a distribution of arrival times at a detector positioned at the end of the ﬂight tube. Measurement of the intensity and ﬂight times of ions arriving at the detector directly produces a mass spectrum. TOF-MS offers several important advantages over other mass analyzers commonly used for elemental analysis. Because of the limited mass range required for complete elemental analysis, spectral generation frequencies of . 20 kHz are typically achieved with moderate acceleration potentials (, 2 kV) and relatively short ﬂight lengths (1 m). Since all m/z values are extracted simultaneously for mass analysis, the spectral skew usually associated with the measurement of transient samples is eliminated. Also, enhanced measurement precision can be achieved with simultaneous multielement ion extraction by eliminating multiplicative noise through use of ratioing techniques. Finally, the simple, open design of TOF mass spectrometers promises to result in enhanced ion transmission efﬁciency and reduced cost. The majority of the species of interest in bioinorganic analysis have not yet been isolated with sufﬁcient purity to be identiﬁed on the basis of chromatographic retention time [48]. Therefore, for these it is important to employ in parallel a molecular (or moiety-) speciﬁc detector to establish the identity of the eluted species. MS in the fast atom bombardment (FAB), electrospray (ES) or matrix-assisted laser desorption ionization mode (MALDI) are available for this purpose. Sometimes it is necessary to combine two or more separation mechanisms in series to insure that a unique species arrives at the detector at any given time. Organolead speciation concerns a relatively small number of well-identiﬁed compounds to be determined at low concentration levels. Important for their determination is speed, sensitivity and accuracy. 34.7.2 Sample preparation The organolead compounds are mostly separated from the matrix by extraction into an organic solvent. TeAL can be transferred directly into the non-polar solvent by extraction using chelating agents such as sodium diethyldithiocarbamate (NaDDTC) or dithizone added prior to extraction for analysis of ionic species, thereby obtaining non-polar complexes. Inorganic lead is present in the samples at concentrations much higher than the organic species. Hence, it is recommended that the inorganic lead be masked with EDTA to avoid memory effects caused by overloading the detector or insufﬁcient derivatization yields in the case of GC separation. The extraction step also includes a preconcentration, which depends on the ratio between the volume of the organic solvent and that of the sample. Another procedure for separation of the analytes from the matrix, in which organolead species from aqueous matrices are adsorbed on ion

1096

Sample preparation for speciation of lead

exchange media, was developed by Hewitt et al. [49]. This method can be applied directly in the ﬁeld, providing very good sample handling and stability of the analyte species. Organolead compounds do not take part in mineralogical processes so they remain physically (adsorption) or chemically (complexation) bound on the surface of the soil. Therefore, a complete digestion of the sediment matrix is not mandatory. Acid leaching of the analytes with simultaneous extraction into an organic solvent after complexation is sufﬁcient to release the species from the matrix. Both DDTC and dithizone are frequently used as chelating agents for this purpose [50]. When analyzing biological tissues or plant material, the matrix must be digested, because organolead compounds are embedded in the tissue of living organisms. This is mostly performed by hydrolysis using tetramethylammonium hydroxide (TMAH) [51,52] or enzymes, such as lipase or protease [53]. These conditions are sufﬁciently moderate to avoid decomposition or transalkylation of the different organolead species. After digestion, the organolead compounds are extracted into an organic solvent after complexation with NaDDTC or dithizone. Separation of organometallic compounds from soil or biological tissue is time consuming using conventional methods. Hydrolysis of biological tissue takes several hours and leaching using ultra-soniﬁcation can take even longer. Application of a focused microwave ﬁeld at low power allows nearly quantitative recovery from these matrices within a few minutes without destroying the carbon –lead bond. Recently, this system was applied to the determination of organotin [54] and organomercury [55] compounds, but it has, to the best of our knowledge, not yet been applied to organolead speciation analysis. An alternative, but rarely used, method for the separation of organolead species from the matrix is supercritical ﬂuid extraction (SFE), where the analytes are extracted into a supercritical ﬂuid (normally, CO2 at high pressure) combining the properties of a gas and a liquid. Due to the polar character of the ionic alkyllead compounds, a suitable modiﬁer must be added for extraction. Johansson et al. [56] added methanol for extraction of organolead from sediment and road dust. Freon chlorodiﬂuoromethane (R-22) was used for extraction of TAL from soil and PVC plastics [57]. The subject of SFE is treated more extensively by Alzaga in Chapter 10. 34.7.3 Sample pretreatment using gas chromatographic separation Sample preparation methods using GC-based hyphenated techniques can generally be divided into two different types; those applying in situ derivatization using tetraalkylborates or tetrahydroborate, and those based on alkylation using Grignard reagents. As organolead hydrides are prone to dismutation reactions, due to their insufﬁcient stability, hydride generation is

1097

F. C. Adams and M. Heisterkamp

seldom applied for determining the different organolead species. An overview of various sample preparation procedures for different matrices using GC-based hyphenated techniques is given in Fig. 34.2. The ﬁrst derivatization method applied for speciation analysis of organolead compounds was alkylation with Grignard reagents, a reaction in which the alkylgroups of alkylmagnesium chlorides or bromides are transferred to the ionic organometal. Fully alkylated species are formed still containing the molecular information of the native organometal. Propylation [58,59] or butylation [60,61] reactions are mostly used for derivatization of the analytes, but pentylation [62] and phenylation [63] are also applied. The former reactions are favorable due to the higher volatility of the generated species, resulting in shorter retention times. Furthermore, derivatives with longer carbon chains are prone to isomerization reactions, leading to peak splitting in the chromatograms. Alkylation with Grignard reagents can only be performed in completely dry media. Hence, organometallic species in water-containing samples have to be extracted into an organic phase, whose dryness must be ensured prior to the alkylation. The destruction of the Grignard reagents in an aqueous medium is the main drawback of this kind of derivatization, resulting in a multi-step and often time-consuming sample preparation procedure. The separation of the analytes from the different matrices and the extraction into an organic solvent is performed as illustrated previously. In recent years, alkylation with alkylborates has become popular for speciation analysis of organometal compounds due to the product’s solubility and stability in aqueous media. Consequently, derivatization can be performed in situ, meaning directly in the water. Even if complex matrices, such as peat or dust, are analyzed, the analytes can be separated from the matrix, derivatized and extracted into an organic solvent simultaneously, making sample preparation a relative fast, one-step procedure. Another advantage of in situ derivatization is the possibility of applying solid phase micro-extraction (SPME) for extraction of the derivatized species from the aqueous medium [63,64]. The analytes are extracted onto a fused silica ﬁber coated with poly-(dimethylsiloxane) either directly in the aqueous medium or using the headspace technique. While only a small fraction of the extraction volume can be injected into the GC when liquid/liquid extraction is applied, SPME allows a quantitative transfer of the extracted analytes into the GC due to the absence of an organic solvent. The ﬁber can be inserted directly into the injection port where the analyte species are thermally desorbed. Hence, the sensitivity of the procedure can be signiﬁcantly increased, resulting in detection limits in the sub-ng l21 range [65]; moreover, the risk of contamination due to insufﬁciently pure solvents is excluded. The commercial availability of sodium tetraethylborate (NaBEt4) has made in situ ethylation an attractive alternative for the speciation analysis of

1098

Sample preparation for speciation of lead

Fig. 34.2. Schematic overview of different sample preparation procedures using GC-based hyphenated techniques.

1099

F. C. Adams and M. Heisterkamp

organotin and organomercury. For organolead speciation, ethylation is only feasible for methylated species [66,67], as both ethyl- and inorganic lead yield TeEL with a consequent loss of species-speciﬁc information. The use of sodium tetrapropylborate (NaBPr4) [68,69] and tetrabutylammonium tetrabutylborate ([Bu4N]þ[BBu4]2) [70,71] allows the speciation analysis of all relevant organolead species. 34.7.4 Extraction recovery TAL and DAL are scavenged from the atmosphere by rainfall and frequently detected at high concentrations in road drainage and surface waters and urban dusts [72] from which they are often not easily extracted. For accurate determinations of these compounds, it is necessary to evaluate extraction recoveries. Major pitfalls have been reported, including poor recovery by ultrasonic extraction, losses related to ﬁltration, etc. In an interlaboratory evaluation of extraction from a spiked urban dust, recoveries using different procedures for extraction and analysis were in the 70–95% range, with standard deviations of 3–8% [73]. The inﬂuence of the reaction medium on the recovery of the different organolead species is shown in Fig. 34.3 for the propylation reaction and detection with ICP-MS. Maximum response is achieved using HAc/NaAc buffer

Fig. 34.3. Inﬂuence of the reaction medium on the recovery of different organolead species after in situ derivatization using NaBPr4.

1100

Sample preparation for speciation of lead

and pH 4.0. At lower pH, the alkylborate is hydrolyzed by the acid according to the equation: NaBPr4 þ HAc ! NaAc þ BPr3 þ PrH

ð34:4Þ

BPr3 þ 3HAc ! BðAcÞ3 þ 3PrH

ð34:5Þ

34.7.5 Separation Since TeAL compounds are volatile and the ionic species are relatively easily derivatized, mostly GC is applied for separation of the analytes. The disadvantage of the derivatization is compensated by the absence of a condensed mobile phase in the measurement system. The use of an inert gas of low viscosity as carrier allows the application of long columns resulting in excellent resolution and high ﬂow rates leading to short separation times. Furthermore, coupling of the GC with the detector is relatively easy and a quantitative transfer of the gaseous eluent into the detector is achieved. 34.7.5.1 Gas chromatographic separation Separation of organolead compounds using GC can be performed with either packed- or open-tubular capillary columns. Both types must be equipped with non-polar stationary phases for the analysis of organolead compounds. In organolead speciation analysis, the stationary phase of the packed columns consists of small, solid particles, mostly covered with a thin layer of a liquid, normally 3 – 10% dimethylpolysiloxane on diatomite (Chromosorb). In recent years, capillary column GC (CGC) of typically 0.25 or 0.32 mm inner diameter, has been preferred due to the higher resolution capability and narrower bandwidth, which is especially recommended when analyzing samples with complex matrices, such as biological tissues. In capillary columns, the glass tubes are coated with a liquid ﬁxed by capillary forces or chemical bonds such as bonded and crosslinked 100% polydimethylsiloxane. Furthermore, capillary columns provide the required inertness necessary for the separation of organometallic compounds, while the active metal–carbon bond may interact with the packed column, often resulting in decomposition or peak tailing. The main drawback of capillary columns is the restricted injection volume (typically 1 ml). This can be overcome by suitable injection systems able to preconcentrate the analytes before releasing them onto the column. Using programmed temperature vaporization (PTV), volumes up to 50 ml can be injected due to a selective evaporation of the solvent while the analytes are simultaneously trapped on a cold liner. The volatility of the solvent needs to be much higher than that of the analytes to ensure a complete elimination of the solvent while the analytes remain adsorbed on the packing of the liner. After evaporation of the solvent, the liner is heated

1101

F. C. Adams and M. Heisterkamp

rapidly and the analytes are transferred onto the column. Another possibility is the use of purge and trap systems for injection, if the analytes are sufﬁciently volatile (boiling point , 1008C) or if they can be easily converted by in situ derivatization to volatile species. The analytes are released from the matrix by a helium ﬂow and trapped on a cold trap before injecting them onto the column by electrical heating of the trap. Recently, GC using multi-capillary columns has been applied to the separation of the different organolead species. These multi-capillary columns consist of a bundle of about 1000 wall-coated open-tubular capillaries, typically of 1 m length with an inner diameter of 40 mm. By reducing the inner diameter of the capillaries, very short columns can be used, allowing highly efﬁcient isothermal separations within a few minutes. The drawback of a reduction in the inner diameter of the capillaries resulting in a lower capacity and, hence, in smaller injection volumes, can be overcome by assembling a large bundle of capillaries to increase the cross-section of the carrier ﬂow. Thus, injection volumes up to 1 ml, which are also typical for common capillary columns, can be achieved. Fast, multi-capillary GC was applied to the analysis of TML in a Certiﬁed Reference Material (CRM 605, road dust) [74] and of TAL species in gasoline [75]. In contrast to capillary columns having inner diameters of about 300 mm, the number of measurements performed per hour can be increased, as the separation time is signiﬁcantly shortened (a few minutes instead of 10– 15 min) and there is no necessity to cool down the column after a GC run. 34.7.5.2 Liquid chromatographic separation High performance liquid chromatography (HPLC) is less often applied to the speciation analysis of organolead compounds. The main advantage of this technique is the possibility of analyzing the naturally occurring species without the need for derivatization. Sample preparation becomes faster and the risk of contamination during the derivatization process is avoided. In contrast to GC, signiﬁcant interactions between the analytes and the mobile phase appear, which makes the composition of the mobile phase an important parameter for obtaining high separation efﬁciency. For the separation of organometallic compounds, different types of HPLC can be applied using various mobile phases. Reversed phase chromatography is mostly used, based on partitioning of the analytes between a non-polar stationary phase (C18) and a polar mobile phase after chelating the species. The complexation can be performed before separation in an extraction step during sample preparation, or the chelating agent can be added directly to the mobile phase. Usually, this phase consists of a mixture of a buffer and methanol, whose composition is changed gradually to improve the separation efﬁciency of the method. Complexation of the organolead species is mostly achieved using dithiocarbamates [76], but dithizone or mercaptoethanol [50] have also been applied. Another LC separation method is ion pair chromatography, which has as its basis

1102

Sample preparation for speciation of lead

the distribution of an ion pair between the stationary and the mobile phase. Both phases have the same composition as those in reversed phase chromatography, but the mobile phase contains an additional ion pairing reagent, such as pentasulfonic acid [77]. A third technique is ion exchange chromatography, with a stationary phase consisting of a porous ion exchange resin and a buffer solution of a particular ionic strength as the mobile phase [78]. The main disadvantages of HPLC-based techniques are its poor resolving power and the necessity to evaporate the liquid mobile phase before transferring the efﬂuent to the detection system. Nebulization using common systems suffers from a very poor efﬁciency, approximately 1%, leading to a loss in sensitivity of 99% compared with the quantitative transfer of the analytes to the detector using GC-based hyphenated techniques. For the speciation analysis of organolead compounds, post-column derivatization using hydride generation [79] or ethylation [50] can improve the sensitivity by circumventing the nebulization of the efﬂuent. Post-column ethylation is suitable for both methyl- and ethyllead speciation, as the species are already separated and characterized by their retention time. 34.7.5.3 Alternative separation techniques Supercritical ﬂuid chromatography (SFC) combines advantages of both GC and LC. By using a supercritical ﬂuid as the mobile phase, its special properties intermediate between gas and liquid can be employed (cf. Chapter 10). Due to its low viscosity, a resolution comparable to that of GC can be obtained and its high partial pressure allows mild separation temperatures, which makes SFC suitable for the analysis of thermally unstable compounds. The coupling to element speciﬁc detection systems is relatively easy, since the supercritical ﬂuid is decompressed to a gaseous state prior to detection. Hence, nearly quantitative transfer efﬁciency to the detector can be obtained by eliminating the need for nebulization of the mobile phase. Consequently, using SFC, the ionic, non-volatile organometals can be separated as native species without derivatization with a resolution and sensitivity comparable to GC. Nevertheless, the use of SFC for separation of organometallic compounds is still in its infancy. Carey et al. [80] applied SFC with plasma mass spectrometric detection for the speciation analysis of organolead and organomercury compounds. Electrophoresis procedures are based on the varying migration rates of ions in an electrical ﬁeld according to their characteristic mobility. Using CE, the analytes are placed in a narrow zone between buffer solutions and the ions migrate through a capillary. A CE capillary coupled to plasma mass spectrometry was recently applied for the speciation analysis of TML and TEL [81]. For the analysis of non-polar compounds, micellar electrokinetic chromatography (MEKC) is applied, where the analytes are trapped in micelles of an ionic surfactant making migration possible in an aqueous solution

1103

F. C. Adams and M. Heisterkamp

through the electrical ﬁeld. MEKC has a higher selectivity towards separation of the organolead species, as a result of the interaction of the analytes with the micelles [82]. 34.7.6 Detection of organolead compounds after chromatography Common detectors in GC, based on thermal conductivity (TCD), ﬂameionization (FID) or ECD, for example, are not sufﬁciently selective to meet the requirements for speciation analysis of organolead compounds. When analyzing complex matrices like dust, biological tissue or peat, element speciﬁc detection methods are especially recommended to obtain reliable results. Therefore, spectrometric detectors measuring element speciﬁc properties like atomic absorption, emission or ﬂuorescence on a characteristic wavelength of the analyte are nowadays applied in addition to mass spectrometers detecting a speciﬁc mass-to-charge (m/z) ratio (see Section 34.7.1). The main advantage of these spectrometric detectors is the possibility of eliminating interferences from matrix compounds (mainly hydrocarbons) due to the lead-speciﬁc measurement and very low detection limits, even with incomplete chromatographic separation. 34.7.6.1 Atomic absorption spectrometry Coupling of GC and AAS can be performed very easily using a ﬂame for atomization by introducing the efﬂuent via a heated transfer line directly into the burner. But low ﬂame temperatures and short residence times of the atoms in the ﬂame and the dilution effects caused by the ﬂame gases result in a poor sensitivity. Therefore, a graphite furnace or, preferably a heated quartz tube (QT) is recommended for analysis, the graphite furnace not being constructed for continuous heating during the extent of the chromatographic run. Interfacing of a heated QT to a GC can be realized by introducing the efﬂuent into the oven via a heated side arm [83]. The supporting gases (hydrogen and air) are heated to avoid condensation of less volatile analytes at the cold surface of the tube or at so-called cold spots in front of it. Graphite furnaces are also used for atomization of the analytes with comparable analytical characteristics, but they are more expensive and difﬁculties occur due to the need for operation at high temperatures for the entire measurement period of a chromatogram; this makes them disadvantageous for routine analysis. Interfacing HPLC with AAS is more sophisticated, due to the necessity of continuously removing the liquid mobile phase. Usually, nebulization is used to convert the efﬂuent into an aerosol subsequent to the separation. While ﬂames can handle wet aerosols, drying and ashing of the efﬂuent is necessary when using graphite furnace (electrothermal) atomization, which makes direct coupling of the column with the furnace practically impossible. Nevertheless, several indirect interfaces have been designed to introduce the efﬂuent into the furnace of the AAS instrument [84].

1104

Sample preparation for speciation of lead

34.7.6.2 Optical emission spectrometry Optical emission spectrometry (OES) is predominantly applied for TeAL speciation using a microwave induced plasma (MIP) as an excitation source for the GC efﬂuent. A commercial instrument is available that was originally developed as a universal detector for GC efﬂuents and was quite appropriate for the detection of Pb, Hg, Sn and other metal species. It is based on the combination of an atomic emission detector (AED) (HP Model 5921A, Hewlett Packard) and a CGC system from the same manufacturer. The realization of a helium MIP operating at low power at atmospheric pressure using a cavity (TM010) designed by Beenakker [85] was the starting point for a development that resulted in a commercially available GC –MIPOES system in 1989. Interfacing of the HP instrument is realized by transferring the efﬂuent via a heated transfer line directly into the discharge tube within a few mm of the plasma, thus resulting in a minimum dead volume. Due to its small size (the plasma is generated in a quartz capillary) and low plasma temperature, the MIP cannot tolerate molecular species, such as hydrocarbons or water. The plasma part of the instrument is separated from the spectrometer part by means of a quartz window at the end of the discharge tube. Detection is performed over a 20 (visible) to 40 nm (UV) wide spectral interval by a photodiode array detector. As such, it is suitable for the detection of lead speciﬁc emissions at and around 405.783 or 261.418 nm. Overloading or even extinguishing of the plasma by solvent vapors can be avoided by venting the solvent before they enter the plasma. Therefore, the chromatographic conditions have to provide a complete separation of the analytes from the solvent prior to detection to allow solvent venting. The use of helium as plasma gas (compared to argon) is very advantageous, as it is a common GC carrier gas and, in particular, provides a simpler spectral background and a signiﬁcantly higher excitation energy, even allowing determination of non-metals. The emitted radiation is viewed axially, avoiding interferences caused by deposits on the wall of the discharge tube. The determination of Pb and the simultaneous determination of Pb, Sn and Hg is possible [75]. Application of the ICP as an excitation source for OES is not suitable for GC separation due to the strong dilution of the efﬂuent by the plasma, which leads to poor sensitivity. ICP-OES was used by Ibrahim et al. [86] as an HPLC detector with a conventional interface based on a glass-frit nebulizer. When transferring the mobile phase into the plasma, several effects must be taken into account. Mobile phases containing a large amount of organic solvent, such as methanol, give rise to plasma instabilities and ﬁnally quench it. Furthermore, the salt content in the mobile phase should be low to prevent clogging of the nebulizer and the torch injector. Other plasma types, such as direct current plasma (DCP), alternating current plasma (ACP) or capacitively coupled plasma (CCP) are not established as excitation sources for OES detection. Their analytical characteristics cannot challenge those offered by MIP-OES or ICP-OES. Some potential for DCP-OES

1105

F. C. Adams and M. Heisterkamp

was shown by Panaro et al. [87] wherein detection limits in the low pg range were achieved. 34.7.6.3 Mass spectrometry Mass spectrometry (MS) of both molecular and atomic ions can be applied for speciation analysis of organolead compounds. The analytes are ionized and detected at speciﬁc m/z ratios using a quadrupole, a magnetic sector, a quadrupole IT or a time-of-ﬂight (TOF) mass analyzer. MS employing QMSs is most frequently used due to its low costs and adequate stability, but its major disadvantage is its inherently low mass resolution, as it is only able to detect nominal masses. If, due to isobaric interferences, higher resolution is required, magnetic sectors with a resolution up to 10,000 can be used. Both quadrupole and magnetic sector instruments are scanning mass analyzers determining sequentially one mass at a time while rejecting all others. The use of TOF analyzers for MS detection (TOF-MS) allows very fast measurement of all ions across the full mass range, which makes them very advantageous for analyses of fast transient signals, as in the efﬂuent from GCs. Relatively inexpensive bench top GC –EI-MS systems using SIM offer very good sensitivity with detection limits in the low pg range, two orders of magnitude better than operating in the full scan mode normally used for structure elucidation. These properties make GC – EI-MS an attractive technique for speciation analysis of organolead compounds [30]. Recently, GC coupled to an ion-trap mass spectrometer with EI ionization in the tandem-MS mode was applied for simultaneous speciation analysis of organolead, tin and mercury compounds after their in situ ethylation and SPME sampling, leading to detection limits at the low ng l21 level [34]. Recently, ICP-MS has attracted great attention as an element speciﬁc detection system for GC, LC, CE and SFC. Generally, a gain in sensitivity of two orders of magnitude over ICP-OES can be obtained. Interfacing of ICP-MS with GC for TeAL analysis can be performed by directly introducing the efﬂuent via a heated transfer line. The argon used as make-up gas is also transferred in a second heated stainless steel tube to avoid condensation of the analytes at the end of the transfer line [88]. Moreover, solvent venting to prevent instability or even extinguishing of the plasma is, contrary to MIP-OES, not necessary. Using this hyphenated system, detection limits for Pb compounds in the low fg range can be achieved [89,90]. An alternative interface design is based on a heated metallic block, which holds tubing housed in copper at the end of the capillary column in a T-joint arrangement. The make-up gas is introduced perpendicularly to the metallic tube without being heated. The T-joint is connected to the torch using a ﬂexible, non-heated PTFE tube [91]. When coupling HPLC to ICP-MS, the same precautions concerning the plasma have to be taken as for using ICP-OES as the detection system. The salt content and the presence of organic solvents in the mobile phase must be minimized, because the sampler and skimmer can be eroded or plugged by

1106

Sample preparation for speciation of lead

large amounts of salt. This affects the ion input aperture regulating the ion beam and, thereby, the sensitivity. Much of the efﬂuent is lost during nebulization and the high dead volume of the spray chamber can cause peak broadening. The employment of a direct injection nebulizer (DIN) placed inside the ICP torch signiﬁcantly increases the nebulization efﬁciency while peak broadening is minimized due to the low dead volume and the absence of a spray chamber. Additionally, the nebulizer allows the use of microscale LC columns and liquid ﬂow rates of 30 –100 ml min21, providing a nearly quantitative transfer of the analytes into the plasma. Thus, absolute detection limits in the sub-pg range can be achieved [92]. One of the many advantages of ICP-MS, apart from its inherent sensitivity, is the possibility of measuring elemental isotope ratios, which allows incorporation of isotope dilution into the analytical procedures. This requires the synthesis of calibrants with veriﬁed purity and stoichiometry [93]. Isotopically enriched TeAL compounds are now available from several sources. Commercially available calibrants of the different TeAL compounds need to be veriﬁed for their purity. Moreover, it was frequently reported that deterioration occurs over a long time period. A Certiﬁed Reference Material (CRM 605) of TML in urban dust was made available for validation work, but there are at present doubts about the long-term stability of this material [94]. Experiments indicate that even when stored in the dark at 48C there are losses of about 30 –40% over a period of 1 year [94]. 34.7.7 Procedures for the determination of organolead compounds in dust material The following procedures were used in the certiﬁcation of CRM 605 and can serve as an example of a number of state-of-the-art procedures [72]. Ethylation/GC – QT-AAS: Mixing of the sample with NaCl/H2O, followed by mechanical shaking and ﬁltering, derivatization with NaBEt4, cryogenic trapping in a U-tube ﬁlled with chromatographic material, and ﬁnal detection by QT-AAS. A variant of this method: a mixture of sample with NaAc –HAc is diluted with water and methanol, subjected to ultrasonic extraction and backextraction into hexane with separation by CGC. Propylation/GC – QT-AAS: After wetting the sample with an aqueous solution of NaCl, the slurry is ﬁltered and the inorganic Pb is complexed with EDTA. Ammonia is added and complexation of the analyte with DDTC is performed followed by hexane extraction; Grignard derivatization is performed with propylmagnesium chloride in diethylether, followed by a clean-up on a silica solid phase extraction column, eluted with hexane. Separation is with GC (U-tube ﬁlled with chromatographic material) followed by QT-AAS detection. Ethylation/CGC – MIP-OES: NaBEt4 derivatization is carried out after addition of ammonium citrate –EDTA and hexane containing Bu4Pb as an internal standard. Separation by CGC is followed by detection with MIP-OES.

1107

F. C. Adams and M. Heisterkamp

SFE/propylation/CGC – MS: NaOH is added to the sample, with buffering using borax –hydrochloric acid. Extraction is by SFE using CO2 with methanol as a modiﬁer. Liquid –liquid extraction of the SFE eluate is performed with n-hexane after complexation with DDTC. Grignard derivatization is carried out with propylmagnesium chloride in diethylether. Separation is with CGC, followed by EI-MS detection of ions m/z 223 and 253. Pentylation/CGC – MS: An aqueous buffer is added to the sample together with DDTC. Extraction is performed twice with pentane, followed by clean-up with activated alumina and elution with hexane –diethylether. Grignard derivatization is carried out with pentylmagnesium bromide. Separation is with CGC, followed by EI-MS and detection of ions at m/z 307 and 309. Butylation/CGC – MS: Extraction is performed with pentane after buffering with ammonium acetate and DDTC complexation. Clean up is carried out with deactivated alumina, followed by hexane elution. Derivatization is performed with butyl-magnesium chloride in THF, followed by addition of H2SO4. Separation is by CGC, followed by MS detection in SIM; monitoring the ions at m/z 208, 223 and 253 (Me3BuPb), m/z 208, 237 and 297 (Et4Pb) and m/z 208 and 379 (Bu4Pb). The results of these methods for the analysis of a candidate CRM (CRM 605, urban dust) are given in Table 34.4. Note the superior precision obtained by the application of isotope dilution mass spectrometry. Detailed procedures for the analysis of water and road dust based on in situ butylation using [Bu4N]þ[BBu4]2 are as follows [69]: Analysis of water samples: A 20 ml aliquot of the water sample is buffered to pH 4.0 with 1 ml HAc/NaAc buffer solution and placed in an extraction vessel together with 0.5 ml of a 0.1 M solution of EDTA to mask the inorganic lead. After adding 400 ml of methanolic [Bu4N]þ[BBu4]2 solution and 500 ml hexane, the mixture is shaken manually for 5 min and TABLE 34.4 Means and standard deviation of results obtained using different techniques for the certiﬁcation of TML in urban dust (CRM 605) [96] Method (see text)

Mean ^ S.D. (mg kg21)

CV (%)

Ethylation/GC-QF-AAS Propylation/GC-QF-AAS Ethylation/CGC-MIPOES SFE/propylation/CGC-MS Pentylation/CGC-MS Butylation/CGC-MS HPLC/ID ICP-MS Certiﬁed value

9.1 ^ 1.5 7.7 ^ 0.9 10.1 ^ 0.8 6.4 ^ 0.4 7.0 ^ 1.0 7.3 ^ 0.4 8.0 ^ 0.2 7.9 ^ 1.2

16.5 11.7 7.9 6.3 14.3 7.3 2.5

1108

Sample preparation for speciation of lead

set aside for another 2 min to enable phase separation. The organic phase is sampled and stored at 208C in the dark until analysis. Analysis of road dust and materials such as peat samples: Approximately 0.2 g of urban dust or 0.5 g of freeze-dried and milled peat is accurately weighed and placed into a centrifugation vessel, together with 30 ml acetate buffer solution. Then, 500 ml of EDTA solution, 400 ml of the methanolic [Bu4N]þ[BBu4]2 and 1.5 ml hexane are added and the mixture is shaken manually for 10 min and centrifuged 4 min at 4000 min21 to enable phase separation. The organic phase is sampled and stored at 208C in the dark until analysis. Reagents: The [Bu4N]þ[BBu4]2 reagent can be purchased from Alfa (Karlsruhe, Germany). A 0.3% (m/v) methanolic solution should be prepared daily. The different acetate buffer HAc/NaAc solutions (0.1 mol l21) can be made by dissolving 13.6 g of sodium acetate tri-hydrate in 1 l of water and the pH values ranging from pH 4.0 to 5.0 adjusted with an appropriate amount of concentrated acetic acid (96%). For preparation of the different citric acid/ammonia (HCi/NH3) buffer solutions (0.1 mol l21), 21.0 g of citric acid monohydrate is dissolved in 1 l of water and the pH values ranging from pH 2.0 to 11.4 are adjusted by adding appropriate amounts of an aqueous ammonium hydroxide solution (25%). 34.7.8 Comparison of the different hyphenated systems An overview of the absolute detection limits obtained with the various hyphenated techniques used for speciation analysis of organolead compounds is illustrated in Fig. 34.4. Generally, GC is superior to HPLC regarding resolution, chromatographic background and transfer efﬁciency to the detector, but the development of new and effective sample introduction systems makes the use of HPLC-based hyphenated techniques an attractive alternative for speciation analysis. Application of SFC is still scarce, despite its promising features; it is gaining interest due to its potential advantages over conventional GC or HPLC systems. Application of ICP-MS as the detection system offers the best sensitivity for both GC and HPLC separation, but instrumentation is relatively costly and the lack of commercially available hyphenated systems hampers its common use. However, ICP-MS has become the detector of choice using HPLC due to its excellent sensitivity and selectivity. There are different commercially available alternatives for GC-based coupled systems. The sensitivity of a GC –MIP-OES system is comparable to GC –ICP-MS and multi-element determination is also possible, making the former one of the most frequently applied GC-based hyphenated techniques in recent years. Another attractive alternative is offered by GC – MS, because this system is relatively inexpensive and routinely applied in almost every lab dealing with organic analyses and structure

1109

F. C. Adams and M. Heisterkamp

Fig. 34.4. Comparison of the absolute detection limits obtained using hyphenated techniques for organolead speciation analysis as compiled from the literature.

elucidation. Detection limits in the low pg range can be obtained, which is sufﬁcient for most samples, but this technique is, despite its advantages, not fully exploited for speciation analysis. Although atomic ﬂuorescence spectrometry (AFS) promises very good capabilities concerning multi-element analysis, sensitivity and simplicity of the interface, this method is seldom applied [95]. The obstacle for application of AFS as a detection system for GC is the lack of light sources reaching high intensity and stability, but improvements in laser techniques making stable tunable lasers of high intensity widely available could pave the way for use of laser excitation AFS as a GC detector. A few years ago, sample preparation was the most time-consuming step in speciation analysis of organometal compounds, but the introduction of reagents allowing in situ derivatization and the development of microwave-assisted methods changed this situation. Chromatographic separation became the timelimiting factor, but the application of multi-capillary columns made possible the isothermal separation of the analytes within a few minutes. Data processing of the detection systems is now the bottleneck of the whole procedure, because the peak widths are very small, requiring fast data handling of the detector. The recent commercial availability of ICP-TOF-MS provides excellent features for detection of fast transient signals due to its high sampling frequencies, with 20,000 complete mass spectra per minute and its high data acquisition rate. A comparison of several gas chromatographic detectors (MS, MIP-OES and ICP-MS with time-of ﬂight) was made for rainwater samples [96]. GC –EI-MS

1110

Sample preparation for speciation of lead TABLE 34.5 Analytical ﬁgures of merit of the procedure cited in Ref. [100]

Absolute detection limit (fg as Pb) Detection limit (fg g21 as Pb) Recovery (%)

TML

DML

TEL

DEL

14 70 94 ^ 3

15 75 95 ^ 2

10 50 94 ^ 5

12 66 93 ^ 2

is the most speciﬁc option as it derives identity from molecular information but cannot detect the compounds in samples at concentrations of ca. 2 pg ml21, the concentration level at which they are present in countries where the leaded gasolines are now banned. GC –ICP-TOF-MS is the most sensitive, with absolute detection limits of about 15 fg for organolead compounds (as Pb), but MIP-OES almost reaches the same sensitivity. Only the most sensitive methods of detection allow the determination of organolead species in remote environmental samples, such as those obtained in the remote polar environment of Greenland and Antarctica. Archiving of organolead compounds in dated snow and ice shows the impact of pollution and the inﬂuence of the lead used as anti-knock agents [97–101]. In central Greenland, ice samples range from approximately 0.05 to 0.5 pg g21 (as Pb) [97,99], thereby requiring extremely sensitive tools for detection. The ﬁgures of merit of the procedure adopted in our laboratory using NaBPr4 derivatization, at pH 4.5 and measurement with ICP-TOF-MS are shown in Table 34.5 [100].

REFERENCES 1 2 3 4 5 6

7 8 9 10

R. Łobin´ski, Analyst, 120 (1995) 615. A.P. Walton, L. Ebdon and G.E. Milward, Appl. Organomet. Chem., 2 (1988) 87. P.J. Craig and S. Rapsomanikis, Environ. Sci. Technol., 19 (1985) 726. R. Pongratz and K.G. Heumann, Chemosphere, 36 (1998) 1935. R. Pongratz and K.G. Heumann, Chemosphere, 39 (1999) 89. R.J.A. Van Cleuvenbergen and F.C. Adams, Organolead compounds. In: O. Hutzinger (Ed.), The Handbook of Environmental Chemistry, Vol. 3. Springer, Berlin, 1990, Part E. D.S. Forsyth, X.D. Marshall and M.C. Collette, Appl. Organomet. Chem., 2 (1988) 233. R.D. Rhue, R.S. Mansell, L.T. Ou, S.R. Cox and Y. Ouyang, Crit. Rev. Environ. Control, 22 (1992) 233. P. Grandjean, Organolead exposures and intoxications. In: P. Grandjean (Ed.), Biological Effects of Organolead Compounds. CRC Press, Boca Raton, FL, 1984. J.E. Cremer, Possible mechanisms for the selective neurotoxicity. In: P. Grandjean (Ed.), Biological Effects of Organolead Compounds. CRC Press, Boca Raton, FL, 1984.

1111

F. C. Adams and M. Heisterkamp 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29

30 31 32 33 34 35 36 37 38

39 40 41 42 43 44 45

46 47

1112

J.O. Nriagu, Sci. Total Environ., 92 (1990) 13. S.P. Nickerson, Chem. Educ., 31 (1954) 560. P. Grandjean and T. Nielsen, Residue Res., 72 (1979) 97. D. Rosner and G. Markowitz, Am. J. Public Health, 75 (1985) 344. Council Regulation (EC) 98/70/EG, Ofﬁcial Journal of the European Communities, 1998, 41, No. L 350, 58. V.M. Thomas, Ann. Rev. Energy Environ., 20 (1995) 301. V.M. Thomas, R.H. Socolow, J.J. Fanelli and T.G. Spiro, Environ. Sci. Technol., 33 (1999) 3942. G. Ducoffre, F. Claeys and P. Bruaux, Environ. Res., 51 (1990) 25. G. Tiravanti and G. Boari, Environ. Sci. Technol., 13 (1979) 849. G. Calingaert, Chem. Rev., 2 (1926) 43. E. Krause and A. von Grosse, Die Chemie der metallorganischen Verbindungen. Borntraeger, Berlin, 1937, p. 372. R.W. Leeper, L. Summers and H. Gilman, Chem. Rev., 54 (1954) 101. R.L. Milde and H.A. Beatty, Adv. Chem. Ser., 23 (1959) 306. H. Shapiro and F.W. Frey, The Organic Compounds of Lead. Wiley, New York, 1968. F.W. Frey and H. Shapiro, Top. Curr. Chem., 16 (1971) 243. K.M. Gilroy, S.J. Price and N.J. Webster, Can. J. Chem., 50 (1972) 2639. G.L. Lewis, P.F. Oesper and C.P. Smyth, J. Am. Chem. Soc., 62 (1940) 3243. G. Calingaert and H.A. Beatty, J. Am. Chem. Soc., 61 (1939) 2748. G. Calingaert and H.A. Beatty, The redistribution reaction. In: H. Gilman (Ed.), Organic Chemistry, an Advanced Treatise, 2nd edn. Organic Chemistry, an Advanced Treatise, Vol. II. Wiley, New York, 1943, p. 1806. D. de Vos and J. Wolters, Rev. Silicon, Germanium, Tin, Lead Compounds, 4 (1980) 209. J.S. Blais and W.D. Marshall, J. Environ. Qual., 17 (1988) 457. G. Chobert and M. Devaud, J. Organomet. Chem., 153 (1978) C23. S. Hancock and A. Slater, Analyst, 100 (1975) 422. V. Cantuti and G.P. Cartoni, J. Chromatogr., 32 (1968) 641. Y.K. Chau, P.T.S. Wong and H. Saitoh, J. Chromatogr. Sci., 14 (1976) 162. A.M. Leach, M. Heisterkamp, F.C. Adams and G.M. Hieftje, J. Anal. At. Chem., 15 (2000) 151. K. Sutton, R.M.C. Sutton and J.A. Caruso, J. Chromatogr. A, 789 (1997) 85. E.H. Evans, J.J. Giglio, T.M. Castilliano and J.A. Caruso, Inductively and Microwave Induced Plasma Sources for Mass Spectrometry. Royal Society of Chemistry, Cambridge, 1995. A. Montasser, Inductively Coupled Mass Spectrometry. Wiley, New York, 1998. J.F. Holland, C.G. Enke, J. Allison, J.T. Stults and B. Pickston, Anal. Chem., 55 (1983) 997A. N. Furuta, J. Anal. At. Chem., 6 (1991) 199. J.T. Watson, Introduction to Mass Spectrometry. Raven Press, New York, 1985. D.W. Koppenaal, C.J. Baronaga and M.R. Smith, J. Anal. At. Chem., 9 (1994) 1053. D.C. Duckworth and C.M. Barshick, Anal. Chem., 70 (1998) 709. K.E. Milgram, F.M. White, K.L. Goodner, C.H. Watson, D.W. Koppenaal, C.J. Barinaga, B.H. Smith, J.D. Winefordner, A.G. Marshall, R.S. Houk and J.R. Eyler, Anal. Chem., 69 (1997) 3714. P.P. Mahony, S.J. Ray and G.M. Hieftje, J. Anal. At. Chem., 11 (1996) 401. P.P. Mahony, S.J. Ray, G. Li and G.M. Hieftje, Anal. Chem., 71 (1999) 178.

Sample preparation for speciation of lead 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83

J. Szpunar, Analyst, 125 (2000) 963. C.N. Hewitt, P.J. Metcalfe and R.A. Street, Water Res., 25 (1991) 91. J.S. Blais and W.D. Marshall, J. Anal. At. Spectrom., 4 (1989) 641. R. Van Cleuvenbergen, D. Chakraborti and F.C. Adams, Anal. Chim. Acta, 228 (1990) 77. C.S. Epler, T.C. O’Harer and G.C. Turk, J. Anal. At. Spectrom., 9 (1994) 79. D.S. Forsyth and W.D. Marshall, Anal. Chem., 55 (1983) 2132. J. Szpunar, V.O. Schmitt, O.F.X. Donard and R. Łobin´ski, Trends Anal. Chem., 15 (1996) 181. C. Gerbersmann, M. Heisterkamp, F.C. Adams and J.A.C. Broekaert, Anal. Chim. Acta, 350 (1997) 273. M. Johansson, T. Berglo¨f, D.C. Baxter and W. Frech, Analyst, 120 (1995) 755. K. Li and S.F.Y. Li, J. Chromatogr. Sci., 33 (1995) 309. R. Łobin´ski and F.C. Adams, Anal. Chim. Acta, 262 (1992) 285. Y. Wang, A.B. Turnbull and R.M. Harrison, Appl. Organomet. Chem., 11 (1997) 889. C. Nerı´n, B. Pons and R. Zuﬁaurre, Analyst, 120 (1995) 751. M.A.R. Abdel-Moati, Environ. Pollut., 91 (1996) 97. V. Minganti, R. Capelli and R. De Pelligrini, Fresenius J. Anal. Chem., 351 (1995) 471. L. Moens, T. De Smaele, R. Dams, P. Van Den Broeck and P. Sandra, Anal. Chem., 69 (1997) 1604. L. Dunemann, H. Hajimiragha and J. Bergerow, Fresenius J. Anal. Chem., 363 (1999) 466. T. De Smaele, L. Moens, P. Sandra and R. Dams, Mikrochim. Acta, 130 (1999) 241. C. Witte, J. Szpunar-Łobin´ska, R. Łobin´ski and F.C. Adams, Appl. Organomet. Chem., 8 (1994) 621. M. Ceulemans and F.C. Adams, J. Anal. At. Spectrom., 11 (1996) 201. T. De Smaele, L. Moens, R. Dams, P. Sandra, J. Van der Eycken and J. Vandyck, J. Chromatogr. A, 793 (1998) 99. M. Heisterkamp and F.C. Adams, Fresenius J. Anal. Chem., 362 (1997) 489. K. Bergmann and B. Neidhart, Fresenius J. Anal. Chem., 356 (1996) 57. M. Heisterkamp and F.C. Adams, J. Anal. At. Spectrom., 14 (1999) 1307. Ph. Quevauviller, R. Harrison, F. Adams and L. Ebdon, Trends Anal. Chem., 19 (2000) 195. Ph. Quevauviler, L. Ebdon, R.M. Harrison and Y. Wang, Appl. Organomet. Chem., 13 (1999) 1. R. Łobin´ski, V. Sidelnikov, Y. Patrushev, I. Rodriguez Periro and A. Wasik, Trends Anal. Chem., 18 (1999) 449. I. Rodriguez Pereiro and R. Łobin´ski, J. Anal. At. Spectrom., 12 (1997) 1381. J. Bettmer, C. Cammann and M.J. Robecke, J. Chromatogr. A, 654 (1993) 177. A.A. Brown, L. Ebdon and S.J. Hill, Anal. Chim. Acta, 286 (1994) 391. D.K. Orren, J.C. Caldwell-Kenkel and P. Mushak, J. Anal. Toxicol., 9 (1985) 258. H.J. Yang and S.J. Jiang, J. Anal. At. Spectrom., 10 (1995) 963. J.M. Carey, N.P. Vela and J.A. Caruso, J. Anal. At. Spectrom., 7 (1992) 1173. D. Schaumlo¨ffel and A. Prange, Fresenius J. Anal. Chem., 364 (1999) 452. P.R. Haddad, M. Macka, E.F. Hilder and D.P. Bogan, J. Chromatogr. A, 780 (1997) 329. W.M.R. Dirkx, M.B. de la Calle, M. Ceulemans and F.C. Adams, J. Chromatogr. A, 683 (1994) 51.

1113

F. C. Adams and M. Heisterkamp 84 85 86 87 88 89 90 91 92 93

94 95 96 97 98 99 100 101

1114

L. Ebdon, S. Hill and R.W. Ward, Analyst, 112 (1987) 1. C.I.M. Beenakker, Spectrochim. Acta, 32B (1976) 483. M. Ibrahim, W. Nisamaneepong, D.L. Haas and J.P. Caruso, Spectrochim. Acta, 40B (1985) 367. K.W. Panaro, D. Erickson and I.S. Krull, Analyst, 112 (1987) 1097. T. De Smaele, P. Verrept, L. Moens and R. Dams, Spectrochim. Acta, 50B (1995) 1409. A. Prange and E. Jantzen, J. Anal. At. Spectrom., 10 (1995) 105. M. Heisterkamp, T. De Smaele, J.P. Candelone, L. Moens, R. Dams and F.C. Adams, J. Anal. At. Spectrom., 12 (1997) 1077. M. Montes Bayo´n, M. Cutie´rrez Camblor, J.I. Garcia Alonso and A. Sanz-Medel, J. Anal. At. Spectrom., 14 (1999) 1317. S.C.K. Shum, H. Pang and R.S. Houk, Anal. Chem., 64 (1992) 2444. S.J. Hill, A. Brown, C. Rivas, S. Sparkes and L. Ebdon, in: Ph. Quevauviller, E. Maier and B. Griepink (Eds.), Quality Assurance for Environmental Analysis. Elsevier, Amsterdam, 1995, p. 412. R. Van Cleuvenbergen, W. Dirkx, P. Quevauviller and F. Adams, Int. J. Environ. Anal. Chem., 47 (1992) 21. A. D’Ulivo and P. Papof, J. Anal. At. Spectrom., 1 (1986) 479. J.R. Baena, M. Gallego, M. Valcarcel, J. Leenaers and F. Adams, Anal. Chem., 73(16) (2001) 3927. R. Lobinski, C. Boutron, J.P. Candelone, S. Hong, J. Szpunar-Lobinska and F. Adams, Environ. Sci. Technol., 28 (1994) 1459. P.-L. Tesseidre, R. Lobinski, M.-T. Cabanis, J. Szunar-Lobinski, J.-C. Cabanis and F.C. Adams, Sci. Total Environ., 153 (1994) 247– 252. R. Lobinski, C.F. Boutron, J.-P. Candelone, S. Hong, J. Szpunar-Lobinska and F. Adams, Environ. Sci. Technol., 28 (1994) 1467– 1471. M. Heisterkamp, K. Van de Velde, F.C. Adams, C. Ferrari and C. Boutron, Environ. Sci. Technol., 33 (1999) 4416. W. Shotyk, D. Weiss, M. Heisterkamp, A.K. Cheburkin, P.G. Appleby and F.C. Adams, Environ. Sci. Technol., 36 (2002) 3823.

Chapter 35

Sample preparation for chromium speciation Miguel de la Guardia and Angel Morales-Rubio

35.1

THE ELEMENT AND ITS REACTIVITY

Chromium was discovered in 1797 by Vangelis and its name comes from the Greek term “chrome”, i.e. color, due to the large number of colored chromium compounds and the aspect of their aqueous solutions [1]. Chromium has four stable isotopes: 50Cr, 52Cr, 53Cr and 54Cr with relative abundance of 4.31, 83.76, 9.55 and 2.38%, respectively, and ﬁve short-lived radioactive isotopes: 48Cr, 49Cr, 51Cr, 55Cr and 56Cr with half-life values of 23 h, 41.9 min, 2.7 days, 3.5 and 5.9 min, respectively. Element number 24 of the periodic table, Cr, is a metal of the fourth period and group 6 (VIA in the Chemical Abstracts notation). It possesses an external electronic structure 3p63d54s1 and can be found in oxidation states Cr(II), Cr(III) and Cr(VI), being also described as Cr(IV) and Cr(V) compounds [2]. Cr(VI) is a powerful oxidant and can be found in only oxo species such as CrO3, CrO4 22 or FeCrO2. Additionally, it is interesting to indicate the ability of CrO4 22 to be dimerized in acidic media, forming the dichromate ion Cr2 O7 22 through the mechanism indicated in Eq. (35.1) CrO4 22 þ Hþ ! CrO3 ðOHÞ2 CrO3 ðOHÞ2 þ Hþ ! H2 CrO4

ð35:1Þ

2CrO3 ðOHÞ2 ! Cr2 O7 22 þ H2 O Cr(V) and Cr(IV) are found as transient intermediates during the reduction of Cr(VI), but cannot be found in aqueous solution as stable species, being easily decomposed forming Cr(III) and Cr(VI). However, some compounds, like Cr(OC4H9)4 or Ba2CrO4 with a tetrahedral structure, or K2CrF6 with an octahedral structure, have been isolated additionally to that of K2(CrOCl5) with an octahedral structure, or K3CrO8 with a quasi-dodecahedral structure. Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

1115

M. de la Guardia and A. Morales-Rubio

The ion CrO4 32 is present in some alkaline and alkaline earth compounds but these solid compounds are hygroscopic and easily hydrolyzed to Cr(III) and Cr(VI) [3]. Cr(III) is the most stable oxidation state of chromium and can be found in solid octahedral compounds or in aqueous solutions in which the CrOH2þ and CrO2 2 species can predominate at pH values higher than 4 and higher than 12, respectively. The reactivity of Cr(VI) is determined by its oxidizing power. The redox potential of Cr(VI) strongly depends on the pH of the medium, E 0 being 1.33 V in acidic media following Eq. (35.2) Cr2 O7 22 þ 14Hþ þ 6e2 ! 2Cr3þ þ 7H2 O

ð35:2Þ

and 2 0.13 V in basic media through the reduction of chromate to Cr(III) hydroxide (Eq. (35.3)) CrO4 22 þ 4H2 O þ 3e2 ! 2CrðOHÞ3 # þ 5OH2

ð35:3Þ

Thus, the presence of Cr(VI) and its CrO4 22 and Cr2 O7 22 chemical forms strongly depends on the redox potential and pH. On the other hand, CrO4 22 can precipitate easily with Pb(II), Ag(I), Tl(I), Ba(II), Hg(II) and, to some extent, with Hg(I), Sr(II), Cu(II) and Fe(II). From an analytical point of view, the reaction of Cr(VI) with H2O2 in acidic media provides an intense blue color, due to the formation of CrO5 that can be extracted in ethyl ether or amyl alcohol, providing a sensitive and practically speciﬁc means to the qualitative identiﬁcation of Cr(VI) in water or dissolved samples. Additionally, oxidation of diphenylcarbazide by CrO4 22 in an alkaline medium to form diphenylcarbazone and its complexation by Cr(III) can serve to differentiate between Cr(III) and Cr(VI). Chromium (III), formed during the redox process, is necessary to obtain the violet-colored complex. For quantitative purposes, the reaction of Cr(VI) with chromotropic acid (1,8-dihydroxynaphtalene-3,6-disulfonic acid) provides a red color but suffers from some interferences from Sb(III), Sn(II) and Sn(IV) that drastically reduces the sensitivity. On the other hand, Cr(III) is less reactive than Cr(VI); the most common chemical forms are the basic species CrOH2þ and CrðOHÞ2 þ as well as the hydroxide Cr(OH)3 and soluble chromite CrO2 2 of Cr(III). In a weakly acidic medium (pH 2), ethylene diamine tetraacetic acid (EDTA) forms a violetcolored 1:1 complex with Cr3þ . This reaction takes place at 1008C, due to the unfavorable kinetics caused by the high stability of the CrðH2 OÞ6 3þ complex. Additionally, EDTA reduces Cr(VI) to Cr(III), thus providing a false positive identiﬁcation of Cr(III) in samples containing only Cr(VI). Because of this, direct identiﬁcation of the presence of Cr(III) is generally based on a previous determination of the absence of Cr(VI) and formation of this latter species through oxidation [1].

1116

Sample preparation for chromium speciation

35.2

THE PRESENCE OF Cr IN NATURE AND INDUSTRIAL PROCESSES

In spite of the fact that Cr is the seventh most abundant element on earth, its presence in soils is relatively small, being the 21st most abundant element in the earth’s crust. Thus, it is clear that the core and mantle contain a higher concentration on Cr than the crust, being highest in igneous materials. The main chromium mineral is chromite (FeCr2O4), but crocoite (PbCrO4) is also of interest. Additionally, chromium is present in a large number of minerals and rocks to which it provides their typical colors, such as in emeralds, granite, crisobile, amphibole and pyroxenes. The concentration of Cr in geological materials varies with the type and nature of the rocks, being especially abundant in igneous rocks, like serpentine, in which concentrations around 1800 mg kg21 can be found, and also in shales and phosphorites. On the other hand, sediments and especially sands, carbonates and granite, contain low Cr concentrations, of the order of 1 mg kg21. In soils, it has been noted that there is a mean world soil concentration of 50 mg kg21, with large differences as a function of their origin, ranging from 5 to 1000 mg kg21. Additionally, Cr in soils is neither easily extracted with water, nor with reducers or complexing agents. As an example, traditional studies based on sequential chemical extraction have shown that the use of a three-stage extraction procedure based on the use of: (i) 1 M ammonium acetate at pH 5, (ii) 1 M hydroxylamine in 25% acetic acid and (iii) 1 M HNO3 at 1208C, are necessaries for speciation of Cr in marine suspended particulate matter [4]. Because of the reduced mobility of Cr in the environment, the typical concentration of Cr in sweat and seawater is at the mg l21 level [5]. It is well accepted that Cr is an essential element for animals and humans but it is not clear what effect it has on plant growth. The Cr concentration in plants is limited by the low concentration of this element in fresh waters and soils and its difﬁculties in removing it from the soil. There are increasing Cr concentrations in plant roots when agricultural amendments obtained from sewage sludges are employed. In fact, the application of the classical ﬁve-step Tessier method to the speciation of Cr in compost samples showed that 50% of Cr remained in the inert fraction of insoluble compounds and that the main form of Cr in compost was Cr(III) with Cr(VI) being less than 12% [6]. Chromium metal is commonly obtained from the reduction of chromate by coal in furnaces in order to obtain a ferrochrome alloy. On the other hand, chromite, treated with fused alkali and oxygen, can be oxidized and sodium dichromate precipitated. The reduction of this latter compound by carbon provides Cr2O3, which is reduced by aluminum to obtain pure chromium, a white, hard and brittle metal that melts at 1890 ^ 108C. Chromium metal in its zero oxidation state is soluble in non-oxidizing acids but can neither be dissolved by nitric acid nor by aqua regia, in spite of the fact that the reduction

1117

M. de la Guardia and A. Morales-Rubio

potential of Cr(II) is 20.91 V and that of Cr(III) is 2 0.74 V. This phenomenon is probably due to the passivation of the metal through the formation of a superﬁcial layer of oxide. This point seems to be conﬁrmed by the high stability of the green chromium oxide, which is frequently used in paints. Metallic chromium is used in the fabrication of special steels, frequently in the production of stainless steel associated with Ni and, in some cases, with W, V, Mo or Co. Chromium is frequently employed in the production of electrodes. The special alloy Nicrom is commonly used in lamps and associated with B or Al. In the paint industry, Cr2O3 is employed as a green pigment and BaCrO4 and PbCrO4 as yellow pigments. Cr2O3 and Cr salts are commonly used in the formation of electroplating baths. K 2Cr 2O 7 is an important oxidant employed as a reagent in laboratories and for dying in the ceramic industry. The use of potassium dichromate in leather tanneries is well known. Chromium acetate is used in the textile industry as a cleaning product and other compounds are used as catalysts, wood preservatives or components in magnetic tips. At the mg levels, Cr is essential for normal glucose and lipid metabolism, and as an insulin receptor [7]. Because of that, the World Health Organization (WHO) has recommended a daily intake of 10–40 mg, based on reported intake data that varies from 5 to 500 mg per day, depending on the diet composition around the world. However, it seems conﬁrmed that Cr(VI) is a carcinogen and irritant and can cause bronchitis, pneumonia, hypersensitivity and gastrointestinal hepatic and renal impairments [8] and, because of this, there is a need for speciation studies in water and foods to establish correctly the safe limits of Cr for human nutrition. 35.3

CHEMICAL SPECIES OF Cr PRESENT IN REAL SAMPLES

In addition to the determination of dissolved and particulate chromium, which is a typical assay conducted during fresh water analysis, the major studies regarding Cr speciation are focussed on the differentiation between the two most stable oxidation states. In aqueous systems, Cr2þ cannot be present at redox potential values higher than 20.4 V in strongly acidic media or between 20.5 and 2 0.8 V in the pH range 4–6, as it is precipitated as Cr(OH)2 for pH values higher than 6.2 [9]. Cr(VI) predominates as Cr(OH)4 between pH 3.5 and pH 9.5 in a reduced redox potential range. The speciﬁc chemical forms of Cr(III) and Cr(VI) strongly depend on the redox and pH values, being controlled, in the absence of complexing agents, by the following equilibria for Cr(III) species: CrðOHÞ3 # þ 3Hþ ! Cr3þ þ 4H2 O

log K ¼ 9:76

ð35:4Þ

CrðOHÞ3 # þ 2Hþ ! CrðOHÞ2þ þ 2H2 O

log K ¼ 5:91

ð35:5Þ

1118

Sample preparation for chromium speciation

CrðOHÞ3 # þ Hþ ! CrðOHÞ2 þ þ H2 O

log K ¼ 20:44

ð35:6Þ

CrðOHÞ3 # ! CrðOHÞ3

log K ¼ 26:84

ð35:7Þ

CrðOHÞ3 þ H2 O ! CrðOHÞ4 2 þ Hþ

log K ¼ 20:44

ð35:8Þ

illustrating clearly that, under reducing conditions, Cr3þ is the dominant species up to pH around 3 as CrðOHÞ2þ is the main Cr(III) species in the pH range between 3.5 and 5. The species CrðOHÞ2 þ is difﬁcult to identify in the presence of covalent Cr(OH)3 due to the ease of precipitation of the hydroxide in the pH range 5–13 necessary to form CrO2 2 . On the other hand, Cr(VI) species can be protonated, H2CrO4 and HCrO4 2 , deprotonated CrO4 22 or dimerized Cr2 O7 22 as a function of pH according to the equilibria: H2 CrO4 ! HCrO4 2 þ Hþ

log K ¼ 20:74

ð35:9Þ

HCrO4 2 ! CrO4 22 þ Hþ

log K ¼ 26:49

ð35:10Þ

2HCrO4 2 ! Cr2 O7 22 þ H2 O

log K ¼ 1:52

ð35:11Þ

Biomethylation, which is a typical mechanism of metal mobilization or metal decontamination in sediments and biota, can produce CH3Cr(H2O3)2þ by reaction of Cr(II) with methylcobalamine through an homolytic pathway. However, the methylchromium is easily decomposed by acids to form Cr(III), with a half-life of this compound being 0.05 min. The essential activity of Cr(III) in the metabolism of lipids and glucose opens the possibility of ﬁnding Cr associated with speciﬁc molecules such as the glucose tolerance factor (GTF). However, the reduced concentration of Cr in serum and biological matrices (on the order of 0.2 ng ml21) offers terrible difﬁculties to establish the actual species present in natural samples, and there is evidence only for the presence of Cr(III) associated with transferrin and albumin. Cr(III) can be complexated by ammonia, but chemical forms of the type of CrðNH3 Þ6 3þ require a high NH4OH concentration and are easily decomposed to yield Cr(OH)3. However, organic anions, such as citrate, tartrate and sugars and polyhydroxylated compounds in general, present a high capacity for Cr(III) complexation and Cr(III) stabilization in front of oxidative processes and, because of that, in samples containing large amounts of organic matter, organic compounds of Cr(III) must also be considered other than aquocomplexes, free or partially hydrolyzed ions. All the aforementioned considerations are based simply on thermodynamic data and, as has been pointed out [10], kinetic and complex system variables must also be taken into consideration.

1119

M. de la Guardia and A. Morales-Rubio

35.4

ANALYTICAL METHODOLOGIES AVAILABLE FOR DETERMINATION OF Cr

All the available modern analytical techniques, from those based on radioactivity to those employing electroanalytical, separation or spectrometric methods, provide appropriate sensitivities for the quantitative determination of total Cr at mg l21 levels [11]. However, in order to undertake Cr speciation studies, the limits of detection (LOD) must move to the mg l21 and ng l21 levels and, because of that, typical LOD values of 5000 ng ml21 for neutron activation analysis (NAA), 500 ng ml21 for X-ray ﬂuorescence (XRF) or 20–200 ng ml21 for ﬂame atomic absorption limit their application to the determination of minority species of Cr in real samples. Polarography, with a LOD of 10–50 ng ml21, provides a good means of speciation of Cr in dissolved samples and spectrometry, with LOD values between 2 and 20 ng ml21 can also be applied to water analysis. Inductively coupled plasma optical emission spectrometry (ICP-OES) with a LOD of 1.5 ng ml21, total reﬂection XRF (LOD ¼ 0.2 –1 ng ml21) and gas chromatography, based on the use of triﬂuoroacetylacetone complexes and electron capture detector (LOD ¼ 0.1 ng ml21 for Cr(III)) provide speciﬁc applications in Cr speciation. Amongst the most sensitive techniques for Cr determination, chemiluminescence (LOD ¼ 0.03 –0.3 ng ml21), electrothermal atomic absorption spectrometry (ET-AAS) (LOD ¼ 0.05 – 0.15 ng ml21) and inductively coupled plasma mass spectrometry (ICP-MS) (LOD ¼ 0.02– 0.05 ng ml21) offer excellent alternatives for both the total determination of Cr at ultratrace levels in macro- and microsamples and the best tool for the determination of minor species. On the other hand, the catalytic effect of Cr(III) on the oxidation of luminol by H2O2, which is in the basis of its chemiluminescence determination, offers a means for the direct determination of traces of Cr(III) in the presence of Cr(VI). Due to their intrinsic selectivity, in the presence of matrix and interelemental interferences, and their possible application as detectors in chromatographic separations, ET-AAS and ICP-MS are the methods of choice in applied studies, and ICP-MS coupled to liquid chromatography is currently the most powerful instrumentation to examine the presence and distribution of Cr species in samples previously dissolved or leached with appropriate reagents. However, the omnipresence of Cr in the environment and the low concentration of this element in actual samples give rise to serious problems of contamination [12]. 35.5

ANALYTICAL METHODOLOGIES FOR Cr SPECIATION IN WATER

Chromium species in waters can be present in both dissolved states or associated with the suspended particles. When the concentration of Cr species

1120

Sample preparation for chromium speciation

is very low, they may undergo alteration during sample handling and analysis, especially when preconcentration procedures are required. Atomic spectrometric detection, both alone or coupled with other techniques to preconcentrate and separate the two main chromium species, Cr(III) and Cr(VI), is the most widely employed methodology reported in the literature. 35.5.1 Speciation of chromium in aqueous matrices Cr(III) and Cr(VI) are frequently determined in waters. However, in the majority of such studies, only a single Cr species was determined together with the total content of Cr, and thus the second species was established by difference. Figure 35.1 shows a schematic presentation of the relative use of atomic spectrometry, molecular spectrophotometry and voltammetry for the determination of Cr(VI), Cr(III) and total chromium (Cr) in the analytical literature compiled from the Analytical Abstracts database for the period 1980 –2001. Complete speciation studies, involving the determination of Cr(III) and Cr(VI) additionally to total Cr was done in approximately 1% of studies in spite of the analytical technique employed. Moreover, Cr(VI) and Cr(III) were determined in 27% of atomic spectrometry-based studies, in 10% of papers involving the use of molecular spectrometry and only in 8% of cases in which electroanalytical determinations were done. The determination of one Cr species and total Cr concentration was undertaken in 10, 13 and 17% of atomicbased, molecular-based and electroanalytical studies, respectively. 35.5.2 Types of samples analyzed Chromium has been determined in all types of water samples, including natural waters and standard or synthetic preparations. The most frequently analyzed samples are efﬂuents, polluted, rinse and waste waters such as residual water from cement, galvanic, plating, sewage treatment, tannery or wood industries. The high concentration of chromium in this type of waters and the presence of speciﬁc forms of Cr as a function of the process involved explain the great number of articles (56%) using molecular spectrophotometry, in front of the most sensitive atomic spectrometric techniques (29%). In most cases, these waters reach the sea, rivers, lakes, etc., and it constitutes a serious contamination risk. On the other hand, it is important to study the contamination levels of natural waters because it affects the safety of drinking waters. Because of that, natural water and bottled, drinking, fresh, mineral, potable, spring, tap and well waters are the second group of the most studied systems. Moreover, the low concentration of chromium in this kind of water also explains why the principal detection techniques employed are atomic spectrometry-based ones

1121

M. de la Guardia and A. Morales-Rubio

Cr(VI) N = 190 30.8%

Cr(VI)+Cr(III) N = 112 18.1%

Cr(III) N = 57 9.2%

Cr N = 174 28.3%

Atomic Spectrometry

Cr(VI) N = 29 10.0%

Cr(VI)+Cr(III) N = 77 26.6%

Cr I)+ (II 26 Cr N = % 4.2

Cr (V N I)+ 8.3 = 51 Cr %

Cr(VI)+Cr(III)+Cr N=7 1.1%

Molecular Spectrometry

Cr(VI) N = 128 54.7%

Cr(III) N = 14 4.8%

Cr (V N I)+ 10 = 24 Cr .2%

Voltammetry Polarography Cr(VI) N = 25 39.7%

Cr(VI)+Cr(III) N=5 7.9%

Cr I)+ 6 (II Cr N = % 2.6

Cr I)+ (II 16 Cr N = % 4.5

Cr (V N I)+ 5.9 = 17 Cr %

Cr(III) N = 35 15.0%

Cr(VI)+Cr(III)+Cr N=1 0.4%

Cr(VI)+Cr(III)+Cr N=4 1.4%

Cr N = 132 45.8%

Cr(VI)+Cr(III) N = 23 10.0%

Cr N = 17 7.3%

Cr(III) N=5 7.9%

Cr I)+ (II 1 Cr N = % 1.6

Cr (V N I)+ 15 = 10 Cr .9%

Cr(VI)+Cr(III)+Cr N=1 1.6%

Cr N = 16 25.4%

Fig. 35.1. Distribution of the chemical forms of chromium in waters determined in the articles reviewed by using different detection techniques. N: number of reports; Cr(VI): chromium VI species; Cr(III): chromium III species; Cr: total chromium.

for both, natural water (49%) and bottled, drinking, fresh, mineral, potable, spring, tap and well waters (57%). In water samples, in which the chromium concentration remains at low levels, atomic spectrometric detection is the most frequently employed analytical methodology. In 67% of the studies that analyzed brackish, coastal, estuarine, harbor, ocean and sea waters; in 51% of determinations in canal, river and stream waters; in 63% of certiﬁed, reference, synthetic and standard waters; in 67% of dig, ground, environmental, surface and underground waters

1122

Sample preparation for chromium speciation

and in 61% of lake, pond and reservoir waters. Rain and snow waters were analyzed mainly by molecular spectrophotometric techniques.

35.5.3 Pretreatments and techniques applied The most frequent pretreatments employed for speciation of chromium are complexation (39%) and preconcentration by column/ionic exchange (22%) of Cr(VI) and/or Cr(III). Some 10% of the pretreatments are based on oxidation of Cr(III) to Cr(VI) and further determination of total chromium as the oxidized species. On the other hand, reduction pretreatments have been employed as well to transform Cr(VI) to Cr(III) and to undertake the determination of total chromium as Cr(III). In quite a number of cases, one species and total chromium concentration were determined together and the concentration of the other species is calculated by the difference between them. The most frequently used pretreatment procedure used with atomic spectrometric detection techniques is preconcentration on columns, which permits the retention and subsequent elution of one or two species. In some cases, one of the two Cr species needs complexation before separation in a column. Complex formation is the most widely applied pretreatment for speciation of chromium, and depending on its oxidation state, chromium can form different complexes. Most of the complexes formed are colored compounds that can be measured by molecular spectrophotometry. For that reason, the percentage for complex formation pretreatments in atomic spectrometric detection techniques is smaller than for the remaining techniques. Liquid extraction processes are frequently used after complex formation to extract the complexes formed prior to their detection. With respect to the techniques employed to determine Cr(VI) and/or Cr(III), atomic spectrometries are those most often employed (48.5%), these being graphite furnace atomic absorption spectrometry (GF-AAS) (15.8%), ﬂame atomic absorption spectrometry (F-AAS) (9.2%), ICP-OES (6.1%), XRF and proton-induced X-ray emission (PIXE) (3.8%), NAA (2.1%), atomic emission spectrometry (AES) (1.3%) and those hyphenated with chromatography (3.6%). In this group, ICP-MS (3.8%) is included along with its hyphenation with chromatography (2.8%) in spite on the fact that MS is an ion measurement technique and not an atom measurement methodology. Molecular spectrometry-based detection is the second most frequently employed (36.3%) group for Cr speciation, with UV–VIS being the most extensively used (26.6%) followed by chemiluminescence (3.1%), ﬂuorimetry (2.6%) and those procedures hyphenated with chromatography (4%). Other techniques, such as voltammetry (4.6%), polarography (2.8%), potentiometry (1.2%) or chromatography (1.7%) have also often been employed to determine Cr(VI) and Cr(III) in water samples.

1123

M. de la Guardia and A. Morales-Rubio

35.5.4 Speciation chromium using atomic spectrometry and MS-based techniques The literature on chromium speciation using these techniques has been classiﬁed into three groups, according to the different pretreatments employed, taking into consideration whether they were accomplished in the solid or liquid phase: group 1—preconcentration on column, ion exchange, adsorption and solid phase extraction; group 2—complex formation, oxidation –reduction, liquid extraction; group 3—precipitation, ﬁltration, centrifugation, acid digestion, microwave-assisted treatments. There are more than 90 studies [13– 102] published using ion exchange, adsorption or preconcentration for the determination of Cr(III) and Cr(VI) as individual species. In this group, preconcentration on columns is the most common treatment. Two different preconcentration strategies have been employed: (A) only one species is preconcentrated, Cr(III) or Cr(VI), while the other species is often complexed or measured directly and (B) both species are preconcentrated, either at the same point or in different places. A summary of the analytical features, derived from the various published atomic techniques, is presented in Table 35.1. For concentrated samples, techniques such as ion chromatography directly coupled plasma atomic emission spectrometry (IC-DCP-AES), F-AAS and HPLC-F-AAS or even HPLC-ICP-MS are recommendable because they can provide the greatest linear range. On the other hand, GF-AAS, ICP-OES, ICP-MS or NAA could be adequate for low levels of chromium species in water samples, as can be seen in Table 35.1, achieving very low LODs. RSD values are of the same order for both direct determinations by atomic spectrometric techniques and those hyphenated with chromatography, the worst being those using GF-AAS. Finally, recovery percentages found for spiked samples are near to 100% in most cases, providing evidence for the absence of analyte losses or contamination using these procedures. 35.5.4.1 Preconcentration of Cr(VI) Preconcentration of Cr(VI) has been accomplished using different types of columns, such as melamine–formaldehyde resin [103] or with melamine – urea–formaldehyde resin [104], a C18-bonded silica reversed phase sorbent with DDTC as the complexing agent [105,106], a glass column containing immobilized sodium diethyldithiocarbamate or ammonium hexamethylenedithiocarbamate on silica gel [107], a column of chromabond NH2 [108], a column containing phosphate-treated sawdust as adsorbent [109], a column containing ZnO (also used to retain Cr(VI)) [110], a column containing TiO2 [35, 111], a column with chitosan [112] or with charcoal [113] and minicolumns such as an alumina microcolumn [42,52] or adsorption of APDC – Cr(VI) complex on polyether ether ketone [114] or polytetraﬂuoroethylene tube reactors [115]. Different types of resins, such as anion exchange resins [76,116,117], resins

1124

TABLE 35.1 Figures of merit for atomic techniques used for chromium speciation in natural waters Technique

Range (ng ml21)

Cr(VI)

Cr(III)

Total Cr

Cr(VI)

Cr(III)

Total Cr

0.8–200 0.01–0.05 0.01–15 0.2–61 0.06–5.5 – 0.02–80 2 0.02 0.6 2–30 35–270 – 0.1–88 0.1–0.5 13–300 0.03

0.2– 80 0.025– 0.3 0.01– 80 1.4– 45 0.06– 0.3 – 0.025– 40 2 0.025 0.6 2– 10 12– 250 – 0.06– 81 0.2– 0.3 42– 400 0.03

0.003– 250 – 0.003– 10 0.2– 2.5 0.2 2– 3 0.8 – – – 4 2 1 0.03– 1 – – –

0.2 –50,000 ,10,000 0.1 –5000 100 –10,000 1 –90 – 100 –10,000 75–350 – ,700 – 5 –100 – 1 –120,000 0.5 –500 ,4500 –

0.2 –1100 0.01–100 0.8 –5000 0.2 –10,000 1 –250 – 100 –10,000 75–350 – ,700 – 5 –100 – 1 –120,000 0.3 –200 ,4500 –

75–7500 – 0.15–300 0.2 –1000 – ,1000 5 –106 – – – 50–250 20–40,000 100 –100,000 0.5 –5000 1 –100 – –

continued

Sample preparation for chromium speciation

F-AAS F-AES GF-AAS ICP-AES ICP-MS DCP-AES HPLC-F-AAS IC-F-AAS HPLC-F-AES HPLC-GF-AAS HPLC-ICP-AES IC-ICP-AES IC-DCP-AES HPLC-ICP-MS IC-ICP-MS XRF NAA

Limit of detection (ng ml21)

1125

1126

TABLE 35.1 (continuation) Technique

Recovery (%)

Cr(VI)

Cr(VI)

0.52–7 2.1–2.4 0.04–12.5 0.8–402 1.2–8 – ,2 3.5 2.1 2.2 1.3–3.8 – – 1.3–3.1 1.2–5 1–7 –

Cr(III) 0.67– 5.9 0.3– 5.6 0.03– 5.8 2.2– 4.2 2.3– 4.4 – ,2 9.6 5.6 3.2 2.3– 5.6 – – 0.9– 3.5 2.3– 5 1– 7 –

Total Cr 1.2– 16 – 1.8– 24.7 0.8– 12 0.2– 3.9 – 1– 6.2 – – – 10 0.3– 1.6 – 2– 5 4.6– 8.3 – 1.1– 5.7

100 –106 99.8 85.4 –97.9 93–103 90–110 – – – 95–98 – 94–96 – – 91–100.8 96–114 84–103 95–105

Cr(III) 93–103 – 87.4–102.4 93–103 90–110 – – – 95–98 – – – – 87 87 84–103 95–105

Total Cr 87–106 95–98 85–114 93–105 90–106 – – – – – .98 – – 98.9–115 – – 95–105

M. de la Guardia and A. Morales-Rubio

F-AAS F-AES GF-AAS ICP-AES ICP-MS DCP-AES HPLC-F-AAS IC-F-AAS HPLC-F-AES HPLC-GF-AAS HPLC-ICP-AES IC-ICP-AES IC-DCP-AES HPLC-ICP-MS IC-ICP-MS XRF NAA

Relative standard deviation (%)

Sample preparation for chromium speciation

with Amberlite diluted in isobutylmethyl ketone (IBMK) [118,119], Chelex 100 chelating resin [120] and liquid anion exchangers such as Amberlite LA-1 or LA-2 [121] have also been used. The pH values employed for the retention of Cr(VI) vary according to the different methodologies from highly acidic [59,86,103–105,109,113] to slightly acidic [42,107,120] and even neutral media [64]. Different solvents were used to elute Cr(VI), such as HNO3 [108,120], NH4OH [52], NaOH [112], ethanol [106,115], sodium acetate [103] or IBMK [59, 114]. In some cases, Cr(III) is oxidized to Cr(VI), total chromium measured, and the Cr(III) content calculated as the difference between total Cr and Cr(VI) [39, 104,105,112]. In other cases, Cr(III) is measured directly [42,52,118,119] or is precomplexed with diphenylcarbazide [22] or with EDTA [76].

35.5.4.2 Preconcentration of Cr(III) Preconcentration of Cr(III) has been accomplished using other types of columns than those employed for Cr(VI), including chelating resin PAPHA [130], with quinoline-8-ol or Muroac A-1 resin [122,123,135], with poly[4-(1-azo-4carboxyphenyl)styrene] [128], with polystyrene divinylbenzene resin [49] or with chelating ion-exchange resin having salicylic acid functional groups [79], Sephadex [126], a reversed phase C18 column [20] and anion exchange columns of Muromac A-1 [123]. Activated alumina can be introduced in an electrochemical cell [134] or in a microcolumn [136]. Columns containing cellulose phosphate [129], ZrO2 [139], TiO2 [30,35], SiO2 [36], activated carbon [125], zeolites [143], Chitosan [132,133], dealginated seaweed biomass [63] and Saccaromyces cerevisiae immobilized on sepiolite [58] have been used for Cr(III) preconcentration. Resins such as the Bio-Rad ion-exchange AGMP-1 after complexation with sulfonated azo-dyes [136], Amberlite XAD-2000 [140], Amberlite XAD-16 [144], exchangeable resin Chelex-100 or Lewatit TP 207 [127] or ion exchange resin [136] can also be used. Finally, chelating collectors such as polyacrylamide [131], polyacrylamidazone-hydrazide lacmoid chelating ﬁber [141] or synthetic ettringite [73] can be used for preconcentration of Cr(III). From the solvents used to elute Cr(III), HCl [30,128 –130,134], HNO3 [122, 123,127,132,133,135,136,138,139], HNO3 in acetone [125,140] and HCl/H2SO4/ HNO3 [141] are the most frequently employed. The pH for the retention of Cr(III) varies according to the process from slightly acidic [20,59,111,122–124, 126,127,130,131] to basic [73,134 –136]. In some cases, reduction of Cr(VI) to Cr(III) is undertaken, total chromium measured and the Cr(VI) content calculated from the difference [49,122,124, 130,134,135]. In other cases, Cr(III) is precomplexed with quinoline-8-ol [49] or with sulfonated azo-dyes [136].

1127

M. de la Guardia and A. Morales-Rubio

35.5.4.3 Preconcentration of both species Retention of Cr(VI) and Cr(III) at the same time Cr(VI) and Cr(III) can be retained on different types of columns packed with materials such as strongly basic anion exchangers with H2SO4 [99], SAX [89], Dionex AG4A [94], DEAE-Sephadex A-25 [61], silica gel loaded with anion exchanger Adogen 464 [81], polyacrylonitrile sorbent modiﬁed with polyethylene polyamine [78], C18 [19,50,70], Eurospher 100-C18 [23,39], Eurosphere RPC18 [15], Dionex CS5 or Dionex ASH [145], anion exchanger Omni Pac PA 100 [146], Dionex ion Pak A57 [67], anion exchanger Ion Pac A65 [65], Excelpak ICS-A23 [28] and Polyspher IC AN anion exchanger [91]. Cr(VI) and Cr(III) can be also retained on a microcolumn packed with activated alumina [36,44,74,92,147], TiO2 [36,64], methyltrioctyl ammonium chloride-loaded silica gel [60] or polymeric Detata sorbent with aminocarboxylic groups [137]. The pH values for retention of the Cr chemical forms vary depending on the species and the procedure employed. Cr(III) can be retained at acidic pH [44,59, 60,70,81,92,99,137], and Cr(VI) can be retained at acidic pH [44,70,81,86,92,99, 137] and in basic pH solutions [60]. The most commonly used solvents for elution of chromium species include HNO3 [44,61,64,74,94,147], water [92], HClO4 [60], HCl [78,137], H2SO4 [99] and methanol [16,19,39,70,86] for Cr(III); and NH4OH [44,64,74,147], boiling water [92], HClO4 [60], HCl [137], methanol [19,39,70,86], IBMK [16], ascorbic acid [99], Na2CO3/NaHCO3 [94] and hydroxylammonium chloride [61] for Cr(VI). Based on these factors, it is possible to do true speciation by an appropriate selection of the experimental variables, thus making the preconcentration of both Cr(III) and Cr(VI), simultaneously or sequentially, with sequential elution. Retention of Cr(VI) and Cr(III) on various supports Cr(VI) and Cr(III) can be sequestered from the same sample using different supports. This can be accomplished using PTFE columns with anion-exchange resins and activated alumina [41] or with a cation-exchange resin column containing Amberlite IR-120 to retain Cr(III) and with Amberlite IRA-400 to retain Cr(VI) [100]. With a C18 column, Cr(III) can be retained and a polyether ether ketone loop can be used to retain the APDC –Cr(VI) complex [16]; with a C18 column with potassium hydrogen phthalate to retain Cr(III) and a Eurospher 100-C18 column to retain Cr(VI) [21]; with a Dowex 50W-X8 ion exchange resin to retain Cr(III) and with a Dowex 1-X8 ion exchange resin to retain Cr(VI) [59]; with an anion-exchange column with Chelex 100 resin to retain Cr(III) and with AG-MP-1 anion resin to retain Cr (VI) [87]. Anionexchange resins can be used to retain Cr(III) and a cation-exchange resin to retain Cr(VI) [45,88]. Cellulose with Cellex P can retain Cr(III) and Cellex T can retain Cr(VI) [142]. A mixture of Anionite AV-17-8 can be used to retain Cr(III) and a mixture of Cationite KB-4 to retain Cr(VI) [47]. A microcolumn with

1128

Sample preparation for chromium speciation

IDAEC chelating cellulose can retain Cr(III) and Cr(VI) with DEAE chelating cellulose [57]. An aromatic sulfonic acid silane column can be used to retain Cr(III) and a Bio-Rad 1-X4 column to retain Cr(VI) [25]. A PAAO resin can be used to retain Cr(III) and a PDTC resin to retain Cr(VI) [148]. An IC-PAKA anion exchanger column can retain Cr(III) and an IC-PAKA cation exchanger column can retain Cr(VI) [96]. A Separon SGC C18 column can retain the Chromazurol S complex of Cr(III) and a Separon SGC C18 column can retain the diethyldithiocarbamate complex of Cr(VI) [86]. The pH for retention of the species varies depending on the chemical forms and the procedure employed: Cr(III) and Cr(VI) at pH 2 [59]; Cr(III) and Cr(VI) at pH 7 [64]; Cr(III) at pH 3.6 and Cr(VI) at pH 0 [86]; Cr(III) at pH 3.8 and Cr(VI) at pH 3 [21]; Cr(III) at pH 2–2.5 and Cr(VI) at pH 3.5 –4.5 [25]; Cr(III) at pH 4–0.8 and Cr(VI) at pH 4–6 [124]. The most commonly employed solvents for elution are HNO3 [41,64,87], HCl [88,100,142], methanol [16,21,86] and NH4OH/HNO3 [25] for Cr(III), and HNO3 [25], NH4OH [41,64], NH4NO3/NH4OH [87], NaOH [142], NaCl [100], methanol [21,86] and IBMK [59] for Cr(VI). 35.5.4.4 Complexation of Cr(VI) and Cr(III) Chemical treatments, like complexation, oxidation or reduction and liquid extraction, have frequently been employed for speciation of chromium where Cr(III) and Cr(VI) are determined as individual species. In this group, the most common pretreatment is complex formation, but this can be accomplished with liquid extraction of the complexes and/or pre-oxidation or pre-reduction pretreatments. Different complexing agents are used, depending on whether the species is Cr(VI) or Cr(III). Cr(VI) can be complexed with different dithiocarbamates, such as ammonium pyrrolidine dithiocarbamate [16,69,151,155 – 158], pyrrolidine1-carbodithioate [159 – 161], ammoniun hexamethylene dithiocarbamate [107], alkylene bisdithiocarbamates (BDTCs) [162], dibenzyldithiocarbamate [152,163,164], Mn(II)-diethyldithiocarbamate [46], Na-diethyldithiocarbamate [86,107,165], trioctylphosphine oxide in hexane [166], benzyl(dodecyl)dimethylammonium bromide [106], chloride [152], ammonium triﬂuoroacetylacetonate [167], diphenylcarbazide [168], propylxanthate [169] and tetrabutylammonium [170]. Cr (VI) can be extracted as an anion-pair complex with dicyclohexyl-18crown-6 in 1,2-dichloroethanol [150]. Cr(III) can be complexed with different hydroquinolines, such as hydroxyethylpiperazine-N0 -3-propanesulfonic acid buffer/methanolic-8-hydroxyquinoline [171], Pd/8-quinolinol/tanic acid [172], hydroxyquinoline [69,173] and methanolic quinoline-8-ol [174]. Triﬂuoroacetylacetone [167,175], Chromazurol S [86], 1-(2-pyridylazo)-2-naphthol [140], pyrocatechol violet [125], EDTA [89], SCN2 [152], Na octyl sulphate [154] and diethyldithiocarbamates [46] have also been used. The pH values for the complexation vary from 2 [158] to 8.5 [46].

1129

M. de la Guardia and A. Morales-Rubio

For the extraction step, IBMK is the most common solvent used [155 –157, 159– 161,169,171,173,174], followed by CHCl3 [165,170], hexane [166], tributylphosphate [176] and 1,2-dichloroethane [150]. In some cases, Cr(III) is oxidized to Cr(VI) and total chromium measured as Cr(VI) [151,165,168,176] or Cr(VI) is reduced to Cr(III) and total chromium measured as Cr(III) [38,149,153,171]. 35.5.4.5 Precipitation of Cr(VI) and Cr(III) The precipitation of chromium species is normally used as a complementary tool for preconcentration and complexation of Cr(VI) and/or Cr(III). Cr(VI) can be precipitated with PbSO4 at pH 3 or with Pb3(PO4)2 in a basic medium [177]; with ZnSO4 and diethyldithiocarbamate at pH 3.5 [179] or with diethyldithiocarbamate at pH 3 [180]; with ammonium pyrrolidinediethylcarbamate at pH 2 for subsequent solvent extraction by IBMK [16,114,115,156] and with Al(OH)3 at pH 9.5 [38]. Cr(III) can be precipitated with Anabaena algae [27]. In this case, Cr(VI) is measured in the supernatant solution after centrifugation of the sample. Cr(III) precipitation can be done with NH3 at pH 5.5 –6.5 [178], with Fe(OH)3 at pH 9.8 [181] and pH 10 –10.5 [182]; with La(OH)3 at pH 9.5 [120,183], with Mg(OH)2 at pH 3 [184], with gallium in H3PO4 at pH 5 [185] and with gallium nitrate at pH 9.3 –10 [184,186]. In the latter two cases, Cr(VI) is determined by difference between total chromium (calculated after reduction of Cr(VI) to Cr(III) and measured as before) and Cr(III). Cr(III) precipitates with Al(OH)3 at pH 5.5 –6.5 [187] or pH 9 [38]. Precipitation by means of Pd/8-quinolinol/tannic acid complex at pH 5.1 –5.3 [172] and other complementary treatments, such as ﬁltration [48,180,188], centrifugation [22,184] and acid digestion with microwave irradiation [74], have also been used. 35.5.5 Determination of chromium speciation using molecular spectrophotometry From the literature on chromium speciation in water using molecular spectrometry [189– 210], it is evident that complexation is the most frequently employed treatment for determination of chromium species, followed by the direct oxidation of reagents or catalyzed by Cr(VI). Amongst molecular spectrometric techniques, UV–VIS absorption is clearly the most employed, covering 85% of the publications. Table 35.2 summarizes the analytical features, obtained using different molecular techniques, for determination of chromium. For samples having high chromium concentration, methods employing UV–VIS detection are the most recommended because they can provide a high linear range. Moreover, chemiluminescence can be adequate for low levels of chromium species in the samples because of the very low LOD

1130

TABLE 35.2 Figures of merit for molecular techniques used for chromium speciation in natural waters Technique

Cr(VI)

Cr(III)

0.04– 24 0.8– 3.3 0.01– 28 0.02– 2 1.8– 40 0.1– 300 10

0.02–190 1 0.01–0.5 0.1–2 2.5–40 0.05–100 50

Range (ng ml21) Total Cr 12–50 0.9 0.04– 1 0.5– 2 40 – –

Relative standard deviation (%)

UV– VIS Fluorimetry Chemiluminescence HPLC-UV– VIS IC-UV– VIS IC-chemiluminescence CE-UV– VIS

Cr(VI)

Cr(III)

Total Cr

2 –100,000 1 –200 0.1 –30,000 0.05 –9600 10–150,000 1 –500 10–100,000

30–1200 1 –150 0.01–800 5 –5000 10–150,000 1 –500 50–50,000

50–2000 1 –60 0.01–10,000 5 –5000 10–150,000 1 –1000 ,100

Recovery (%)

Cr(VI)

Cr(III)

Total Cr

Cr(VI)

Cr(III)

Total Cr

0.4– 4.1 1.2– 6 1.8– 5 1.8– 6 1– 3 2.7– 4.5 1– 4.5

0.9–2 1.6 0.4–6.8 2.1–6 – 2.1–4.3 1.3–4.5

0.8–8 – 1.2–6 0.2–5 – – ,4.5

87–108 96–105 92–105 91–104 .95 – 98–103

85–117 – 92–108 106 – – 98–103

85–115 – 94–110 – – – 94–103

Sample preparation for chromium speciation

UV– VIS Fluorimetry Chemiluminescence HPLC-UV– VIS IC-UV– VIS IC-chemiluminescence CE-UV– VIS

Limit of detection (ng ml21)

1131

M. de la Guardia and A. Morales-Rubio

levels achieved. RSDs are of the same order for all molecular spectrophotometric techniques, even if they are hyphenated with chromatography. Recoveries are near to 100% in most cases, and the wide range in some reports is a consequence of the large number of reagents employed. 35.5.5.1 Determination of Cr(VI) using UV– VIS Determination of Cr(VI) has been accomplished using complexation with different reagents, such as diethylaminophenol [211–213], ﬂuorone compounds [203,214– 218], rhodamine [219,220], tetramethylbenzidines [221,222], diantipyrinyl compounds [223 –233], ammonium pyrrolidine-1-carbodithioate [210], phenylenediamine compounds [234,235], diphenylbenzamidine compounds [236,237], aminodiethylaniline [238,239], diphenylcarbohydrazide [240,241], chrome azurol S [242,243], xylenol orange [244] malachite green [245], (5bromo-2-thiazolylazo)resorcinol [246], 4-(6-methyl-2-benzothiazolylazo)phloroglucinol [247], dithizone [249], diphenylhidrazine [250], N-(1-naphthyl)ethylenediamine dihydrochloride [251] and 2-(2-thiazolylazo)-5-diethylaminobenzoic acid [314]. Complex formation varies from the simplest 1:1 form, as with Cr:2(3,5-dibromo-2-pyridylazo)-5-diethyl aminophenol [213], to the ternary ones 1:2:2, as Cr:salicylﬂuorone:TDPC [214], Cr:2-nitrophenylﬂuorone:HDTMAB [217] or CrO2:N1-hydroxy-N1N2-diphenylbenzamidine:N-arylacetamide [236]. In some procedures, surfactants such as cetyltrimethylammonium bromide (CTMAB) [203,228,242], tetradecyltrimethylammonium hydroxide (TDTMAH) [201], hexadecyltrimethylammonium bromide (HDTMAB) [216,217,248], cetyldimethylaminoacetic acid (CDMAA) [215], tetradecylpyridinium chloride (TDPC) [214], hexadecylpyridinium bromide (HDPB) [243], tetrabutylammonium bromide [252], Tween-60 [225,229] or Tween-80 [226,230] are employed to enhance the sensitivity. The most employed reagent for determination of Cr(VI) in waters is diphenylcarbazide, employed in more than 70 publications being used as the reference reaction when comparing new procedures for Cr(VI) determination, and monitored at a wavelength of around 540 nm. Other reagents employed for Cr(VI) determination make use of its oxidizing properties. In this sense, reagents such as 2-(5-bromo-2-pyridylazo)-5-diethylaminophenol [253], 2,3,7-trihydroxy-9-dibromohydroxyphenylﬂuorone [254], aniline blue [255], the arsenazo family of compounds [256–263], beryllon III [264], brucine [265], or p-acetylchlorophosphonazo [266], carbazone compounds [267–269], or KI/acridine red/polyvinyl alcohol [270] are oxidized by Cr(VI) wherein the color developed or extent of decoloration is proportional to the chromium concentration. In some cases, Cr(VI) has been determined based on its catalytic effect on some oxidation reactions [271 –287] and surfactants, as cetyltrimethylammonium bromide [271–273], hydroxyethylidene-1,1-diphosphonic acid [274] or cetylpyridinium chloride [275] were added for enhancement of sensitivity.

1132

Sample preparation for chromium speciation

35.5.5.2 Determination of Cr(III) using UV– VIS Determination of Cr(III) has been accomplished using a series of reagents, such as diethylaminophenol [213,253,288 – 294] or dimethylaminophenol compounds [295,296], ﬂuorone compounds [203,297,298], 8-hydroxyquinoline [195,299], Alizarin red S [300], ammonium pyrrolidine dithiocarbamate [200], Arsenazo III [192,198], benzyl-(3-nitrilopropyl)dithiocarbamate [301], bromopyrogallol red/ethyl violet [302], Chrome Azurol S [303,304], Chromotrope 2C [305], cupferron [306], diphenylcarbazide [196,204,206,307], EDTA [209,308], Morine [309], m-sulfo chlorophosphonazo [310], Neutral red/2,20 -bipyridyl [311], Nitrosulfophenol K [312], Nitrosulfophenol M [312], Quercetin [309,313] or Sulfochlorophenol S [312]. As with Cr(VI) determinations, in some procedures, surfactants are employed to improve the sensitivity [203,253,290–292,296,297,303,304,313]. 35.5.5.3 Determination of chromium using spectroﬂuorimetry Fluorimetry has been mainly employed for determination of Cr(VI) in waters, the reagents most frequently used being ﬂuorone compounds [316–319], rhodamine compounds [320,321], crystal violet [322], L -tyrosine [323], pyronine Y [324], Victoria blue 4 R [325], 8-hydroxyquinoline-5-sulphonic acid [326], p-acetamidophenol [327], safranine T [328] and thiamine hydrochloride [329]. Few applications have been found using ﬂuorimetry for determination of Cr(III) [199]. Surfactants such as CTMAB [319], HDTMAB [316] or octylphenyl polyglycol ether (OPPGE) [317] have been employed to enhance the sensitivity. A sequential extraction-ﬂuorimetric method for determining Cr(III) and Cr(VI) using a cryptand 2.2.1 ligand in CH2Cl2 has been described. In this study, Cr(III) was determined by measuring the ﬂuorescence intensity of the organic phase at 549 nm (excitation at 534 nm) and Cr(VI) determined in the aqueous phase after reduction to Cr(III) with ethanol and extraction with CH2Cl2. 35.5.5.4 Chemiluminescence for determination of chromium Determination of chromium in water samples by chemiluminescence has been accomplished mainly for Cr(III) species based on the catalytic oxidation of luminol by H2O2 in basic media [189,190,193,330–337] or with the luminol/ CN2/NaOH system [338]. In some cases, Cr(VI) has been determined by the same method after a previous reduction step [189,190,193,333,334]. On the other hand, chemiluminescence determination of Cr(VI) has been accomplished using ferrocyanide/luminol/NaOH [339,340], rhodamine B/H2O2/NaOH [341], lophine/H2O2/KOH [342] or quercetin/H2O2/KOH [343] reagent mixtures. With this last procedure, Cr(III) has been determined after oxidation to Cr(VI). A chemiluminescence system for separation and determination of both chromium species in environmental waters has been reported by Lu et al. [193]. Water was passed through a 711 anion-exchange column, Cr(VI) was retained

1133

M. de la Guardia and A. Morales-Rubio

and Cr(III) was determined in the eluate after treatment with luminol/H2O2 in an alkaline medium. On the other hand, to determine Cr(VI), a new aliquot of sample was passed through a 732 cation-exchange column, Cr(III) was retained and the eluted Cr(VI) was reduced with acidic H2O2 and determined as above. Other procedures determine one of the species and total concentrations of chromium. In such procedure proposed by Zhang et al. [334], the water sample was divided into two portions, the ﬁrst one was reacted directly with a mixture of luminol/H2O2 at pH 12 for Cr(III) chemiluminescence measurement; the second one was passed through a reduction column of Cu-coated Zn granules and the online generated Cr(III) plus original Cr(III) were measured by chemiluminescence, as above. On the other hand, in a procedure proposed by Han et al. [343], total chromium concentration, after oxidation of Cr(III) by PbO2, was measured by the chemiluminescence of a mixture of quercetin/H2O2/ KOH. From another aliquot of water, Cr(VI) concentration was directly determined using the same reagent mixture. 35.5.5.5 Determination of Cr(III) and Cr(VI) using hyphenated techniques An ion-chromatographic separation of chromium species in water coupled with diphenylcarbohydrazide-based spectrophotometric detection has been proposed by Pobozy et al. [197]. Samples were injected into the pH 3.5 phthalate buffer mobile phase and Cr(III) was not retained on the PRP-X100 anionexchange column, whereas Cr(VI) was. For the post-column detection of Cr(III), the eluate was mixed with Ce(IV) in diluted sulfuric acid for oxidation of Cr(III) to Cr(VI), and then mixed with diphenylcarbohydrazide for its determination at 540 nm. Subsequent elution of Cr(VI) from exchange column and detection at 540 nm provides the concentration of Cr(VI) in the sample. de-Beer and Coetzee [205] proposed the ion chromatographic separation of Cr(III) and Cr(VI) and their spectrophotometric detection in industrial efﬂuents. Water samples, containing EDTA, were injected into a stream of Na2CO3 for ion chromatography on a Dionex 2000i AS5 separator column. Post-column derivatization of Cr(VI) was effected by reaction with diphenylcarbazide and the absorbance of the Cr(III) –EDTA and the Cr(VI)– diphenylcarbazide complexes was measured at 545 nm. Janos [344] reported a procedure for the ion chromatographic determination of Cr(VI) in electroplating waste water on a column of Separon HEMA-S 1000 Q-L using a mobile phase of phosphate buffer at pH 7.5 and detection at 370 nm. Andrle and Broekaert [210] reported the determination of chromium species in plating waste water, accomplishing the separation by reversed phase HPLC on a column of LiChrosorb RP-18 of the previously formed ammonium pyrrolidine-1-carbodithioate complexes of Cr(III) and Cr(VI). Detection was completed at 259 nm. Saleh et al. [126] developed an ion chromatography UV–VIS detection method for Cr(III) hydrolytic polymerization products in natural waters.

1134

Sample preparation for chromium speciation

Hydrolytic monomers, dimers and oligomers present in waters were separated on an IC-PAC-CS5 sulfonate/quaternary amine mixed-resin column and mono-, di- and tri-meric Cr(III) detected by scanning with a photodiode-array detector from 200 to 600 nm. The authors reported that the trimeric Cr(III) peak was predominant in the ion chromatograms of surface water, and accounted for 6.5 –35.9% of total Cr(III). The simultaneous determination of Cr(III) and Cr(VI) in electroplating waste water by reversed phase ion pair HPLC has been carried out by Jen et al. [209]. Water samples were treated with an excess of EDTA to ensure Cr(III) chelation and were then analyzed by HPLC on a Supelcosil LC-8 column on which chromium species were separated and later detected at 242 nm. An online preconcentration of Cr(VI) –diphenylcarbazide complex on a Silasorb C18 precolumn and further elution of the retained analyte onto an HPLC Silasorb C18 analytical column have been described by Padarauskas et al. [345] and applied to drinking, surface and natural water analysis. An integrated separation detection system based on ion chromatography coupled with chemiluminescence detection was developed and applied to the study of chromium speciation in natural waters [189]. An aliquot of sample was injected into an efﬂuent stream containing EDTA and passed through an AS4A anion exchange column in which separation of both chromium species occurred. The column eluate was merged with a stream of sulfur dioxide to reduce Cr(VI) to Cr(III) and then with a stream of luminol/H 2O 2 at pH 11.5 for chemiluminescence measurement. Separation of Cr(III) and Cr(VI) species in fresh water by using dual column has been described by Williams et al. [190]. The system involved the simultaneous injection of a sample into a Dionex CG2 cation-exchange column and a similar Dionex AG4A anion-exchange column in parallel for separation of Cr(III) and Cr(VI), using a post-column reducing agent of K2SO3 to reduce Cr(VI) and a post-column reagent mixture of luminol/H2O2/KOH for chemiluminescence detection. 35.5.5.6 Cr(III) oxidation or Cr(VI) reduction Speciation of Cr using molecular spectrometry sometimes involves the use of redox reactions to change the oxidation state of Cr. In some cases, Cr(III) is oxidized to Cr(VI) by (NH4)2SO4 [320], (NH4)2S2O8 [234,235,238,263,264,346, 347], KMnO4 [215,221,239,307,317], PbO2 [343], KIO4 [202], bromine water in alkaline media [348], Ce(IV) [349– 351], hydroxylamine hydrochloride [211, 291] or PbO2 [352] and total chromium is measured as Cr(VI). In other cases, Cr(VI) is reduced to Cr(III) by sulﬁte [299], H2O2 in HCl media [333], Zn metal [334] or hydroxyl ammonium chloride [212,218,315] and total chromium is measured as Cr(III). On the other hand, following a separation of the chromium species, Cr(III) is measured as Cr(VI) after its oxidation by ammonium persulfate [191], Ce(IV) [197] or KMnO4 [219], or Cr(VI) is determined as Cr(III) after its reduction

1135

M. de la Guardia and A. Morales-Rubio

by ascorbic acid [196], sulfur dioxide [189], H2O2 in acidic media [193] or K2SO3 [190]. 35.6

ANALYTICAL METHODOLOGIES FOR Cr SPECIATION IN BIOLOGICAL FLUIDS

The human body content of Cr is about 6 mg and this decreases with age [353, 354]. Ingestion, especially from foods containing coordination complexes of Cr(III) with nicotinic acid, amino acids, such as glycine, glutamic acid or cysteine and, in general, animal proteins, is the main source of Cr intake [355, 356], followed by inhalation and skin contact, which can be important factors for occupational health problems [357]. When ingested, Cr(III) is absorbed at levels lower than 1% [358] and Cr(VI) on the order of ,2% [359]. The presence of Cr in the body ﬂuids is thus strongly dependent on the chemical form of Cr and on the functional state of the stomach and intestines due to the common reduction of Cr(VI) to Cr(III) by gastric juices [360]. Cell membranes are permeable to Cr(VI) but not to Cr(III) [354] and Cr(VI) is reduced to Cr(III) in the mitochondria and cell nuclei [361]. The highest levels of Cr are found in hair, followed by the lung, liver and kidney [362] and, in general, Cr(VI) accumulates in the reticule endothelial system, including bone marrow, as well in the liver, kidney, spleen and testes [360]. Cr passes the placenta and is secreted in breast milk [363]. Chromium competes with Fe for transferring binding sites [364] and is present in blood, making it a good indicator of recent exposure to soluble Cr(III) compounds [354,360]. Chromium in serum and plasma is always Cr(III) and this parameter is not a good indicator of Cr-tissue storage or metal deﬁciency, except in extreme cases [355]. On the other hand, Cr in urine is the best index of recent exposure to soluble Cr compounds; its level being higher for diabetics than for healthy people [365]. In fact, kidney and liver favor the reduction of Cr(VI) to Cr(III) and the elimination of Cr(III) is relatively fast with half times between 0.5 and 83.4 days [360,362]. Thus, urine and sweat are the main body ﬂuids in which Cr can be determined [354,359]. 35.6.1 Speciation of chromium in biological ﬂuids Total chromium has been determined in the bulk of publications (81%) relating to biological ﬂuids. Moreover, only one of the two species of chromium, Cr(VI) or Cr(III), was determined in the 10% of the cases that take into consideration all detection techniques employed. Determination of Cr(VI) was done in approximately 15% of the studies, whereas Cr(III) was determined in 12%. Furthermore, the two chromium species have been determined together only in the 7% of such studies and

1136

Sample preparation for chromium speciation

ﬁnally, determination of one of the species and total chromium concentrations were done in the 2% of the cases studied. 35.6.2 Types of samples analyzed The most frequently analyzed sample for Cr speciation is urine, studied in the 42% of cases, followed by blood, serum and plasma with 19.1, 18.6 and 5.5% of published papers, respectively. Other body ﬂuids studied were semen (1.5%), synovial (0.5%), cerebrospinal (0.5%) and lung (0.5%) ﬂuids. Milk has been studied in the 9.5% of the cases. As noted earlier, it is important to study the levels of chromium in body ﬂuids because these can indicate a contamination source in working areas and how it can affect the safety and health of workers. Because of this, and since urine, blood and serum are easy to obtain, these are the matrices most frequently analyzed (79.4%). The number of studies in which speciation has been conducted is considerably lower than those involving total Cr determination, probably due to matrix complexity. In fact, urine can be considered the simplest of biological ﬂuid matrices and, because of that, 67% of chromium speciation studies have been focused on it, followed by matrices such as blood, serum or plasma with 12.1, 12.1 and 9.1%, respectively, of the chromium speciation studies. 35.6.3 Pretreatments and techniques applied The most frequent pretreatments employed for Cr(VI) and/or Cr(III) determination in biological ﬂuids are complexation (41%), separation by column/ionic exchange (24%) and preconcentration on different supports (21%). Seventeen percent of the pretreatments are based on separation of Cr(III) and Cr(VI) species and further independent determination of them. On the other hand, pretreatments such as acid digestion, liquid extraction and precipitation have been employed in 10% of the cases. These processes are frequently used after complex formation and extraction prior to Cr determination. In some cases, different pretreatments are sequentially employed in the same publications. Atomic spectrometric techniques are the most often employed for Cr speciation in biological ﬂuids (71.9%), GF-AAS being used 40.6% of the time, F-AAS (6.3%), ICP-OES (6.3%), XRF (3.1%) and those hyphenated with chromatography (15.6%). This group also includes ICP-MS hyphenated with chromatography (9.4%). Molecular spectrophotometry has been employed in 21.9% of studies, as UV–VIS, hyphenated with chromatography, the most extensive strategy employed (9.4%), followed by chemiluminescence (6.2%) and UV–VIS spectrophotometry (6.2%).

1137

M. de la Guardia and A. Morales-Rubio

Other techniques, such as voltammetry (6.2%) or chromatography (3.1%), have also often been employed to determine Cr(VI) and Cr(III). 35.6.4 Chromium speciation using atomic spectrometry detection Table 35.3 summarizes samples, techniques and analytical ﬁgures of merit dealt with in the studies on the determination of Cr(III) and Cr(VI) in biological ﬂuids. 35.6.4.1 GF-AAS for determination of Cr species The determination of chromium species in urine by GF-AAS has been accomplished by Qi et al. following chelate coprecipitation [46] and by Shan et al. after solvent extraction with a high-molecular-weight secondary alkylamine [376]. In the last case, urine was digested with concentrated HNO3 and H2SO4 and the solution was evaporated almost to dryness. The residue was dissolved in water, pH adjusted to 2.8 and Cr(VI) extracted with a solution of the amine N-235 in benzene. Portions of the extract were analyzed by GF-AAS. The amount of Cr originally present as Cr(VI) can be determined by direct extraction. An original methodology for “in situ” chromium speciation, based on the different GF-AAS atomization properties of both species, has been proposed [51,379]. In initial studies, the volatile Cr(III) complex was not completely volatilized at low temperature, and similarly not all the Cr(VI) was retained for determination at high temperatures [379]. In the procedure employed by Arpadyan and Krivan [51], urine was diluted with a solution containing 1,1,1triﬂuoropentane-2,4-dione and sodium acetate. A 20 ml portion of the resulting solution was injected into the graphite tube and the furnace was heated to 4008C for 5 min for volatilization of Cr(III); the temperature was then raised over 10 s to 12008C during 40 s and then to 27008C for volatization and measurement of Cr(VI). Long and Han [377] employed co-precipitation with aluminum hydroxide for determination of Cr(III) and Cr(VI) in urine by GF-AAS. The Cr(III) was coprecipitated directly with Al(OH)3, whereas Cr(VI) could be co-precipitated only after reduction to Cr(III) with ascorbic acid. Precipitates were aged for 30 min before collection and organic material removed by digestion with HNO3 –H2O2 before determination by atomic absorption. Determination of total chromium and Cr(VI) in UHT milk by GF-AAS has been reported by Lameiras et al. [367]. For total Cr determination, milk was mixed with Triton X-100 and diluted with H2O. For determination of Cr(VI), proteins were separated by precipitation with acetate buffer at pH 3.5, the supernatant solution was passed through a column of Chromabond NH2 and Cr(VI) was retained. After elution of Cr(VI) with 2 M HNO3, both total Cr and Cr(VI) were quantitated by GF-AAS.

1138

TABLE 35.3 Figures of merit for the determination of Cr(III) and/or Cr(VI) in biological ﬂuids using atomic spectrometric techniques Treatment

Species

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

Blood

PC

(III) (VI)

F-AAS

–

–

–

1996 [20]

Milk

CPX PC SPE PC PTT SPE EXT CPX – PC CPX PC OXI CPX PC

(III)

AAS

24 (III) 75 (VI) –

–

–

–

2000 [366]

(VI) Cr

GF-OAS

0.15

,50

,6

.93

1998 [367]

(III) (III) (VI) (VI) (III) (VI) (VI) Cr

AAS HPLC-UV-AAS GF-AAS IC-DCP-AES F-AAS

– – 0.2 1 0.6

– – ,1000 ,100,000 ,100

– – 0.8– 4 – 2

– – 97–102 – 96–107

2000 [368] 1987 [369] 1994 [370] 1989 [93] 2001 [151]

(III)

AAS

–

–

–

–

2000 [371]

Milk

Plasma Plasma Serum Serum Urine

Urine

continued

Sample preparation for chromium speciation

Matrix

1139

1140

TABLE 35.3 (continuation) Matrix

Urine

Urine Urine Urine

Urine

SPE CPX PC RED CPX PC SPT PC OXI CPX PC SPT PC SPE

Species

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

(III) (VI)

F-AAS

24 (III) 75 (VI)

–

3.8

–

1996 [20]

(III) (VI)

HPLC-ICP-MS

3 pg

,1000

0.9 (III) 1.3 (VI)

–

1995 [145]

(III) (VI)

IC-ICP-AES

,1000

0.3– 1.6

–

1995 [372]

(VI) (III) (VI)

ASV-ICP-MS IC-ICP-MS

12–15 (III) 35–47 (VI) – 0.3– 0.8 (III) 0.4– 1 (VI)

– ,1000

– 1.3 (III) 2.4 (VI)

62 –

1994 [373] 1994 [374]

(VI)

GF-OAS

–

–

–

1992 [375]

–

M. de la Guardia and A. Morales-Rubio

Urine

Treatment

TABLE 35.3 (continuation) Treatment

Species

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

Urine

CPX PTT EXT CPX PC SPT PTT RED PC CPX SPT FTT EXT CTF PC – PTT

(III) (VI)

GF-AAS

–

–

GF-AAS GF-AAS

0.01 –

0.1 –5 –

.94 (III) .87 (VI) – 97–104

1991 [46]

(VI) (III) (VI)

5.8 (III) 2 (VI) – –

1990 [376] 1986 [51]

(III) Cr

GF-AAS

–

,60

5

–

1986 [377]

(III) (III) (VI)

ICP-OES GF-AAS

0.92 –

,1000 –

12 –

– –

1986 [378] 1985 [379]

(III) (VI)

XRF GF-AAS

3 0.1

,60 –

18 –

74 95–100

1983 [380] 1983 [381]

(III) (VI) (VI) (III) (VI)

ICP-OES GF-AAS GF-AAS

0.028 – –

– – –

21 – –

– – –

1983 [148] 1981 [382] 1981 [383]

Urine Urine

Urine Urine Urine Urine Urine Urine Urine Urine

Sample preparation for chromium speciation

Matrix

1141

M. de la Guardia and A. Morales-Rubio

35.6.4.2 F-AAS for determination of Cr species An interesting study was reported by Paleologos et al. [151] for the speciation of chromium in urine based on surfactant micelle methodology. The method involves the preconcentration of the ammonium pyrrolidinedithiocarbamate metal chelates at the micelle cloud point. The condensed surfactant phase, along with the metal chelate(s), is introduced into the F-AAS in which discrimination of Cr species is feasible by calculating the Cr(III) concentration from the difference between total Cr and Cr(VI).

35.6.4.3 ICP-AES for determination of Cr species Cox and McLeod determined Cr(III) in urine by FI-ICP-AES after preconcentration of these species on an activated alumina microcolumn. The Cr(III) was adsorbed from the sample onto the column using 0.02 M NH3 as carrier and was then eluted with 2 M HNO3 [378].

35.6.4.4 Hyphenated techniques for determination of Cr species Gaspar et al. [20] used an online chromatographic separation of Cr(III) and Cr(VI) for their determination in blood and urine. Digested samples were injected into a water carrier and passed to a column of Eurospher 100-C18. The Cr(III) was retained and concentrated and the unretained Cr(VI) was determined by F-AAS. Determination of Cr(III) was achieved after elution with H2O/methanol (1:4) and measured by F-AAS. IC-DCP-OES has been employed for direct determination of Cr(III) and Cr(VI) in serum [93]. Sample solutions were treated by ion chromatography on an HPIC-AS-7 strongly basic anion-exchange resin or HPIC-CS-2 strongly acidic cation-exchange resin columns. The mobile phases employed were HNO3 or trilithium citrate/H2C2O4 for the anion or cation exchange columns, respectively. Moreover, on-column sample preconcentration was achieved by multiple injections into the column followed by rapid regeneration of the column by elution with HCl for Cr(III) (both columns) or with NaOH for Cr(VI) (anion-exchange column). Hyphenation of HPLC-ICP-MS, together with a direct injection nebulizer, has been employed by Zoorob et al. for chromium speciation in a freeze-dried urine standard reference material [145]. Tomlinson et al. [374] employed IC-ICP-MS hyphenation for speciation of chromium in urine. The Cr(III) was complexed with 2,6-pyridine dicarboxylic acid prior to separation of both chromium species on an HPLCCS5 column. IC-ICP-OES hyphenation was employed for chromium speciation in urine [372]. Samples were injected into a mobile phase of HNO3 and the separation of Cr(III) and Cr(IV) was achieved on a column of latex anion-exchanger OmniPac

1142

Sample preparation for chromium speciation

PAX100 Guard, the Cr(III) being eluted in the dead volume of the column and Cr(VI) 90 s later. 35.6.5 Chromium speciation using molecular spectrophotometry detection Table 35.4 summarizes samples, techniques and analytical ﬁgures of merit dealing with studies published on chromium speciation in biological ﬂuids using molecular spectrophotometry. As can be seen, complexation is the most common treatment. UV–VIS detection can provide the highest linear range and is the most widely recommended technique for samples having high chromium concentration. On the other hand, chemiluminescence can be adequate for detection of low levels of chromium species in samples. RSDs and recovery values are of the same order for all molecular spectrophotometric techniques employed for Cr speciation in biological ﬂuids, even if they are hyphenated with chromatography. Cr(VI) in urine was determined by colorimetric reaction with 1,5diphenylcarbazide. When the reagent was added to the urine, an immediate red-violet color appeared, having a characteristic absorption peak at 544 nm [387]. Gao employed chemiluminescence for the determination of trace amounts of chromium in urine [388]. Urine was evaporated to dryness and the residue was treated with 30 ml H2O and 2 ml 60 g l21 sulfurous acid. The solution was boiled and concentrated and KBr and EDTA were added. The mixture was diluted with H2SO4 of pH 2.5 and portions of the solution were mixed with luminol and H2O2 and the generated chemiluminescence measured. Escobar et al. [384] determined Cr(III) in urine and blood serum by means of ﬂow-injection chemiluminescence analysis. Volumes of 1 ml urine or blood were digested with 3 ml HNO3 in a microwave oven for 5 min. After the addition of 2 ml H2O2, the reaction mixture was evaporated to dryness. The digestion was then completed with 1 ml H2SO4 and 2 ml H2O. The transparent residue was dissolved in water and diluted to 10 ml following the addition of sufﬁcient EDTA and H3PO4 to give a pH of 4. Finally, the detection method was based on the Cr(III)-catalyzed chemiluminescent reaction between luminol and H2O2. An anion-exchange HPLC determination of Cr(VI) in lung preparations after “in vivo” and “in vitro” treatment with sodium chromate has been proposed [389]. Lung tissue was homogenized in Tris –HCl buffer (pH 7.4) and centrifuged at 38,000g for 10 min. The precipitate was homogenized again and centrifuged as above. The supernatant solution was analyzed by HPLC on a column of Mono Q, HR 5/5 anion-exchange resin, using gradient elution with Tris –HCl and detection of Cr(VI) at 370 nm. Suzuki employed anion-exchange HPLC for determination of water-soluble Cr(III) and Cr(VI) complexes in biological materials [369,386]. Rat plasma,

1143

1144 TABLE 35.4

Matrix

Treatment

Species

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

Blood Blood Culture Urine Urine Urine Serum Serum Lung Plasma

CPX – CPX CPX CPX CPX CPX – CTF CPX

(III) (VI) (III) (VI) (VI) (III) (III) (III) (VI) (VI) (III) (VI)

Chemiluminescence UV– VIS HPLC-UV– VIS UV– VIS Chemiluminescence Chemiluminescence Chemiluminescence UV– VIS HPLC-UV– VIS HPLC-UV– VIS

– – 7 ng 370 – – – – 0.5 ng –

,6 – ,7000 ,20,000 – ,6 ,6 – ,8000 –

0.9– 2 – ,3 – – 0.9– 2 0.9– 2 – ,2.1 –

98–140 – 97 ^ 2 – 97–103 89–100 98–104 – 91–96 –

1998 [384] 1979 [385] 1986 [386] 2000 [387] 1999 [388] 1998 [384] 1998 [384] 1979 [385] 1989 [389] 1987 [369]

M. de la Guardia and A. Morales-Rubio

Figures of merit for determination of Cr(III) and/or Cr(VI) in biological ﬂuids using molecular techniques

Sample preparation for chromium speciation

erythrocyte lysate and liver supernatant solutions were treated in vitro with Cr(VI) or Cr(III). Samples were then analyzed by HPLC with use of a fastprotein liquid chromatograph equipped with a Mono Q HR 5/5 anion-exchange column, using gradient elution. Detection of Cr(VI) was by continuous monitoring at 370 nm and AAS, and other Cr species were monitored by AAS with all instruments connected in series to the column outlet. 35.7

ANALYTICAL METHODOLOGIES FOR SPECIATION OF Cr IN SOLID SAMPLES

35.7.1 Speciation of chromium in solid samples The low solubility of Cr(OH)3 at neutral pH and the association of Cr in chromate-based minerals, in addition to its accumulation in kidney and liver, favor the presence of chromium species in solid samples, from geological materials to foods, technological products and tissues. Total chromium has been determined in the bulk of publications (87.0%). Moreover, only one of the two most common species of chromium, Cr(VI) or Cr(III), was determined in 10.5% of all detection techniques employed. Determination of Cr(VI) was done in approximately 12.2% of the studies, whereas Cr(III) was determined in 6.4%. Furthermore, the two chromium species have been determined together only in the 2.9% of such studies and, ﬁnally, determination of one of the species and total chromium concentrations were done in the 2.1% of the cases. 35.7.2 Types of samples analyzed It is well known that when performing analysis by means of instrumental techniques, usually the most difﬁcult and laborious part of the work is sample pretreatment, being more difﬁcult and time consuming when the sample is more complex. Sometimes the matrix plays a decisive role in the analytical procedure, because its components can vary the oxidation state of the analyte during treatment of the sample with acids or solvents. This circumstance has a signiﬁcant importance when analyzing solid samples, given the drastic conditions often needed for their dissolution or for the extraction of the analyte. Studies reviewed have been divided in four groups, depending on the origin of the sample analyzed, in order to evaluate the different speciation strategies developed: (i) industrial materials, (ii) soils, sediments, sludges and geological samples, (iii) clinical and biological samples and (iv) meal and food samples. 35.7.2.1 Determination of Cr species in industrial materials References concerning determination of chromium in industrial samples report the analysis of aerosols, airborne particulate, alloys, aluminum ashes, building materials, cement, coal, dust, ﬁbers, industrial wastes, iron, leather, metals,

1145

M. de la Guardia and A. Morales-Rubio

pharmaceutical products, steel, welding dust and welding fumes [390–463]. However, in the majority of studies, no true speciation was done and the fact that Cr(III) or Cr(VI) was measured depends on the sample treatment applied before detection more than on the actual presence of these oxidation states in the original sample. The analysis of aerosol samples is quite interesting since aerosol particles are introduced into the lungs during breathing. Thus, accurate determination of the Cr(VI) content on/in these particles has a special importance when monitoring metal processing factories. Furthermore, Cr(VI) is the most carcinogenic inorganic constituent in aerosols [7]. In addition, the high oxidation potential of Cr(VI) can play an important role in atmospheric chemistry if it is present in/on aerosol particles [390]. Sample pretreatment generally affects the speciation of analytes. To determine certain species, the procedure mostly employed is complexation before determination by UV–VIS spectrophotometry. Other procedures isolate the species to be determined by extraction or by using column separation processes. Complexing agents used for Cr(III) involve EDTA, which has been the most frequently utilized when analyzing aerosols [392], while phenanthraquinone monoxime has been applied to studies of alloys [406]. Naphthalene was used for aluminum samples as well as with steel samples [408]. trans-1,2-Diaminecyclohexane-N,N0 ,N0 ,N-tetra acetic acid was employed with building materials and leather [416] to obtain an anionic form of Cr(III). Zephiramine was also applied when analyzing steels and iron [294] and Cr(III) has been determined in steel samples using Chrome azurol [304] or 4-(2-pyridylazo)resorcinol and xylometazoline hydrochloride [451] as complexing agents. For complexing Cr(VI), diphenylcarbazide is the most widely used agent for cements [419,464], dust discharge deposit [424], ash [216], timbers [452,453] and welding fume analysis [458] followed by determination of this complex using UV–VIS spectrophotometry. In addition, N1-hydroxy-N1,N2-diphenyl benzamidine and N-aryl acetamide were used on cement, coal dust and ash samples [236]. Disodium 1-(4-hydroxysalicylideneamino)-8-hydroxy naphthalene-3,6-disulfonate has been used on ash samples to measure the ﬂuorescence [413]. The Cr(VI) in metals has been complexed with salicylﬂuorone, with o-nitrophenylﬂuorone or with o-chlorophenylﬂuorone [431]. A variety of agents were reported for use with steel sample analysis, such as phenylﬂuorone [446], 1,8-dihydroxy-2-(40 -chloro-20 -phosphonophenylazo)-7-(600 ,800 -disulfonaphthyl azo)-3,6-disulfonaphthalene [447], dibromohydroxy phenylﬂuorone [448], 2(5-bromo-2-pyridylazo)-5-diethylaminophenol [450] and diantipyrylstyryl methane [232]. Another interesting aspect of selective extraction of chromium species is the separation or isolation procedures used. Prior to separation, an acid treatment of the sample is commonly applied [390]. Separation procedures reported basically include anion exchange, extraction and ﬁltration. Anion exchange is frequently used to separate the species to be determined directly or after a

1146

Sample preparation for chromium speciation

previous extraction or complexation. Thus, anion-exchange chromatography was used on aerosol samples for determination of Cr(III) [392], Cr(III) and Cr(VI) [390], while anion-exchange resins have been applied for the analysis of Cr(III) in air [394] and in the analysis of airborne particulates for Cr(III) and Cr(VI) [F399] and Cr(VI) [F400]. Welding dusts were also subjected to a similar anion exchange using amberlite resin to determine Cr(VI) [119,457]. Extraction processes were employed mainly for determination of Cr(VI). In spite of this, Cr(III) was extracted from a spent catalyst with bis(2,4, 4-trimethylpenthyl) dithiophosphonic acid before being determined by ICP-AES [417]. In some cases, samples must be ﬁltered before extraction, such as for determination of Cr(VI) in dust [424]. Neotetrazolium chloride was used to extract Cr(VI) from steel samples [443]. Other substances have also been used for extracting Cr(VI) from steel samples, such as rhodamine 6G [219] and benzyltributylammonium [449]. Liquid –liquid extraction was also applied to the determination of Cr(III) and Cr(VI) in welding fumes, using hydrochloric acid, IBMK and amberlite [459] and in pharmaceutical products, using the formation of chromyl chloride prior to its extraction into IBMK [432]. For sampling of particulate air components, the ﬁlter sampling technique was applied to the determination of both species, [399,401] or only Cr(VI) [400]. It has also been applied to welding fume samples [463]. In general, ﬁltration is only the ﬁrst treatment step, followed by extraction for determination of Cr(VI) in dust and melting [424] or in welding fumes [460]. The coupling of separation and measurement techniques was also used to determine Cr(III) and Cr(VI) in ashes using HPLC-F-AAS and HPLC-ICP-MS detection [414] and in dyes by LC-AAS and LC-ICP-OES [425]. 35.7.2.2 Determination of Cr species in soils, sediments, sludges and geological samples Table 35.5 summarizes the samples, techniques and analytical ﬁgures of merit dealing with studies published on speciation of chromium in geological materials. As can be seen, the most commonly used procedure for chromium speciation in this group of samples is the extraction of species of interest by heating the sample with suitable organic or inorganic solutions or solvents. Depending on the nature and complexity of the matrix, a previous digestion of the sample could be necessary. In addition, the extraction sometimes involves the complexation of the analyte species. Several solvents or solutions were compared in order to extract the same species; thus, Cr(VI) was extracted from soil samples with acidic or alkaline solutions [492], nitric acid, acetate buffer, KCl or IBMK [495] or NaOH solution [503]; from soils and sediments with phosphate buffer or alkaline media with or without sonication [473] and from soils, loam, ores, sand and sediments with K2HPO4/KH2PO4, CO3 22 /NaOH or NaOH solutions [467]. Total chromium was

1147

1148

TABLE 35.5 Chromium speciation in soils, sediments, sludges and geological samples Treatment

Species

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

Geological

EXT OXI FUS CPX FUS CPX EXT CTF CPX FTT – – EXT CTF CPX PC CPX CPX FTT EXT CTF CPX –

(VI) Cr (VI)

UV– VIS

–

–

2.2

–

1996 [219]

HPLC-UV

–

20– 200

–

–

1995 [465]

(VI) Cr (VI)

UV– VIS

3.2

,60

0.3–1.2

–

1994 [466]

UV– VIS

–

–

–

–

1995 [467]

(VI) (VI) (VI) (VI)

UV– VIS; AAS UV– VIS Titrimetry UV– VIS

– 3 – –

– ,400 ,52,000 –

– ,1.4 0.9 –

– 97– 102 99– 101 –

1985 [424] 2000 [468] 1999 [469] 1995 [467]

(III) (VI) (VI) (VI) (VI)

ICP-AES Polarography UV– VIS UV– VIS; AAS UV– VIS

– – – – –

– 50– 3200 – – –

1.4 – – – –

– – – – –

1994 [470] 1990 [471] 1988 [450] 1985 [424] 1995 [467]

(VI)

UV– VIS

–

,1200

0.4–0.7

–

2001 [255]

Geological Geological Loam

Melted Ores Ores Ores

Ores Ores Ores Ores Sand

Sediment

M. de la Guardia and A. Morales-Rubio

Matrix

TABLE 35.5 (continuation) Treatment

Species

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

Sediment Sediment

– EXT FTT CTF EXT EXT CTF CPX PC EXT CPX EXT EXT CTF CPX – – FTT CPX – – –

(III) (VI)

F-AAS UV– VIS

23–305 ng g21 –

– –

,9 –

.95 –

2001 [472] 1997 [473]

(III) (VI)

UV– VIS UV– VIS

– –

100 –2500 –

– –

– 97–100

1996 [474] 1995 [467]

(VI)

UV– VIS

–

–

–

–

1989 [475]

(VI) (VI)

UV– VIS UV– VIS

– –

10– 2100 –

– –

– 97–100

2001 [476] 1995 [467]

(VI) (III) (III) (VI) (VI) (VI) (III) (VI)

GF-AAS ICP-AES UV– VIS

20 ng g21 – –

– – –

6.5 – –

93–101 – –

2000 [477] 1988 [478] 1984 [479]

UV– VIS XRA CZE-UV

– – 100 150

,1200 – ,10,000

0.4– 0.7 6.1 –

– – –

2001 [255] 2001 [480] 2001 [481]

Sediment Sediment

Sediment

Sludge (sewage) Sludge

Sludge (sewage) Sludge (sewage) Sludge Soil Soil Soil

1149

continued

Sample preparation for chromium speciation

Matrix

1150

TABLE 35.5 (continuation) Treatment

Species

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

Soil

–

–

–

–

2001 [482]

EXT – – EXT – EXT – EXT EXT

UV– VIS ICP-MS AAS UV– VIS UV– VIS UV– VIS Voltammetry AAS Voltammetry UV– VIS F-AAS

–

Soil Soil Soil Soil Soil Soil Soil Soil Soil

2001 [483] 2001 [434] 2001 [484] 2001 [476] 2000 [485] 2000 [486] 2000 [487] 1999 [488] 1998 [86]

– 10– 400 – –

– 1.1 – – – – 5.2 – 7.2 5.1 – 1.8 – –

– – – – – – – – –

– – – EXT FTT CTF EXT MW

– – – – – – 0.003 – 0.2 2.5 – – – –

– 100 –1800 – 10– 2100 – – 0.05–5.2 – –

Soil Soil Soil Soil

(VI) Cr (VI) (VI) (VI) (VI) (VI) (VI) (VI) (VI) (III) (VI) (VI) (VI) (VI) Cr (VI)

– – – –

1998 [489] 1998 [439] 1998 [490] 1997 [473]

(VI) (III) (VI) Cr (VI) Cr (VI)

UV– VIS AAS Polarography

– –

– –

– 98.6

1997 [443] 1996 [491]

UV– VIS

– 1.5 0.8 – 20

,5500

–

–

1996 [492]

Polarography

0.1

0.2 –200

–

97– 100

1996 [493]

Soil Soil

Soil Soil

EXT CPX FUS

UV– VIS UV– VIS Voltammetry UV– VIS

M. de la Guardia and A. Morales-Rubio

Matrix

TABLE 35.5 (continuation) Treatment

Soil

EXT

Soil

EXT CTF CPX CPX EXT EXT

Soil Soil Soil

Soil

CPX EXT EXT CPX PC OXI CTF FTT EXT CPX OXI

Soil

EXT

Soil Soil Soil

Species

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

Cr (III) (VI) (VI)

F-AAS GF-AAS UV– VIS

–

–

–

–

1995 [494]

–

–

–

97– 100

1995 [467]

(VI) Cr (III) (VI) (III) (VI) (VI) Cr (VI) Cr (VI) Cr

UV– VIS ICP-OES LC-ICP-OES

–

–

–

–

1995 [495]

4 ng g21 0.5 ng g21 250 270 –

–

–

,96

1995 [496]

,40,000

–

–

1994 [497]

–

–

–

1994 [498]

0.3

0.5– 500

–

96.1

1993 [499]

0.3

200– 400

4.5

–

1992 [500]

(VI) Cr (III)

LC

0.1

–

2.8

109

1991 [501]

GF-AAS

0.12

–

10

–

1991 [502]

HPLC-ICP-OES UV– VIS AAS Voltammetry Polarography UV– VIS GF-AAS HPLC-GF-AAS

1151

continued

Sample preparation for chromium speciation

Matrix

1152

TABLE 35.5 (continuation) Treatment

Species

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

Soil

(VI)

GF-AAS

–

–

–

98

1990 [503]

–

–

–

–

1990 [298]

UV– VIS ICP-OES HPLC-DCP-OES

– – –

– – –

– – –

– – –

1989 [504] 1988 [478] 1986 [505]

Soil

CPX

(VI) Cr (VI) (III) (III) (VI) (III)

UV– VIS

Soil Soil Soil

EXT CTF FTT RED CPX CPX – CPX

UV– VIS

–

,500

–

–

1984 [294]

Soil

M. de la Guardia and A. Morales-Rubio

Matrix

Sample preparation for chromium speciation

also extracted from soil samples by comparing different extracting agents [495]. From these studies, it seems difﬁcult to choose the most appropriate agent for total recovery of Cr(VI) [495]. In other studies, a speciﬁc extractant could be used but alkaline extractions are strongly recommended, such as Na2CO3 with [473] or without [467] the addition of NaOH, as well as a combination of NaOH and NH4NO3 [503]. Other authors proposed the use of a single extraction agent for soil analysis [491,493,496,498,500,502]. Determination of total chromium and Cr(VI) was reported using NaHCO3 [493] or a combination of Na2CO3 and NaOH [498]. Extractions with KH2PO4 [500] and with CaCl2 [496] were proposed for determination of Cr(III) and Cr(VI). In addition, NaOH was also used for quantitation of total chromium, Cr(III) and Cr(VI) [491], Cr(III) being selectively extracted using pelargonic acid [502]. With respect to digestion procedures, samples were not always subjected to the same hard conditions. Alkaline fusion with Na2O2 at 7008C can be applied to geological samples for determination of Cr(VI) through the preparation of a ternary complex [465], whereas treatment with citric acid solution (pH 5) was strong enough to determine Cr(VI) in sediments where the corresponding diphenylcarbazide complex was used [475]. As described for the use of complexing agents in industrial materials, EDTA was proposed for determination of Cr(III) and Cr(VI) in soils [494] and in slags and sludges [479], after performing an appropriate extraction procedure. Acetic acid was proposed for the determination of both species [494]. In addition, diphenylcarbazide can be used for complexing Cr(VI) in sediments [475] as well as in soils [298] and soil leachates [504]. These procedures have also been reported to enable the quantiﬁcation of total chromium or Cr(III) by difference between total chromium and Cr(VI). 35.7.2.3 Determination of Cr species in solid clinical and biological samples Table 35.6 summarizes the literature on this topic, the most commonly analyzed samples being feces and hair. Garcia-Rico et al. [506] determined Cr(III) in feces, wherein they were dried for 12 h at 1008C and ground so as to pass through a 0.5 mm screen. Samples were weighed into a crucible, heated on a hot plate for 10 min and ashed at 6508C for 4 h. The ashes were dissolved in 10 ml 70% HNO3 and digested in a microwave oven (7 min at 570 W, 10 min at 540 W and 20 min at 510 W). A 6 ml aliquot of 30% H2O2 was added followed by further digestion (1 min at 570 W and 7 min at 540 W). After digestion, the sample was cooled and diluted to 100 ml and quantitated by F-AAS. Aguilera et al. [507] determined Cr(III) in feces as Cr2O3. The sample was dry-ashed at 450–5508C in a nickel crucible, fused with Na2CO3 – K2CO3 – KNO3 (5:5:2), the cooled melt dissolved in H2O and the absorbance of the solution measured at 372 nm.

1153

1154 TABLE 35.6

Matrix

Treatment

Species

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

Feces Feces Hair Hair Hair Hair

MW FUS CPX CPX CPX PrCP RED CPX CPX EXT – –

(III) (III) (VI) (VI) (III) (III) Cr (VI) (III) (VI) (VI) (VI)

F-AAS UV– VIS UV– VIS UV– VIS Chemiluminescence F-AAS

– – 0.04 6.6 – 0.8

,4000 ,30,000 ,80 ,46 0.02– 6 ,150

4.5 0.7 2.8 –3.2 2.7 –3.3 0.9 –2.0 2.7

91.3 99.3 102 –103 98–99 109 –115 –

1999 [506] 1988 [507] 2001 [256] 2000 [257] 1998 [384] 1996 [508]

Fluorimetry RP-HPLC-UV UV– VIS PIXE PIXE

2 2 2 ng – –

2 –60 ,400 – – –

,4.7 0.4 –3.9 – – –

– – – – –

1994 [317] 1994 [305] 1997 [443] 1999 [509] 1997 [510]

Hair Hair Plants Tissue (renal) Tissue (renal)

M. de la Guardia and A. Morales-Rubio

Chromium speciation in solid clinical and biological samples

Sample preparation for chromium speciation

A representative example of the analytical procedures for speciﬁc Cr(III) quantitation in solid biological samples is the methodology proposed for hair samples that involves dry ashing for 3 h at 7508C followed by HPLC with UV– VIS detection [305]. Escobar et al. [384] determined Cr(III) in hair using ﬂow injection and chemiluminescence Cr(III)-catalyzed reaction between luminol and H2O2. Hair samples were washed with n-hexane, ethanol and water and dried at 908C. Portions of 0.4 –0.5 g were digested with 5 ml HNO3 in a microwave oven for 5 min. After the addition of 2 ml H2O2, the reaction mixture was evaporated to dryness. The digestion was then completed with 1 ml H2SO4 and 2 ml H2O. The transparent residue was dissolved in water and diluted to 10 ml following the addition of necessary reagents to produce the chemiluminescence. Zou et al. [508] determined Cr(III) and total chromium in hair by means of online coprecipitation and F-AAS detection. Human hair was digested with HNO3/HCl/H2O2 and the digest was treated with 5% phenanthroline, 5% sulfosalicylic acid and 1% hydroxylamine hydrochloride (I), adjusted to pH 3 and mixed with 0.5% lanthanum nitrate. To determine Cr(III), reduction with I was omitted. Prepared samples were merged with a stream of NH4OH/NH4Cl buffer of pH 9.4. Coprecipitation of Cr(III) with lanthanum hydroxide occurred in a knotted reactor. The precipitate was eluted from the walls of the reactor with a stream of HCl and carried to the FAS for detection of Cr. 35.7.2.4 Determination of Cr species in foods Table 35.7 summarizes the samples, techniques and analytical ﬁgures of merit dealing with studies published on speciation of chromium in solid food materials. Determination of Cr(III) in a variety of foods, through the formation of different complexes, has been reported [288,511,512]. As a previous step for Cr(III) determination, the dry ashing of ﬂour at 7508C for 3 h was proposed [305]. Fishbone meal samples could be treated with a NaOH solution at pH ¼ 10 [514]. In addition, determination of total chromium requires a previous mineralization of samples using acid mixtures. Thus, HNO3, HCl and HF were proposed for animal feed samples [513], while HNO3 and HClO4 were recommended for ﬁshbone meal samples [514]. Finally, Cr(VI) can be determined from the difference between total chromium and Cr(III) [514] or by a speciﬁc extraction procedure with NaOH, proposed for the analysis of animal feeds [513], and catalytic polarography, used when analyzing unpolished rice [519]. 35.7.3 Solid sample treatments for speciation of chromium Dissolution of samples or quantitative leaching of chromium species is necessary in most cases before the measurement of Cr species by appropriate techniques. These pretreatment steps, in which acids, bases or other reagents

1155

1156

TABLE 35.7 Chromium speciation in food samples Treatment

Specie

Technique

LOD (ng ml21)

Range (ng ml21)

RSD (%)

R (%)

Year [Ref.]

Cereals Chinese peach Feeds (animal)

(III) (III) (VI) Cr (III) (III) (III) Cr (III) (VI) (VI) (III) (III) (III) (III)

UV– VIS Voltammetry GF-AAS

,800 0.2–100 0.4–50 0.5–5 ,400 ,800 –

– – 3 –7 2 –4 0.4 –3.9 – 3 –4

90–104 – –

1993 [511] 1991 [512] 1994 [513]

RP-HPLC-UV UV– VIS F-AAS

– 0.1 0.4 0.5 2 – –

– 90–104 100–106

1994 [305] 1993 [511] 1994 [514]

UV– VIS GF-AAS UV– VIS UV– VIS Potentiometry RP-HPLC-UV HPLC-UV

– 1.8 – – – 2 2

,800 1.8–50 ,1600 8– 600 0.03– 0.3 ,400 4– 400

– 4.1 –6.5 3.7 –4.4 3.6 – 0.4 –3.9 –

90–104 .93 96–98 95–103 97–100 – –

1993 [511] 2000 [515] 1999 [516] 1996 [292] 1995 [517] 1994 [305] 1992 [288]

(VI) (VI) (III) (VI)

Polarography Polarography UV– VIS

200 – –

,5000 50–3200 –

– – –

– – 95–108

1990 [518] 1990 [519] 1995 [198]

Tuna

CPX CPX EXT CTF CPX CPX PrCP FTT CPX SPT CPX CPX PC CPX CPX CTF CPX CPX SPT CPX Cr PC

F-AAS

–

–

1996 [20]

CPX

24 75 –

–

Vegetables

(III) (VI) (III)

,800

–

90–104

1993 [511]

Flour Liver (pig) Meal (ﬁshbone) Meat Powered milk Powered milk Pumpkin Rice Rice Rice Rice Rice Tea soup

UV– VIS

M. de la Guardia and A. Morales-Rubio

Matrix

Sample preparation for chromium speciation

are employed, must be carried out carefully to assure that no change in the original oxidation state of chromium occurs during the pretreatment. On the other hand, some pretreatments employed for leaching chromium from samples cannot achieve total extraction, and then the speciation is accomplished only from the leached chromium, not from the total content of the element in the sample. The most frequent pretreatments employed for Cr(VI) and/or Cr(III) in solid samples are complexation (36%), liquid extraction (30%) and ﬁltration (17%). Other treatments, such as preconcentration, centrifugation and separation, have been employed in 15% of the cases, and reduction, oxidation, acid digestion, microwave assisted or fusion have been employed as well in 15% of the cases. It must be noted that some pretreatments were employed sequentially in the same work. Molecular techniques are the most frequently employed for speciation studies in solid samples (56.3%), UV–VIS being used most frequently (48.5%), followed by ﬂuorimetry/chemiluminescence (3.9%) and those hyphenated with chromatography (3.9%). Atomic spectrometry has been employed in 31% of the studies, these being GF-AAS, the most extensive technique employed (12.1%), followed by XRF (6.3%), F-AAS (5.8%), ICP-OES (3.4%) and hyphenated techniques with chromatography (3.4%). Finally, other techniques such as voltammetry and polarography (6.3%) and those related to mass spectrometry detection (ICP-MS, ID-MS, SIMS and HPLC-ICP-MS) with 4.9% of the publications have been also employed to speciate Cr(VI) and Cr(III).

35.8

FINAL CONSIDERATIONS

Speciation of chromium has focussed on the determination of the two common oxidation states of this element and, in most cases, speciation studies were based on the cationic nature of Cr(III) and anionic Cr(VI), thus involving ionic separation and/or selective complexation sample treatment. However, there is a lack of literature on the determination of speciﬁc chemical forms of chromium. Amongst the strategies suggested for Cr speciation in waters, those based on chromium preconcentration and selective elution of its ionic forms from solid supports are probably the most interesting in order to provide accurate determinations for in-ﬁeld sampling and stabilization of microamounts of Cr(III) and Cr(VI) and selective and sensitive determination based on online elution followed by as sensitive as possible detection. Additionally, the electrostacking methodology developed by He et al. [520] provides a single and convenient means of online hyphenation of cation-based and anion-based methodologies, such as ICP-MS and ion chromatography, suitable for the simultaneous determination of Cr(III) and Cr(VI) species. However, this aspect has not yet developed, currently relying on sequential measurements by F-AAS or GF-AAS.

1157

M. de la Guardia and A. Morales-Rubio

Considering analysis of body ﬂuids, it is astonishing to see the absence of complete studies involving the recognition of transport proteins and speciﬁc sites of Cr activity in biochemical processes. These aspects require in-depth studies for which the use of gel permeation preparative chromatography followed by hyphenated methods of HPLC and atomic or ionic spectrometry could probably improve the state of the art of our knowledge. For the speciation of Cr in solid samples, it is absolutely necessary to insist on the development of accurate and well-validated methodologies to assure the quantitative recovery of species without changing their nature in the bulk sample. Thus, recovery studies on each species and mixtures of species would contribute to improvement in the conﬁdence of the reported results and to eradicate doubts about matrix – reagent and matrix – species interactions during the pretreatment steps. Validations of methodologies available for Cr speciation are the key points and for that it is absolutely necessary that the use of reference materials for Cr species be developed. In this sense, the preparation of stabilized solutions containing Cr(III) and Cr(VI) made by the BCR [521], and that of the same institution concerning a lyophilized solution containing Cr(III) and Cr(VI) [522] are of tremendous interest in order to be able to conﬁrm the applicability of academic studies.

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14

1158

F. Burriel-Martı´, F. Lucena-Conde, S. Arribas-Jimeno and J. Herna´ndez-Mendez, Quı´mica Analı´tica Cualitativa, 1st edn. Paraninfo, Madrid, 1985. F.A. Cotton and G.W. Wilkinson, Advanced Inorganic Chemistry, 2nd edn. Wiley, New York, 1966. R. Scholder and H. Schwarz, Z. Inorg. Chem., 326 (1963) 11. F. Bafﬁ, M. Ravera, M.C. Lanni, F. Soggia and E. Magi, Anal. Chim. Acta, 306 (1995) 149. M. Bernhard, F.E. Brinckman and P.J. Sadler (Eds.) The Importance of Chemical Speciation in Environmental Processes. Springer, Berlin, 1986. E. Sterlinska and W. Golebiewska, Chem. Anal. (Warsaw), 39 (1994) 77. R.A. Anderson, Trace Elements in Health and Disease. Norstedt, Stockholm, 1985. J.O. Nriagu and E. Nielboer (Eds.) Chromium in Natural and Human Environments. Wiley, New York, 1988. M. Pourbaix, Atlas of Electrochemical Equilibria in Aqueous Solutions. Pergamon Press, Oxford, 1966. L. Campanella, in: S. Caroli (Ed.), Element Speciation in Bioinorganic Chemistry. Wiley, New York, 1996. M. Sperling, in: A. Townshend (Ed.), Encyclopedia of Analytical Sciences. Academic Press, London, 1995. G.E. Batley, Trace Element Speciation. Analytical Methods and Problems. CRC Press, Boca Raton, FL, 1989. B. Deng and J.X. Luo, Fenxi Shiyanshi, 13 (1994) 4. H.M. Kingston, D. Huo, Y. Lu and S. Chalk, Spectrochim. Acta B, 53 (1998) 299.

Sample preparation for chromium speciation 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53

H. Groll, G. Schaldach, H. Berndt and K. Niemax, Spectrochim. Acta B, 50 (1995) 1293. A. Gaspar, C. Sogor and J. Posta, Fresenius J. Anal. Chem., 363 (1999) 480. F. Sacher, B. Raue, J. Klinger and H.J. Brauch, Int. J. Environ. Anal. Chem., 74 (1999) 191. W. Tirler and W. Bloedorn, LaborPraxis, 22 (1998) 52. J. Posta, A. Gaspar, R. Toth and L. Ombodi, Magy. Kem. Foly., 102 (1996) 535. A. Gaspar, J. Posta and R. Toth, J. Anal. At. Spectrom., 11(11) (1996) 1067. J. Posta, A. Gaspar, R. Toth and L. Ombodi, Fresenius J. Anal. Chem., 355 (1996) 719. S.X. Li, G.Q. Huang and S.H. Quan, Fenxi Kexue Xuebao, 11 (1995) 42. N. Jakubowski, B. Jepkens, D. Stuewer and H. Berndt, J. Anal. At. Spectrom., 9 (1994) 193. J. Posta, H. Berndt, S.K. Luo and G. Schaldach, Anal. Chem., 65 (1993) 2590. C.A. Johnson, Anal. Chim. Acta, 238 (1990) 273. S. Peraniemi and M. Ahlgren, Anal. Chim. Acta, 315 (1995) 365. J.L. He, Fenxi Huaxue, 23 (1995) 1047. Y. Inoue, T. Sakai and H. Kumagai, J. Chromatogr. A, 706 (1995) 127. G.S. Qian, Lihua Jianyan Huaxue Fence, 37 (2001) 180. P. Liang, C.X. Li, Y.C. Qin, B. Hu and Z.C. Jiang, Fenxi Kexue Xuebao, 16 (2000) 300. M. Pantsar-Kallio and P.K.G. Manninen, Anal. Chim. Acta, 318 (1996) 335. L.Y. Qiu and E.Y. Zhao, Guangpuxue Yu Guangpu Fenxi, 13(5) (1993) 111. Y.L. Wu, B. Hu, T.Y. Peng and Z.C. Jiang, Fresenius J. Anal. Chem., 370 (2001) 904. E. Vassileva, Analusis, 28 (2000) 878. W.H. Ma, R.X. Cai and D.H. Chen, Lab. Rob. Autom., 11 (1999) 141. E. Vassileva, K. Hadjiivanov, T. Stoychev and C. Daiev, Analyst, 125 (2000) 693. N.K. Djane, K. Ndung’u, C. Johnsson, H. Sartz, T. Tornstrom and L. Mathiasson, Talanta, 48 (1999) 1121. L. Li, Y.J. Feng and G.Q. Huang, Guangpuxue Yu Guangpu Fenxi, 18 (1998) 354. J. Posta, A. Gaspar, R. Toth and L. Ombodi, Microchem. J., 54 (1996) 195. M. Pantsar-Kallio and P.K.G. Manninen, Fresenius J. Anal. Chem., 355 (1996) 716. Q.A. Kong, Q.F. Wu and X.H. Guo, Fenxi Huaxue, 24 (1996) 1. J.L. Manzoori, M.H. Sorouraddin and F. Shemirani, Talanta, 42 (1995) 1151. A.M. Naghmush, K. Pyrzynska and M. Trojanowicz, Anal. Chim. Acta, 288 (1994) 247. M. Sperling, S. Xu and B. Welz, Anal. Chem., 64 (1992) 3101. Z. Hong, H. Chen and M. Huang, Fenxi Huaxue, 19 (1991) 484. W. Qi, J. Cao, S. Lin, S. Chen and K. Sakai, Bunseki Kagaku, 40 (1991) 39. V.K. Ososkov and A.M. Zhelezko, Izv. Vyssh. Uchebn. Zaved. Khim. Khim. Tekhnol., 34 (1991) 49. W. Qi, S. Chen and S. Lin, Guangpuxue Yu Guangpu Fenxi, 10 (1990) 46. K. Isshiki, Y. Sohrin, H. Karatani and E. Nakayama, Anal. Chim. Acta, 224 (1989) 55. A. Syty, R.G. Christensen and T.C. Rains, J. Anal. At. Spectrom., 3 (1988) 193. S. Arpadyan and V. Krivan, Anal. Chem., 58 (1986) 2611. A.G. Cox, I.G. Cook and C.W. McLeod, Analyst, 110 (1985) 331. D.E. Leyden, K. Goldbach and A.T. Ellis, Anal. Chim. Acta, 171 (1985) 369.

1159

M. de la Guardia and A. Morales-Rubio 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90

1160

T.L. Mullins, Anal. Chim. Acta, 165 (1984) 97. I.S. Krull, K.W. Panaro and L.L. Gershman, J. Chromatogr. Sci., 21 (1983) 460. Y.L. Chang and S.J. Jiang, J. Anal. At. Spectrom., 16 (2001) 858. Z. Horvath, A. Lasztity, I. Varga, E. Meszaros and A. Molnar, Talanta, 41 (1994) 1165. H. Bag, A.R. Turker, M. Lale and A. Tunceli, Talanta, 51 (2000) 895. E. Menendez-Alonso, S.J. Hill, M.E. Foulkes and J.S. Crighton, J. Anal. At. Spectrom., 14 (1999) 187. K. Hilman and T. Goto, Bunseki Kagaku, 44 (1995) 921. M. Hiraide and A. Mizuike, Fresenius J. Anal. Chem., 335 (1989) 924. A. Fajgelj and L. Kosta, Vestn. Slov. Kem. Drus., 34 (1987) 175. M.E. Romero-Gonzalez, C.J. Williams and P.H.E. Gardiner, J. Anal. At. Spectrom., 15 (2000) 1009. S.J. Kumar, P. Ostapczuk and H. Emons, At. Spectrosc., 20 (1999) 194. C. Barnowski, N. Jakubowski, D. Stuewer and J.A.C. Broekaert, J. Anal. At. Spectrom., 12 (1997) 1155. M.J. Tomlinson and J.A. Caruso, Anal. Chim. Acta, 322 (1996) 1. F.A. Byrdy, L.K. Olson, N.P. Vela and J.A. Caruso, J. Chromatogr. A, 712 (1995) 311. C.R. Lan, C.L. Tseng, M.H. Ynag and Z.B. Alfassi, Analyst, 116 (1991) 35. E.K. Paleologos, C.D. Stalikas, S.M. Tzouwara-Karayanni, G.A. Pilidis and M.I. Karayannis, J. Anal. At. Spectrom., 15 (2000) 287. T.P. Rao, S. Karthikeyan, B. Vijayalekshmy and C.S.P. Iyer, Anal. Chim. Acta, 369 (1998) 69. J. Posta, A. Alimonti, F. Petrucci and S. Caroli, Anal. Chim. Acta, 325 (1996) 185. J.W. Olesik, J.A. Kinzer and S.V. Olesik, Anal. Chem., 67 (1995) 1. A. Isozaki, T. Narukawa and T. Okutani, Bunseki Kagaku, 44 (1995) 111. M.L. Mena, A. Morales-Rubio, A.G. Cox, C.W. McLeod and P. Quevauviller, Quim. Anal., 14 (1995) 164. L.B. Allen and J.A. Koropchack, J. Chromatogr. A, 657 (1993) 192. W. Fong and J.C.G. Wu, Spectrosc. Lett., 24 (1991) 931. S.B. Roychowdhury and J.A. Koropchak, Anal. Chem., 62 (1990) 484. I.Yu. Andreeva, E.K. Bocharova, E. Danilova and A.I. Gunchenko, Zh. Anal. Khim., 44 (1989) 675. E.B. Milosavljevic, L. Solujic, J.H. Nelson and J.L. Hendrix, Mikrochim. Acta, 3 (1985) 353. G. Kura, T. Hashiguchi and T. Tarutani, Fresenius Z. Anal. Chem., 316 (1983) 716. P. Battistoni, S. Bompadre, G. Fara and G. Gobbi, Talanta, 30 (1983) 15. A. Gaspar, J. Posta and R. Toth, Magy. Kem. Foly., 103 (1997) 321. S. Ahmad, R.C. Murthy and S.V. Chandra, Analyst, 115 (1990) 287. D.J. Butcher, A. Zybin, M.A. Bolshov and K. Niemax, Rev. Anal. Chem., 20 (2001) 79. A. Zybin, G. Schaldach, H. Berndt and K. Niemax, Anal. Chem., 70 (1998) 5093. P. Rychlovsky, M. Krenzelok and R. Volhejnova, Collect. Czech. Chem. Commun., 63 (1998) 2015. P.A. Sule and J.D. Ingle Jr., Anal. Chim. Acta, 326 (1996) 85. D. Oktavec, J. Lehotay and E. Hornackova, At. Spectrosc., 16 (1995) 92. D.M. Adria-Cerezo, M. Llobat-Estelles and A.R. Mauri-Aucejo, Talanta, 51 (2000) 531. S.K. Luo and H. Berndt, Fresenius J. Anal. Chem., 360 (1998) 545.

Sample preparation for chromium speciation 91 92 93 94 95 96 97 98 99 100 101 102

103 104 105 106 107 108 109 110 111 112 113 114 115 116 117 118 119 120 121 122 123 124 125 126 127

J. Prokisch, B. Kovacs, Z. Gyori and J. Loch, J. Chromatogr. A, 683 (1994) 253. Z. Huang, X. He and J. Deng, Lihua Jianyan Huaxue Fence, 26 (1990) 100. I.T. Urasa and S.H. Nam, J. Chromatogr. Sci., 27 (1989) 30. H. De-Beer and P.P. Coetzee, S. Afr. J. Chem., 41 (1988) 152. F. Nakata, S. Hara, H. Matsuo, T. Kumamaru and S. Matsushita, Anal. Sci., 1 (1985) 157. M. Pantsar-Kallio and P.K.G. Manninen, J. Chromatogr. A, 750 (1996) 89. C.M. Andrle, N. Jakubowski and J.A.C. Broekaert, Spectrochim. Acta B, 52 (1997) 189. M.J. Powell, D.W. Boomer and D.R. Wiederin, Anal. Chem., 67 (1995) 2474. H. Luo and H. Yao, Henliang Fenxi, 9 (1993) 99. M.S. Jimenez, L. Martin, J.M. Mir and J.R. Castillo, At. Spectrosc., 17 (1996) 201. Q. Wu, L. Zhou, L. Luang, S. Zhou and Q. Chen, Lihua Jianyan Huaxue Fence, 27 (1991) 195. N.I. Shcherbinina, G.R. Ishmiyarova, I.E. Nikitina, G.V. Myasoedova, L.A. Polosukhina, I.I. Uralov, A.D. Simonva and E.Y. Daniova, Zh. Anal. Khim., 45 (1990) 766. B. Demirata, I. Tor, H. Filik and H. Afsar, Fresenius J. Anal. Chem., 356 (1996) 375. B. Demirata, Mikrochim. Acta, 136 (2001) 143. M. Sperling, X. Yin and B. Welz, Analyst, 117 (1992) 629. V. Otruba, J. Pivnicka and V. Kanicky, Collect. Czech. Chem. Commun., 65 (2000) 1865. I. Karadjova, Mikrochim. Acta, 130 (1999) 185. J. Lameiras, M.E. Soares, M.L. Bastos and M. Ferreira, Analyst, 123 (1998) 2091. L. Ajmal, R.A.K. Rao and B.A. Siddiqui, Water Res., 30 (1996) 1478. A.C. Sahayam, J. Arunachalam and S. Gangadharan, Can. J. Anal. Sci. Spectrosc., 43 (1998) 4. J.C. Yu, X.J. Wu and Z.L. Chen, Anal. Chim. Acta, 436 (2001) 59. A.F. Xue, S.H. Qian, G.Q. Huang and L.Y. Chen, J. Anal. At. Spectrom., 15 (2000) 1513. S. Dahbi, M. Azzi and M. de-la-Guardia, Fresenius J. Anal. Chem., 363 (1999) 404. A. Gaspar and J. Posta, Anal. Chim. Acta, 354 (1997) 151. S. Nielsen and E.H. Hansen, Anal. Chim. Acta, 366 (1998) 163. A. Cuesta, J.L. Todoli and A. Canals, Spectrochim. Acta B, 51 (1996) 1791. H. Liang, H. He and Z. Zhang, Fenxi Huaxue, 18 (1990) 1142. K. Vercoutere, R. Cornelis, S. Dyg, L. Mees, J.M. Christensen, K. Byrialsen, B. Aaen and P. Quevauviller, Mikrochim. Acta, 123 (1996) 109. S. Dyg, R. Cornelis, B. Griepink and P. Quevauviller, Anal. Chim. Acta, 286 (1994) 297. T. Yabutani, K. Chiba and H. Haraguchi, Bull. Chem. Soc. Jpn., 74 (2001) 31. C. Minoia, A. Mazzucotelli, A. Cavalleri and V. Minganti, Analyst, 108 (1983) 481. S. Hirata, K. Honda, O. Shikino, N. Maekawa and M. Aihara, Spectrochim. Acta B, 55 (2000) 1089. S. Hirata, Y. Umezaki and M. Ikeda, Anal. Chem., 58 (1986) 2602. A. Isozaki, K. Kumagai and S. Utsumi, Anal. Chim. Acta, 153 (1983) 15. I. Narin, M. Soylak, L. Elci and M. Dogan, Talanta, 52 (2000) 1041. F.Y. Saleh, G.E. Mbamalu, Q.H. Jaradat and C.E. Brungardt, Anal. Chem., 68 (1996) 740. F. Bafﬁ, A.M. Cardinale and R. Bruzzone, Anal. Chim. Acta, 270 (1992) 79.

1161

M. de la Guardia and A. Morales-Rubio 128 129 130 131 132 133 134 135 136 137 138 139 140 141 142 143 144 145 146 147 148 149 150 151 152 153 154 155 156 157 158 159 160 161 162 163

1162

N.N. Basargin, V.Y. Anikin, V.D. Salikhov and Y.G. Rozovskii, Zavod. Lab. Diagn. Mater., 66 (2000) 14. P.M. Padilha, J.C. Rocha, J.C. Moreira, J.T.S. Campos and C.C. Federici, Talanta, 45 (1997) 317. R.M. Cespon-Romero, M.C. Yebra-Biurrun and M.P. Bermejo-Barrera, Anal. Chim. Acta, 327 (1996) 37. M. Ozcan, S. Akman, C. Erbil and S. Sarac, Fresenius J. Anal. Chem., 355 (1996) 665. K.H. Lee, M. Oshima, T. Takayanagi and S. Motomizu, Anal. Sci., 16 (2000) 731. K.H. Lee, M. Oshima and S. Motomizu, Bunseki Kagaku, 49 (2000) 529. E. Beinrohr, A. Manova and J. Dzurov, Fresenius J. Anal. Chem., 355 (1996) 528. B. Pasullean, C.M. Davidson and D. Littlejohn, J. Anal. At. Spectrom., 10 (1995) 241. O. Abollino, C. Sarzanini, E. Mentasti and A. Liberatori, Spectrochim. Acta A, 49 (1993) 1411. I. Kubrakova, T. Kudinova, A. Formanovsky, N. Kuz’min, G. Tsysin and Y. Zolotov, Analyst, 119 (1994) 2477. A.G. Cox and C.W. McLeod, Anal. Chim. Acta, 179 (1986) 487. E. Vassileva and N. Furuta, Fresenius J. Anal. Chem., 370 (2001) 52. I. Narin, M. Soylak, L. Elci and M. Dogan, Anal. Lett., 34 (2001) 1935. J.X. Chang, X.Y. Luo, Z.X. Su, G.Y. Zhan and W.Y. Gao, Fresenius J. Anal. Chem., 349 (1994) 438. A.M. Naghmush, K. Pyrzynska and M. Trojanowicz, Anal. Chim. Acta, 288 (1994) 247. M. Foldesova, P. Dillinger and P. Lukac, J. Radioanal. Nucl. Chem., 245 (2000) 435. S. Tokalioglu, S. Kartal and L. Elci, Anal. Sci., 16 (2000) 1169. G. Zoorob, M.J. Tomlinson, J.S. Wang and J.A. Caruso, J. Anal. At. Spectrom., 10 (1995) 853. D. Jensen and W. Bloedorn, GIT Fachz. Lab., 39 (1995) 654. M.J. Marques, A. Morales-Rubio, A. Salvador and M. de la Guardia, Talanta, 53 (2001) 1229. M. Zhuang and R.M. Barnes, Spectrochim. Acta B, 38 (1983) 259. M. Korn, M.G.A. Korn, B.F. Reis and E. de Oliveira, Talanta, 41 (1994) 2043. M. Yamamoto and T. Honjo, Bunseki Kagaku, 50 (2001) 567. E.K. Paleologos, C.D. Stalikas and M.I. Karayannis, Analyst, 126 (2001) 389. Y. Hou, X. Chi and S. Lin, Guangpuxue Yu Guangpu Fenxi, 9 (1989) 51. C. Yoshimura and T. Huzino, Hyomen Gijutsu, 41 (1990) 318. D. Wu, Z. Zhou and G. Wang, Fenxi Huaxue, 15 (1987) 208. D. Baralkiewicz and J. Siepak, Chem. Anal. (Warsaw), 44 (1999) 879. S.C. Nielsen, S. Sturup, H. Spliid and E.H. Hansen, Talanta, 49 (1999) 1027. A. Gaspar and J. Posta, Magy. Kem. Foly., 102 (1996) 503. E.E. Menden, F.H. Rutland and W.E. Kallenberger, J. Am. Leather Chem. Assoc., 85 (1990) 363. K.S. Subramanian, J. Res. Natl. Bur. Stand., 93 (1988) 305. M. He, Yankuang Ceshi, 6 (1987) 250. X. Chi, M. Tan and L. Zhan, Guangpuxue Yu Guangpu Fenxi, 5 (1985) 40. T.P. Hsieh and L.K. Liu, Anal. Chim. Acta, 282 (1993) 221. M. Tsuji, K. Kawasaki, T. Niizeki, M. Saitou and T. Hattori, Int. J. PIXE, 10 (2000) 57.

Sample preparation for chromium speciation 164 165 166 167 168 169 170 171 172 173 174 175 176 177 178 179 180 181 182 183 184 185 186 187 188 189 190 191 192 193 194 195 196 197 198 199 200 201 202 203 204 205

J. Pavel, J. Kliment, S. Stoerk and O. Suter, Fresenius J. Anal. Chem., 321 (1985) 587. C.M. Wai, L.M. Tsay and J.C. Yu, Mikrochim. Acta, 2 (1987) 73. M. Yamada, S. Kusakabe, J. Prekopova and T. Sekine, Anal. Sci., 12 (1996) 405. Y.S. Fung and W.C. Sham, Analyst, 119 (1994) 1029. G.Q. Huang, S.H. Qian and S. Zhong, Fenxi Kexue Xuebao, 11 (1995) 32. T. Balaji, P. Chiranjeevi and G.R.K. Naidu, Anal. Lett., 31 (1998) 1081. M. Yamada and T. Sekine, Solvent Extr. Res. Dev. Jpn., 399 (1996) 108. M.J. Gardner and J.E. Ravenscroft, Fresenius J. Anal. Chem., 354 (1996) 602. Q.B. Zhang, H. Minami, S. Inoue and I. Atsuya, Anal. Chim. Acta, 401 (1999) 277. E. Beceiro-Gonzalez, J. Barciela-Garcia, P. Bermejo-Barrera and A. BermejoBarrera, Fresenius J. Anal. Chem., 344 (1992) 301. E. Beceiro-Gonzalez, P. Bermejo-Barrera, A. Bermejo-Barrera, J. Barciela-Garcia and C. Barciela-Alonso, J. Anal. At. Spectrom., 8 (1993) 649. J. Ueda, H. Satoh and S. Kagaya, Anal. Sci., 13 (1997) 613. J.L. Manzoori and F. Shemirani, J. Anal. At. Spectrom., 10 (1995) 881. J. Obiols, R. Devesa, J. Garcia-Berro and J. Serra, Int. J. Environ. Anal. Chem., 30 (1987) 197. Y. Shan, Lihua Jianyan Huaxue Fence, 25 (1989) 93. F. Sugimoto, N. Tsunekage and Y. Maeda, Nippon Kagaku Kaishi, 6 (1992) 667. D. Atanassova, V. Stefanova and E. Russeva, Talanta, 47 (1998) 1237. S. Long and L. Zhang, Fenxi Shiyanshi, 5 (1986) 8. L. Li and G.Q. Huang, Lihua Jianyan Huaxue Fence, 30 (1994) 160. T. Yabutani, S. Ji, F. Mouri, A. Itoh, K. Chiba and H. Haraguchi, Bull. Chem. Soc. Jpn., 73 (2000) 895. B. Ouddane, M. Skiker, J.C. Fischer and M. Wartel, Analusis, 25 (1997) 308. S. Kagaya and J. Ueda, Bull. Chem. Soc. Jpn., 68 (1995) 2843. A. Boughriet, L. Deram and M. Wartel, J. Anal. At. Spectrom., 9 (1994) 1135. S. Long and L. Han, Fenxi Huaxue, 14 (1986) 529. J. Anwar, M.A. Aslam and M. Ahmad, J. Chem. Soc. Pak., 18 (1996) 287. H.G. Beere and P. Jones, Anal. Chim. Acta, 293 (1994) 237. T. Williams, P. Jones and L. Ebdon, J. Chromatogr., 482 (1989) 361. W. Shi and T. Hu, Lihua Jianyan Huaxue Fence, 34 (1998) 447. S.B. Savvin, O.P. Shvoeva, V.P. Dedkova, S.V. Rudakov and A.V. Mikhailova, Zh. Anal. Khim., 51 (1996) 308. J. Lu, X. Zhang, B. Zhang, W. Qin and Z. Zhang, Gaodeng Xuexiao Huaxue Xuebao, 14 (1993) 771. F.L. Zhou and Y.M. Gao, Fenxi Huaxue, 20 (1992) 806. T.A. Bol’shova, A.Yu. Malykhin and E.M. Basova, Vestn. Mosk. Univ. Ser. 2 Khim., 30 (1989) 172. H.L. Liu, S.S. Liu and Y.M. Chen, Fenxi Huaxue, 26 (1998) 963. E. Pobozy, E. Wojasinska and M. Trojanowicz, J. Chromatogr. A, 736 (1996) 141. H.J. Fan and B. Qu, Fenxi Kexue Xuebao, 11 (1995) 37. E. Andres-Garcia and D. Blanco-Gomis, Analyst, 122 (1997) 899. M. Bittner and J.A.C. Broekaert, Anal. Chim. Acta, 364 (1998) 31. S. Pozdniakova and A. Padarauskas, Analyst, 123 (1998) 1497. T. Prochackova and M. Foltin, Chem. Listy, 91 (1997) 585. Z.X. Guo, Fenxi Shiyanshi, 16 (1997) 14. L.Q. Li, C.D. Wu, H.G. Zheng and Y.D. Li, Fenxi Huaxue, 23 (1995) 242. H. de-Beer and P.P. Coetzee, S. Afr. J. Chem., 44 (1991) 105.

1163

M. de la Guardia and A. Morales-Rubio 206 207 208 209 210 211 212 213 214 215 216 217 218 219 220 221 222 223 224 225 226 227 228 229 230 231 232 233 234 235 236 237 238 239 240 241 242 243 244 245 246 247 248

1164

Z. Huang, X. He and Z. Wu, Fenxi Huaxue, 16 (1988) 1101. S. Du, S. Song and Z. Huang, Fenxi Huaxue, 12 (1984) 382. O.P. Semenova, A.R. Timerbaev, R. Gaegstadter and G.K. Bonn, J. High Resolut. Chromatogr., 19 (1996) 177. J.F. Jen, G.L. Ou-Yang, C.S. Chen and S.M. Yang, Analyst, 118 (1993) 1281. C.M. Andrle and J.A.C. Broekaert, Fresenius J. Anal. Chem., 346 (1993) 653. C.S. Lin, X.S. Zhang and H. You, Fenxi Shiyanshi, 11 (1992) 18. C. Lin and X. Zhang, Analyst, 112 (1987) 1659. F. Zhang and G. Chen, Fenxi Huaxue, 12 (1984) 740. P.Y. Gao, X.X. Gu and T.Z. Zhou, Anal. Chim. Acta, 332 (1996) 307. F.J. Zou, Fenxi Shiyanshi, 14 (1995) 29. J. Yang, Lihua Jianyan Huaxue Fence, 23 (1987) 341. W. Qi and L. Zhu, Talanta, 33 (1986) 694. G. Xi and D. Liang, Fenxi Huaxue, 12 (1984) 209. V. Maheswari and N. Balasubramanian, Chem. Anal. (Warsaw), 41 (1996) 569. Zh.M. Arstamyan and R.S. Karinyan, Arm. Khim. Zh., 43 (1990) 442. Y.B. Ren, K. Zhao and W.X. Li, Fenxi Shiyanshi, 17 (1998) 51. K. Zhao, Y.B. Ren and W.X. Li, Fenxi Huaxue, 25 (1997) 742. S.P. Zhou, G.Y. Yang and J.Y. Yin, Fenxi Huaxue, 28 (2000) 890. H. Zhao, Y.Q. Xiang, X.G. Chen, Z.D. Hu, X.Z. Wu and Y. Nagaosa, Anal. Sci., 15 (1999) 1129. Q.H. Zhao, Y.H. Xu, Q.H. Xu and Q. Li, Lihua Jianyan Huaxue Fence, 33 (1997) 360. W.R. Yang, Q. Li, Q.H. Xu and H.T. Wang, Fenxi Huaxue, 24 (1996) 243. H.T. Wang, J. Pan and Q.H. Xu, Fenxi Huaxue, 24 (1996) 195. Q.E. Cao, Y.H. Liang, Q.H. Xu and Q. Li, Fenxi Huaxue, 23 (1995) 525. Q. Li and Q.H. Xu, Fenxi Huaxue, 22 (1994) 393. H. Hu, Lihua Jianyan Huaxue Fence, 28 (1992) 92. Y. Yang and Y. Chen, Fenxi Huaxue, 19 (1991) 92. J. Sun, M. Zhao, J. Gao, H. Jiang and Q. Liu, Fenxi Shiyanshi, 7 (1988) 9. M. Ding and J. Zhang, Huaxue Tongbao, 2 (1988) 22. M.H. Zhang, Q.R. Zhang, Z. Fang and Z.C. Lei, Talanta, 48 (1999) 369. S.X. Cao and M.B. Zhang, Fenxi Shiyanshi, 18 (1999) 79. C. Agarwal, K.S. Patel and R.K. Mishra, Int. J. Environ. Anal. Chem., 36 (1989) 95. A. Golwelker, K.S. Patel and R.K. Mishra, Bull. Chem. Soc. Jpn., 63 (1990) 605. S.X. Cao and M.G. Zhang, J. Trace Microprobe Tech., 17 (1999) 157. L.F. Huang and B.W. Wang, Fenxi Huaxue, 25 (1997) 208. Dionex Technical Note, Technical Note 26, July 2000, p. 6. Dionex Application Note, Application Note 80, July 2000, p. 2. Y.M. Li and H.M. Shen, Lihua Jianyan Huaxue Fence, 32 (1996) 175. Z.Q. Wu, Lihua Jianyan Huaxue Fence, 28 (1992) 337. Y.P. Chen, J. Zhang, H.Y. Peng, B. Yao and P.Z. Li, Gaodeng Xuexiao Huaxue Xuebao, 13 (1992) 459. R. Parkash, R. Bansal, A. Kaur and S.K. Rehani, Talanta, 38 (1991) 1163. Z.J. Yang, F. Liang, C.Y. Wang, H.S. Zhang and J.K. Cheng, Fenxi Kexue Xuebao, 16 (2000) 371. Y.X. Miao, H. Wang, H.S. Zhang and J.K. Cheng, Fenxi Shiyanshi, 13 (1994) 1. F. Zhou and T. Liu, Yankuang Ceshi, 7 (1988) 278.

Sample preparation for chromium speciation 249 250 251 252 253 254 255 256 257 258 259 260 261 262 263 264 265 266 267 268 269 270 271 272 273 274 275 276 277 278 279 280 281 282 283 284 285 286 287 288 289 290 291 292 293 294 295

Y. Shi, Y. Feng and D. Sun, Fenxi Shiyanshi, 9 (1990) 13. K.L. Liang and Q.H. Duan, Yejin Fenxi, 11 (1991) 44. P.K. Tarafder, S.B. Singh and D.P.S. Rathore, Fresenius J. Anal. Chem., 354 (1996) 124. X.L. Wu and J. Gao, Fenxi Huaxue, 26 (1998) 248. G. Fang and C. Miao, Analyst, 110 (1985) 65. Z. Ou, X. Lu and J. Pan, Gaodeng Xuexiao Huaxue Xuebao, 14 (1993) 486. D.W. Huang, Y.Y. Lu and J.H. Cheng, Fenxi Shiyanshi, 20 (2001) 72. Q. Lu and R.X. Cai, Fenxi Kexue Xuebao, 17 (2001) 138. D.L. Zhang, C.H. Cheng and X.W. Du, Fenxi Kexue Xuebao, 16 (2000) 481. C.B. Xia, Fenxi Huaxue, 28 (2000) 784. C.B. Xia, Fenxi Ceshi Xuebao, 19 (2000) 56. J.H. Yu and Q.T. Yang, Fenxi Huaxue, 27 (1999) 1309. Z.Q. Zhao, R.M. Gao, J.T. Li, S.R. Liu and H. Liu, Microchem. J., 58 (1998) 1. J.Y. Sun, L. Sun and H.F. Xia, Lihua Jianyan Huaxue Fence, 33 (1997) 35. J.H. Yu, R.Z. Cui and L.B. Wang, Fenxi Shiyanshi, 12 (1993) 43. R.M. Gao, Z.Q. Zhao, Q.Z. Zhou and D.X. Yuan, Talanta, 40 (1993) 637. P.C.C. Oliveira and J.C. Masini, Analyst, 123 (1998) 2085. Q.Z. Qu and J.H. Li, Lihua Jianyan Huaxue Fence, 31 (1995) 39. M. Hou and M. Shao, Fenxi Shiyanshi, 19 (2000) 1. C.S. Devaragudi and K. Hussain-Reddy, Talanta, 44 (1997) 1973. S. Osaki, T. Osaki and Y. Takashima, Talanta, 30 (1983) 683. X.D. Liu, S.P. Liu and Y.M. Huang, Fenxi Huaxue, 23 (1997) 1202. H.Z. Wu, C.Y. Qiu and Z.S. Zheng, Fenxi Huaxue, 27 (1999) 668. H.Z. Wu, Y.L. Liu, Z.L. Chen and Z.S. Zheng, Fenxi Kexue Xuebao, 15 (1999) 213. H.Z. Wu, Z.R. Wu, J. Lin and Z.S. Zheng, Fenxi Shiyanshi, 18 (1999) 26. X.C. Wu and T.Y. Liu, Fenxi Shiyanshi, 14 (1995) 15. L.L. Ma and C.Y. Yang, Fenxi Huaxue, 27 (1999) 990. C.J. Liu and J.J. Jiang, Lihua Jianyan Huaxue Fence, 29 (1993) 230. Z. Liu and X.Y. Wang, Fenxi Huaxue, 24 (1996) 164. D. Li, Lihua Jianyan Huaxue Fence, 25 (1989) 221. A.A. Mohamed, S.A. Ahmed and M.F. El-Shahat, J. Trace Microprobe Tech., 19 (2001) 297. H.Z. Wu, Z.S. Zheng, G.B. Hu and F. Wang, Fenxi Huaxue, 20 (1992) 1445. R.H. He and J.H. Wang, Fenxi Shiyanshi, 19 (2000) 24. C. Reis, J. Carlos-de-Andrade, R.E. Bruns and R.C.C.P. Moran, Anal. Chim. Acta, 369 (1998) 269. F. Alba, J.M. Estela and V. Cerda, Quim. Anal. (Barcelona), 7 (1988) 219. Y. Cao, Fenxi Shiyanshi, 10 (1991) 22. J. Wang, R. He and J. Wang, Lihua Jianyan Huaxue Fence, 27 (1991) 215. S. Liu and F. Wang, Talanta, 38 (1991) 801. Z.M. Zhang, Fenxi Huaxue, 26 (1998) 325. S. Liu, M. Zhao and C. Deng, J. Chromatogr., 598 (1992) 298. Z.M. Xue, X.D. Zhao and S.H. Zhang, Lihua Jianyan Huaxue Fence, 33 (1997) 121. F. Zeng, K.Y. Cui and W. Lu, Lihua Jianyan Huaxue Fence, 33 (1997) 57. Z. Bei, J. Yan and X. Hong, Lihua Jianyan Huaxue Fence, 26 (1990) 357. N. Yang, W.Y. Zhou and E.P. He, Lihua Jianyan Huaxue Fence, 32 (1996) 353. Z.M. Xue, X.R. Wang and Y.T. Wang, Lihua Jianyan Huaxue Fence, 36 (2000) 16. X. Wang, Fenxi Huaxue, 12 (1984) 943. R. Anton, H. Marini and R. Olsina, Anal. Sci., 11 (1995) 431.

1165

M. de la Guardia and A. Morales-Rubio 296 297 298 299 300 301 302 303 304 305 306 307 308 309 310 311 312 313 314 315 316 317 318 319 320 321 322 323 324 325 326 327 328 329 330 331 332 333 334 335 336 337

1166

Z. Yi and H. Gang, Talanta, 41 (1994) 1247. Z.X. Guo, L.J. Yu and S.L. Zhang, Lihua Jianyan Huaxue Fence, 34 (1998) 62. J. Chen, J. Chai and D. Wang, Huaxue Shiji, 12 (1990) 54. E.K. Paleologos, C.D. Stalikas, S.M. Tzouwara-Karayanni and M.I. Karayannis, Anal. Chim. Acta, 436 (2001) 49. Y.S. Yan, J.Y. Wei, K.Q. Shi, X.G. Zhou and Z.J. Tan, Lihua Jianyan Huaxue Fence, 31 (1995) 98. Y.J. Park and J.K. Hardy, J. Chromatogr., 481 (1989) 287. R.M. Gao and H.G. Liu, Fenxi Huaxue, 21 (1993) 306. W. Qi and B. Pu, Huaxue Xuebao, 42 (1984) 27. G. Fang and J. Luo, Fenxi Huaxue, 13 (1985) 769. S.P. Liu, M.Q. Zhao and C.Y. Deng, Talanta, 41 (1994) 279. M. Saitoh, K. Furuya, H. Inoue and T. Shirai, Bunseki Kagaku, 38 (1989) 331. J.L. Manzoori, M.H. Sorouraddin and F. Shemiran, Anal. Lett., 29 (1996) 2007. B. Baraj, M. Martinez, A. Sastre and M. Aguilar, J. Chromatogr. A, 695 (1995) 103. A.A.Y. El-Sayed, E.A. Saad, B.M.M. Ibrahime and M.T.M. Zaki, Mikrochim. Acta, 135 (2000) 19. B. Du, C.N. Luo, Q. Wei and Y.L. Dan, Lihua Jianyan Huaxue Fence, 30 (1994) 359. J.B. Lu, J.H. Wang and S. Wang, Lihua Jianyan Huaxue Fence, 35 (1999) 392. Q.L. Yang and S.P. Liu, Yejin Fenxi, 13 (1993) 7. M.J. Gonzalez-Alvarez, M.E. Diaz-Garcia and A. Sanz-Medel, Talanta, 36 (1989) 919. H.Q. Yang, S.Q. Mu, G. Zhang, X.Q. Wan and L. Li, Lihua Jianyan Huaxue Fence, 32 (1996) 298. J. Feng, Z. Wang, B. Zhang and D. Liang, Fenxi Huaxue, 11 (1983) 348. X.L. Zhao, G.Q. Gong and Y. Wang, Fenxi Shiyanshi, 12 (1993) 43. T.S. Chen, Z.G. Chen and D. Wu, Fenxi Huaxue, 22 (1994) 129. H. Ji, G. Gong and H. Wang, Fenxi Huaxue, 17 (1989) 841. H.Y. Ma, H.G. Tuo, X. Li and X.Q. Yao, Fenxi Shiyanshi, 20 (2001) 80. J.S. Shen and X.R. Li, Fenxi Huaxue, 29 (2001) 944. S.L. Geng, R.Y. Wang and J. Fan, Fenxi Huaxue, 28 (2000) 61. V. Kabasakalis, Anal. Lett., 26 (1993) 2269. N.Q. Jie, J.H. Rang and J. Guo, Anal. Lett., 25 (1992) 1447. S.L. Feng, A.N. Tang and J. Fan, Fenxi Huaxue, 29 (2001) 558. S.P. Liu, R. Yang and Z.F. Liu, Fenxi Huaxue, 26 (1998) 1432. N. Jie and J. Jiang, Analyst, 116 (1991) 395. N.Q. Jie, J.H. Yang, W. Fu and R.X. Zhang, Fenxi Huaxue, 22 (1994) 864. L.H. Chen and D.S. Wang, Fenxi Huaxue, 25 (1997) 120. N. Jie and W. Yu, Yingyong Huaxue, 7 (1990) 71. P. Campins-Falco, L.A. Tortajada-Genaro and F. Bosch-Reig, Talanta, 55 (2001) 403. A. Economou, A.K. Clark and P.R. Fielden, Anal. Commun., 35 (1998) 389. G.Z. Fang and D.D. Wang, Fenxi Ceshi Xuebao, 16 (1997) 27. R. Escobar, Q.X. Lin, A. Guiraum and F.F. de-la-Rosa, Int. J. Environ. Anal. Chem., 61 (1996) 169. H.S. Zhang, X.C. Yang and L.P. Wu, Fenxi Huaxue, 23 (1995) 1148. R. Escobar, Q.X. Lin, A. Guiraum and F.F. de la Rosa, Analyst, 118 (1993) 643. Z. Zheng and Z. Wang, Fenxi Huaxue, 12 (1984) 10. D. Prugo and N. Radic, Hem. Ind., 37 (1983) 179.

Sample preparation for chromium speciation 338 339 340 341 342 343 344 345 346 347 348 349 350 351 352 353 354 355 356 357 358 359 360 361 362 363 364 365 366 367 368 369 370

G.H. Li and Z.N. Yu, Huaxue Tongbao, 4 (1992) 42. Z.J. Zhang, W. Qin and S.N. Liu, Anal. Chim. Acta, 318 (1995) 71. J. Lu, X. Zhang and Z. Zhang, Fenxi Huaxue, 20 (1992) 61. F. Zhang and Y. Chen, Fenxi Shiyanshi, 5 (1986) 1. T. Kamidate, K. Yamaguchi, T. Segawa and H. Watanabe, Anal. Sci., 5 (1989) 429. H.Y. Han, Z. He, Q.Y. Luo and Y. Zeng, Lihua Jianyan Huaxue Fence, 34 (1998) 295. P. Janos, Fresenius J. Anal. Chem., 342 (1992) 195. A. Padarauskas, A. Judzentiene, E. Naujalis and V. Paliulionyte, J. Chromatogr. A, 808 (1998) 193. M. Foltin, T. Prochakova and J. Kandrac, Chem. Listy, 93 (1999) 399. S. Matsuoka, Y. Tennichi, K. Takehara and K. Yoshimura, Analyst, 124 (1999) 787. N. Balasubramanian and V. Maheswari, J. AOAC Int., 79 (1996) 989. Y. Luo, S. Nakano, D.A. Holman, J. Ruzicka and G.D. Christian, Talanta, 44 (1997) 1563. J. Ruz, A. Rios, M.D. Luque de Castro and M. Valcarcel, Fresenius J. Anal. Chem., 322 (1985) 499. J.C. de Andrade, J.C. Rocha and N. Baccan, Analyst, 110 (1985) 197. Q. Wei, S. Wang and L. Wang, Gaodeng Xuexiao Huaxue Xuebao, 11 (1990) 140. D.L. Tsalev and Z.K. Zaprianov, Atomic Absorption Spectrometry in Occupational and Environmental Health Practice. CRC Press, Boca Raton, FL, 1984. R.L. Zielhuis, Exposure Limits to Metals for the General Population. CEC WHO, Amsterdam, 1981. E.A. Sheard, M.W. Johnson and R.J. Caster, in: A.C. Brown and R.G. Crounse (Eds.), Hair Trace Elements and Humans Illness. Praeger, New York, 1980. H. Shahristani, K.M. Shihab and M. Jalil, Nuclear Activation Techniques in the Life Sciences. IAEA, Vienna, 1979. H. Teraoka, Arch. Environ. Health., 36 (1981) 155. R.A. Anderson, Sci. Total Environ., 17 (1981) 13. S. Langard, N. Gundersen, D.L. Tsalev and B. Gylseth, Acta Pharmacol. Toxicol., 42 (1978) 142. S. Langard and A. Hensten-Pettersen, in: D.F. Williams (Ed.), Systemic Aspects of Biocompatibility. CRC Press, Boca Raton, FL, 1981. S. Langard, Biol. Trace Element Res., 1 (1979) 45. S. Langard and T. Norseth, in: L. Friberg, G.F. Nordberg and V.B. Vouk (Eds.), Handbook of the Toxicology of Metals. Elsevier, Amsterdam, 1979. G.K. Murthy, Crit. Rev. Environ. Control, 4 (1974) 1. G. Pethes, Elemental Analysis of Biological Materials. IAEA, Vienna, 1980. D. Rondia, in: E. Diferrante (Ed.), Trace Metals Exposure and Health Effects. Pergamon Press, Oxford, 1979. I.M.M. Kenawy, M.A.H. Hafez, M.A. Akl and R.R. Lashein, Anal. Sci., 16 (2000) 493. J. Lameiras, M.E. Soares, M.L. Bastos and M. Ferreira, Analyst, 123 (1998) 2091. Z.H. Wang, M. Song, Q.L. Ma, H.M. Ma and S.C. Liang, Mikrochim. Acta, 134 (2000) 95. Y. Suzuki, J. Chromatogr. Biomed. Appl., 59 (1987) 317. J.L. Manzoori and A. Saleemi, J. Anal. At. Spectrom., 9 (1994) 337.

1167

M. de la Guardia and A. Morales-Rubio 371 372 373 374 375 376 377 378 379 380 381 382 383 384 385 386 387 388 389 390 391 392 393 394 395 396 397 398 399 400 401 402 403 404 405 406 407 408 409 410 411 412

1168

I.M.M. Kenawy, M.A.H. Hafez, M.A. Akl and R.R. Lashein, Anal. Sci., 16 (2000) 493. D. Jensen and W. Bloedorn, GIT Fachz. Lab., 39 (1995) 654. J.R. Pretty, E.A. Blubaugh, J.A. Caruso and T.M. Davidson, Anal. Chem., 66 (1994) 1540. M.J. Tomlinson, J. Wang and J.A. Caruso, J. Anal. At. Spectrom., 9 (1994) 957. B. Neidhart and C. Tausch, Mikrochim. Acta, 109 (1992) 137. X. Shan, Y. Zheng and Z. Ni, At. Spectrosc., 11 (1990) 116. S. Long and L. Han, Fenxi Huaxue, 14 (1986) 529. A.G. Cox and C.W. McLeod, Anal. Chim. Acta, 179 (1986) 487. K. Dungs, H. Fleischhauer and B. Neidhart, Fresenius J. Anal. Chem., 322 (1985) 280. L. Ping, K. Matsumoto and K. Fuwa, Anal. Chem., 55 (1983) 1819. C. Minoia, A. Mazzucotelli, A. Cavalleri and V. Minganti, Analyst, 108 (1983) 481. C. Minoia, M. Colli and L. Pozzoli, At. Spectrosc., 2 (1981) 163. T. Shimizu, R. Kimoto, Y. Shijo and K. Sakai, Bunseki Kagaku, 30 (1981) 66. R. Escobar, M.S. Garcia-Dominguez, A. Guiraum and F.F. de la Rosa, Fresenius J. Anal. Chem., 361 (1998) 509. E. Thomsen and R.M. Stern, Scand. J. Work Environ. Health, 5 (1979) 386. Y. Suzuki, Ind. Health, 24 (1986) 23. B.D. Paul, K.K. Martin, J. Maguilo Jr. and M.L. Smith, J. Anal. Toxicol., 24 (2000) 233. Q. Gao, Lihua Jianyan Huaxue Fence, 35 (1999) 162. Y. Suzuki and K. Fukuda, J. Chromatogr. Biomed. Appl., 81 (1989) 283. R. Nusko and K.G. Heumann, Fresenius J. Anal. Chem., 357 (1997) 1050. J. Goschnick, M. Lipp and H.J. Ache, Fresenius J. Anal. Chem., 346 (1993) 365. S.B. Roychowdhury and J.A. Koropchak, Anal. Chem., 62 (1990) 484. D. Marlow, J. Wang, T.J. Wise and K. Ashley, Am. Lab., 32 (2000) 26. W. Matczak and J. Chmielnicka, Chem. Anal. (Warsaw), 34 (1989) 431. X. Wang, Fenxi Huaxue, 11 (1983) 879. T. Shoji, F.E. Huggins and G.P. Huffman, Energy Fuels, 16 (2002) 325. J.M. Boiano, M.E. Wallace, W.K. Sieber, J.H. Groff, J. Wang and K. Ashley, J. Environ. Monit., 2 (2000) 329. J. Wang, K. Ashley, D. Marlow, E.C. England and G. Carlton, Anal. Chem., 71 (1999) 1027. O. Rohling and B. Neidhart, Fresenius J. Anal. Chem., 351 (1995) 33. D.L. Ehman, V.C. Anselmo and J.M. Jenks, Spectroscopy, 3 (1988) 32. K. Sawatari, Ind. Health, 24 (1986) 111. L. Smirnova and M. Vackova, Chem. Prum., 38 (1988) 340. H.L. Sun, H.M. Liu and S.J.J. Tsai, J. Chromatogr. A, 857 (1999) 351. H. Katsumata, N. Teshima and T. Kawashima, Bull. Chem. Soc. Jpn., 70 (1997) 2151. B.A. Uzoukwu and C.I. Ukegbu, Indian J. Chem., 36A (1997) 351. A. Wasey, R.K. Bansal, M. Satake and B.K. Puri, Microchem. J., 33 (1986) 352. C. Li and X. Yu, Lihua Jianyan Huaxue Fence, 37 (2001) 43. S.K. Jain, B.K. Puri, A.L. Singla and A.L.J. Rao, Microchem. J., 37 (1988) 167. F. Goodarzi and F.E. Huggins, J. Environ. Monit., 3 (2001) 1. D.W. Huo and H.M. Kingston, Anal. Chem., 72 (2000) 5047. J. Wang, K. Ashley, E.R. Kennedy and C. Neumeister, Analyst, 122 (1997) 1037. J. Prokisch, S.A. Katz, B. Kovacs and Z. Gyori, J. Chromatogr. A, 774 (1997) 363.

Sample preparation for chromium speciation 413 414 415 416 417 418 419 420 421 422 423 424 425 426 427 428 429 430 431 432 433 434 435 436 437 438 439 440 441 442 443 444 445 446 447 448 449 450 451

Z.G. Zhang, Y.P. Mao, Y. Tang and Y.Z. Fang, Fenxi Huaxue, 24 (1996) 1109. J. Lintschinger, K. Kalcher, W. Gossler and M. Novic, Fresenius J. Anal. Chem., 351 (1995) 604. F.E. Huggins, N. Shah, J. Zhao, F. Lu and G.P. Huffman, Energy Fuels, 7 (1993) 482. A. Padarauskas and G. Schwedt, Talanta, 42 (1995) 693. K.S. Panesar, O.V. Singh and S.N. Tandon, Talanta, 41 (1994) 1341. K. Kersting, M. Adelmann and D. Breuer, Gefahrstoffe Reinhalt Luft., 59 (1999) 247. A. Bobrowskli, M. Gawlicki and J. Malolepszy, Environ. Sci. Technol., 31 (1997) 745. A. Yamada, K. Hodouchi and N. Yamazaki, Nippon Kagaku Kaishi, 5 (1991) 433. J.L. Vandenbalck and G.J. Patriarche, Sci. Total Environ., 60 (1987) 97. J.L. Vandenbalck and G.J. Patriarche, Anal. Lett., 17 (1984) 2309. A.G. Coedo, T. Dorado, I. Padilla and F.J. Alguacil, J. Anal. At. Spectrom., 15 (2000) 1564. J. Mocak, M. Vanickova and J. Labuda, Mikrochim. Acta, II (1985) 231. S.C. Stephen, Anal. Proc., 30 (1993) 440. K. Rieder, F. Baertschi, L. Muralt and U. Mueller, Mitt. Lebensmittelunters Hyg., 92 (2001) 178. J. Kowalzik, J. Poppe, M. Sietz and A. Sonnenberg, GIT Labor Fachz., 45 (2001) 60. J.X. Yu and H.H. Zeng, Fenxi Ceshi Xuebao, 17 (1998) 59. O. Nygren and J.E. Wahlberg, Analyst, 123 (1998) 935. M. Melich, S. Palagyi and M. Kern, Isotopenpraxis, 25 (1989) 354. Z. Wang, G. Xu and H. Shen, Fenxi Huaxue, 16 (1988) 193. V. Maheswari and N. Balasubramanian, Analusis, 25 (1997) 2. A.R. Timerbaev, O.P. Semenova, W. Buchberger and G.K. Bonn, Fresenius J. Anal. Chem., 354 (1996) 414. M.B. Melwanki and J. Seetharamappa, Indian J. Chem. Sect. A, 40A (2001) 659. S.L. Feng, A.N. Tang and J. Fan, Fenxi Huaxue, 29 (2001) 558. D.D. Malkhede, P.M. Dhadke and S.M. Khopkar, Indian J. Chem. Sect. A, 38 (1999) 1079. M. Theisen and R. Niessner, Fresenius J. Anal. Chem., 365 (1999) 332. E. Pappert, J. Flock and J.A.C. Broekaert, Spectrochim. Acta B, 54 (1999) 299. M. Kamburova, Mikrochim. Acta, 128 (1998) 177. Z.Y. Cui and H.Y. Ren, Lihua Jianyan Huaxue Fence, 34 (1998) 86. X.L. Liu, Y. Zhou and J.Z. Xiao, Lihua Jianyan Huaxue Fence, 34 (1998) 111. D.T. Burns and C.D.F. Dangolle, Anal. Chim. Acta, 356 (1997) 145. M. Kamburova, Anal. Lett., 30 (1997) 305. V.V. Lebedev, Zavod. Lab., 63 (1997) 55. H.Z. Li, D.T. Zhai, C.X. Yang and X.H. Jiang, Lihua Jianyan Huaxue Fence, 33 (1997) 313. W.B. Xu and C.S. Ren, Lihua Jianyan Huaxue Fence, 32 (1996) 107. X.G. Chen, Y.J. Xu and Z.D. Hu, Anal. Lett., 26 (1993) 605. X. Lu, Z.P. Ou, E.Y. Shao and J.M. Pan, Lihua Jianyan Huaxue Fence, 29 (1993) 210. S.A. Barakat, D.T. Burns and M. Harriot, Anal. Chim. Acta, 272 (1993) 135. X. Zhao, X. Liu, R. Zhang and Z. Hu, Huaxue Shiji, 10 (1988) 173. Y. Anjaneyulu, M.R.P. Reddy and C.S. Kavipurapu, Analyst, 111 (1986) 1167.

1169

M. de la Guardia and A. Morales-Rubio 452 453 454 455 456 457 458 459 460 461 462 463 464 465 466 467 468 469 470 471 472 473 474 475 476 477 478 479 480 481 482 483 484 485 486 487 488 489

1170

M. Sikovec, M. Franko, F.G. Cruz and S.A. Katz, Anal. Chim. Acta, 330 (1996) 245. M. Sikovec, F.G. Cruz, M. Franko and S.A. Katz, Spectrosc. Lett., 29 (1996) 465. K.J. Sreeram and T. Ramasami, J. Environ. Monit., 3 (2001) 526. K. Ndung’u, N.K. Djane, F. Malcus and L. Mathiasson, Analyst, 124 (1999) 1367. R. Milacic and M. Benedik, J. Pharm. Biomed. Anal., 18 (1999) 1029. K. Vercoutere, R. Cornelis, S. Dyg, L. Mees, J.M. Christensen, K. Byrialsen, B. Aaen and P. Quevauviller, Mikrochim. Acta, 123 (1996) 109. H.H. Fricke, D. Dahmann, D. Breuer, P. Muellers and B. Schleser, Staub Reinhalt Luft., 55 (1995) 275. S. Dyg, T. Anglov and J.M. Christensen, Anal. Chim. Acta, 286 (1994) 273. H.D. Buer, D. Dahmann, H.H. Frike, S. Herwald and B. Neidhart, Staub Reinhalt Luft., 51 (1991) 125. J.M. Arber, D.S. Urch and N.G. West, Analyst, 113 (1988) 779. K. Sawatari and F. Serita, Ind. Health, 24 (1986) 51. L. Olah and J. Pivoluska, J. Radioanal. Nucl. Chem., 96 (1985) 575. K. Kersting, M. Adelmann and D. Breuer, Staub Reinhalt Luft., 54 (1994) 409. S. Tian and G. Schwedt, Anal. Chim. Acta, 317 (1995) 189. C.J. Liu and W. Zhang, Fenxi Shiyanshi, 13 (1994) 48. B.R. James, J.C. Petura, R.J. Vitale and G.R. Mussoline, Environ. Sci. Technol., 29 (1995) 2377. A.A. Mohamed and M.F. El-Shahat, Anal. Sci., 16 (2000) 151. H. Katsumata, N. Teshima, M. Kurihara and T. Kawashima, Talanta, 48 (1999) 135. X.J. Chang, X.Y. Luo, Z.X. Su, G.Y. Zhan and W.Y. Gao, Fresenius J. Anal. Chem., 349 (1994) 438. N. Yoshikuni, Talanta, 38 (1991) 515. M. Soylak, F. Armagan, L. Elci and M. Dogan, Ann. Chim. (Rome), 91 (2001) 637. R.J. Vitale, G.R. Mussoline, K.A. Rinehimer, J.C. Petura and B.R. James, Environ. Sci. Technol., 31 (1997) 390. M.E. Malla, M.B. Alvarez and D.A. Batistoni, Quim. Anal., 15 (1996) 129. S.K. Sahoo and S.M. Khopkar, Indian J. Environ. Prot., 9 (1989) 241. I.S. Balogh, I.M. Maga, A. Hargitai-Toth and V. Andruch, Talanta, 53 (2000) 543. R. Milacic and J. Scancar, Analyst, 125 (2000) 1938. D.J. Hawke and A. Lloyd, Analyst, 113 (1988) 413. K. Ohls and G. Riemer, Fresenius J. Anal. Chem., 317 (1984) 774. R.E. Shaffer, J.O. Cross, S.L. Rose-Pehrsson and W.T. Elam, Anal. Chim. Acta, 442 (2001) 295. Z.L. Chen, R. Naidu and A. Subramanian, J. Chromatogr. A, 927 (2001) 219. M. Pantsar-Kallio, S.P. Reinikainen and M. Oksanen, Anal. Chim. Acta, 439 (2001) 9. J. Kalembkiewicz, S. Kopacz and E. Soco, Chem. Anal. (Warsaw), 46 (2001) 369. A. Goetzl and W. Riepe, Talanta, 53 (2001) 1187. E. Desimoni, P. Genevini, F. Tambone, S. Pikovic and P. Banﬁ, Electroanalysis, 12 (2000) 337. Y.I. Korenman, S. Kopach, Y. Kalembkievich, L. Filar and B. Papchak, J. Anal. Chem., 55 (2000) 25. M. Korolczuk, Anal. Chim. Acta, 414 (2000) 165. H. Rudel and K. Terytze, Chemosphere, 39 (1999) 697. D. Huo, Y. Lu and H.M. Kingston, Environ. Sci. Technol., 32 (1998) 3418.

Sample preparation for chromium speciation 490 491 492 493 494 495 496 497 498 499 500 501 502 503 504 505 506 507 508 509 510 511 512 513 514 515 516 517 518 519 520 521 522

M. Korolczuk and M. Grabarczyk, Chem. Anal. (Warsaw), 43 (1998) 257. L.M. Flores-Velez, M.E. Gutierrez-Ruiz, O. Reyes-Salas, S. Cram-Heydrich and A. Baeza-Reyes, Int. J. Environ. Anal. Chem., 61 (1996) 177. J.W. Grate and R.H. Taylor, Field Anal. Chem. Technol., 1 (1996) 39. X.Y. Wei, X.M. Wang, Y.H. Liu and J.X. Tan, Fenxi Huaxue, 24 (1996) 1170. A. Sahuquillo, J.F. Lopez-Sanchez, R. Rubio and G. Rauret, Mikrochim. Acta, 119 (1995) 251. P. Fodor and L. Fischer, Fresenius J. Anal. Chem., 351 (1995) 454. J. Prokisch, B. Kovacs, Z. Gyori and J. Loch, Commun. Soil Sci. Plant Anal., 26 (1995) 2051. J. Prokisch, B. Kovacs, Z. Gyori and J. Loch, J. Chromatogr. A, 683 (1994) 253. G. Solano, S.A. Katz, J. Holzbecher and A. Chatt, J. Radioanal. Nucl. Chem., 179 (1994) 173. A.R. Paniagua, M.D. Vazquez, M.L. Tascon and P. Sanchez-Batanero, Electroanalysis, 5 (1993) 155. R. Milacic, J. Stupar, N. Kozuh and J. Korosin, Analyst, 117 (1992) 125. W. Qi, Y. Zhu, S. Ding and H. Ye, Yankuang Ceshi, 10 (1991) 97. A.V. Granzhan, G.M. Kuchuk and A.K. Charykov, Zh. Anal. Khim., 46 (1991) 812. K. Furtmann and D. Seifert, Fresenius J. Anal. Chem., 338 (1990) 73. L.A. Heringer-Donmez and W.E. Kallenberger, J. Am. Leather Chem. Assoc., 84 (1989) 110. G.J. DeMenna, Chromatogr. Int., 19 (1986) 16. L. Garcia-Rico, R.E.R. Ruiz and L.G. Coronado, J. AOAC Int., 82 (1999) 575. J.F. Aguilera, C. Prieto, E. Molina and M. Lachica, Analusis, 16 (1988) 454. H.F. Zou, S.K. Xu and Z.L. Fang, At. Spectrosc., 17 (1996) 112. R.L. Keith, A.J. Gandolﬁ, L.C. McIntyre, M.D. Ashbaugh and Q. Fernando, Nucl. Instrum. Methods. Phys. Res. Sect., B149 (1999) 167. R.L. Keith, A.J. Gandolﬁ, L.C. McIntyre, M.D. Ashbaugh and Q. Fernando, Nucl. Instrum. Methods. Phys. Res. Sect. B, B130 (1997) 358. Q.H. Xu, J.S. Wang and Y.X. Xu, Lihua Jianyan Huaxue Fence, 29 (1993) 169. X. Jiang, Q. Cai, W. Shi and X. Lu, Fenxi Huaxue, 19 (1991) 1311. M.E. Soares, M. Lourdes-Bastos and M.A. Ferreira, J. Anal. At. Spectrom., 9 (1994) 1269. J.Y. Yao, Q.L. Hu and W.B. Xie, Guangpuxue Yu Guangpu Fenxi, 14 (1994) 69. M.E. Soares, M.L. Bastos and M. Ferreira, J. AOAC Int., 83 (2000) 220. L.F. Huang, Y.W. Huo and B.W. Wang, Lihua Jianyan Huaxue Fence, 35 (1999) 170. W.R. Jin, X.L. Jia and J.M. Lu, Electroanalysis, 7 (1995) 962. K. Saraswathi, K. Yamuna and K.A. Emmarnel, J. Electrochem. Soc. India, 39 (1990) 122. K. Saraswathi, K. Yamuna and K.A. Emmarnel, Bull. Electrochem., 6 (1990) 953. Y. He, M.L. Cervera, M.I. Garrido-Ecija and M. de-la-Guardia, Anal. Chim. Acta, 421 (2000) 57. S. Dyg, R. Cornelis, B. Griepink and P. Quevauviller, Anal. Chim. Acta, 288 (1994) 245. K. Vercoutere, R. Cornelis, L. Mees and P. Quevauviller, Analyst, 123 (1998) 965.

1171

This page intentionally left blank

Chapter 36

Sample preparation for metal-based drugs R.R. Barefoot

36.1

INTRODUCTION

About 25 elements are essential for animal life. These elements are absorbed through the intake of food, water, and supplementary preparations. Both essential and nonessential elements can be used in drugs for therapeutic purposes. Some inorganic compounds and metal-containing organic compounds recommended for treatments of various health problems and physical and mental conditions are classiﬁed as drugs. The objective of this section is to provide information on sample preparations for analytical work involving metal-based drugs, and speciﬁcally on speciation analyses. The prescriptions of these drugs in health care depend upon a knowledge of the ways in which drugs interact in the body and in recognizing side effects and toxicity levels. Although measurements of total elements derived from metal-based drugs in body ﬂuids and organs have been reported in many clinical studies, relatively little information is available on the detailed chemical and biological transformations which occur after the drugs have been administered. Such information is usually obtained from studies of speciation. Most of the reports on speciation studies of metal-based drugs pertain to platinum and gold compounds. Thus, most of this section is devoted to these drugs. Some reports involving ruthenium, mercury and vanadium compounds is also included.

36.2

PLATINUM-BASED DRUGS

Some of the most important metallo-drugs, namely those based on platinum compounds, are used for treatment of some forms of cancer. Of these, cisplatin, or cis-diamminedichloroplatinum(II), is the anticancer drug which has received the most attention since the discovery of its properties over 30 years ago. Over this period of time, many platinum compounds have been tested in treatment programs to assist patients suffering from cancer. Cisplatin has been the most Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

1173

R. R. Barefoot TABLE 36.1 Status of some platinum anticancer drugs Drugs

Clinical status

Cisplatin Carboplatin Oxaliplatin Lobaplatin JM216 ZD0473

Worldwide approval Worldwide approval Approved in France; phase II clinical trials Approved in China; phase II clinical trials Phase III clinical trials Phase I clinical trials

successful one and it is the platinum drug that is most widely used around the world. The platinum drugs listed in Table 36.1 have been approved or are being studied along with others in on-going trials. The current status of platinumcontaining drugs has been discussed by Wong and Giandomenico [1]. Some ruthenium compounds also are being studied as potential anti-cancer drugs. Dosages of administered drugs must be monitored carefully because of their toxicities and in order to assess their effectiveness. Side effects are of great concern in setting limits on levels which the body can tolerate. Measurements of changes in concentrations of intact drugs, and of biotransformation and degradation species, in body ﬂuids during and after administration of drugs, have been important in establishing their pharmacokinetics. Thus, sampling and sample preparation for subsequent analyses is a major concern. When cisplatin or other platinum anticancer drug is administered, a signiﬁcant portion of the drug reacts immediately with proteins and thiol compounds such as L -methionine, L -cystein and glutathione in biological ﬂuids. This portion of the drug is called bound platinum, and it does not exhibit anticancer properties. The rest is called free or ultraﬁlterable platinum; its concentration as measured in plasma ultraﬁltrate (PUF) is the most signiﬁcant fraction of the drug from a pharmacological and toxicological points of view. Studies of cisplatin have shown that the monohydrated species, cis[PtCl(NH3)2(H2O)]þ, is the most cytotoxic of the cisplatin species. Thus, speciation analyses comprise procedures for measurements of total and free platinum as well as other procedures designed to obtain more detailed information about compositions. The latter involve means of detection and identiﬁcation of intact drugs and major transformation species. Measurements of total and free platinum have been carried accomplished conveniently by ﬂame and graphite furnace atomic absorption spectrometry (GF-AAS) techniques, mainly as described by Barefoot and Van Loon [2]. High performance liquid chromatography (HPLC) has played a leading role in separations of drug species. Methods of detection and measurement in

1174

Sample preparation for metal-based drugs

HPLC studies have included atomic absorption spectrometry, UV/Vis spectrometry, inductively coupled plasma-mass spectrometry (ICP-MS) and mass spectometry (MS). Recent work in this area has been reviewed by Barefoot [3]. Nuclear magnetic resonance (NMR) spectroscopy has also been used for identiﬁcations of biotransformation and degradation products of platinum drugs. Investigations of 195Pt and 15N NMR have yielded information on changes in structures of cisplatin and carboplatin, or cis-diammine(1,1cyclobutanedicarboxylato) platinum (II) in aqueous media, as reported by Barnham et al. [4]. 36.2.1 Preparation of biological samples Most of the work reported on analyses of samples involving platinum drugs is centered on cisplatin. Samples have included whole blood, plasma, PUF, leukocytes, urine and tissues. The preparation of PUF from whole blood is very important. PUF is the most frequently analyzed type of sample for its content of free platinum, intact drugs and transformation species. Cisplatin, carboplatin and other drugs undergo rapid biotransformations. Sample preservation and treatment requires rapid working procedures, cooling the samples as much as possible and freezing materials, which are being retained for subsequent work. Details of these operations are recorded under speciﬁc drugs and in Table 36.2. TABLE 36.2 Preparation of PUF for platinum speciation analyses Compound

Procedure

Detection/method

Reference

Cisplatin

Centrifugation for 20 min at 48C and 4000g, 10,000 MW cut-off ﬁlter; storage 2708C Centrifugation for 30 min at 48C and 2000g, 25,000 MW cut-off ﬁlter; storage 2808C Centrifugation for 15 min at 48C and 2000g, 30,000 MW cut-off ﬁlter; storage 2208C Centrifugation for 45 min at 48C and 1000g, CF 25 membrane cone, SPE; storage 2208C Methanol extraction for 18 h and 2208C centrifugation for 20 min and 2000g; storage 2208C

HPLC

[7]

GF-AAS

[17]

HPLC

[18]

HPLC

[21]

HPLC

[22]

Oxaliplatin

Ormaplatin

Lobaplatin

JM216

1175

R. R. Barefoot

36.2.2 Cisplatin A typical sampling procedure is given here for the preparation of blood plasma and leukocyte fractions from whole blood. It is applicable to cisplatin, carboplatin and the other platinum drugs listed in Table 36.2. Blood samples are collected in heparinized test tubes on ice. As soon as possible, plasma is separated by centrifugation for 10 min at 48C and 1000g [5]. The layer containing blood leukocytes is removed and frozen at 2208C [6]. PlatinumDNA adducts, formed as a result of administrations of cisplatin or carboplatin, are obtained from this fraction. A puriﬁcation step is necessary in order to separate proteins from the adducts. PUF is prepared from blood plasma with minimum delay. Table 36.2 contains a selection of procedures used for preparations of PUF in studies of cisplatin, carboplatin and other drugs. The procedure for cisplatin is also used for carboplatin [6]. Precipitation of proteins from whole blood and blood plasma with ice-cold ethanol was compared with centrifugation for the preparation of plasma and PUF by Johnsson et al. [8]. A volume of 0.5 ml of whole blood or plasma was mixed with 1 ml of ice-cold ethanol (99.5%). Each sample was centrifuged for 5 min at 48C and 1000g; and then frozen at 2 708C. Platinum levels in plasmas from both procedures were similar. The authors recommended ethanol deproteinization in place of ultraﬁltration on the basis of simplicity and cost effectiveness of the procedure. The method for thawing frozen samples is also important [9]. There were no losses of cisplatin in samples thawed by immersion in water at 808C for 30 s. A reduction of 5% in cisplatin concentration occurred during thawing at room temperature for 20 min. Urine samples are collected in ice-cold test tubes. If they are not analyzed immediately, the samples are stored at 2208C [10,11]. Speciation of platinum in urine was studied by means of HPLC by Tang et al. [10]. Kidney samples from rats exposed to cisplatin are homogenized with 0.25% w/w Triton X-100 (1:4 w/v) [11] and GF-AAS was used for the determination of platinum content. For the determination of the cisplatin content in tumor tissues, the tissues are decomposed in 65% nitric acid at 378C for 2 d [12] and the solutions are subsequently analyzed for platinum by GF-AAS. The stabilities of cisplatin and its monohydrated complex in samples of whole blood, plasma and PUF at 378C and pH 7.4 were measured by Andersson and Ehrsson [13]. The half-lives of cisplatin and its complex were 1.43 and 0.36 h, respectively, in blood, and 0.88 and 0.05 h, respectively, in plasma. A small quantity of the monohydrated complex was detected when cisplatin was added to plasma under the same conditions. Free platinum levels in blood and plasma did not decrease signiﬁcantly over a period of 2 h at 208C. Gradual decreases in levels were noted in samples stored at 2208C over a 14-d period, but no decreases were detected at 2708C for

1176

Sample preparation for metal-based drugs

3 months [8]. Dilute solutions of cisplatin in 0.9% sodium chloride are stable for several days. It is important to note the formation of reaction products of cisplatin and its aquo complexes with solvents and reagents used in speciation procedures, particularly in HPLC. Such reaction products upset speciation equilibria and lead to erroneous conclusions about compositions [14]. Reagents to be avoided are acetonitrile, carboxylic acids (formic, acetic, triﬂuoroacetic), and buffers (phosphates, Tris, HEPES). Methanol and triﬂic acid (triﬂuoromethanesulfonic acid) are unreactive, as are some ion-pairing agents. 36.2.3 Carboplatin Carboplatin, or cis-diammine(1,1-cyclobutanedicarboxylato)platinum(II), is a less toxic analogue of cisplatin. Carboplatin exhibits cross-resistance with cisplatin; that is, carboplatin is active against the same types of tumors as cisplatin. Preparations of samples for speciation and determinations of carboplatin in PUF [5], leukocytes [6], urine [10,11] and kidney tissues [11], have been described in Section 36.2.2 on cisplatin. No degradation of carboplatin in either plasma or PUF was detected over periods of time typical for routine analyses. Solutions containing 3.2 mg ml21 of carboplatin in water and in 5% glucose were stable for 1 month in the dark at 48C. Decomposition of carboplatin was accelerated by exposure to light and at higher temperatures [15]. Concentrations of carboplatin ranging from 0.5 –10 mg ml21 in 0.9% sodium chloride, and of 1 mg ml21 glucose were stored at 238C. A decrease of 10% of drug in the salt solution was observed after 168 h, but only 5% loss was noted in the glucose solution [16]. 36.2.4 Oxaliplatin Oxaliplatin, or trans-L -1,2-diaminocyclohexaneoxalatoplatinum(II), exhibits no cross-resistance with cisplatin, and only mild nephrotoxicity. The latter property is one of the main limitations in cisplatin therapy. The preparation of plasma from whole blood is similar to that described in Section 36.2.2, except that samples are stored at 2 808C. A procedure for PUF preparation is found in Table 36.2 [17]. Oxaliplatin biotransformations in samples of PUF and urine were studied by HPLC-ICP-MS [18]. PUF samples are analyzed directly, and urine samples are ﬁltered and then diluted with mobile phase. In order to prepare a solution for the determination of platinum in red blood cells (RBC) by GF-AAS, 0.5 ml of RBC is digested in nitric acid in a ratio of 2:1 (volume). The digested sample is mixed with Triton X-100 [17]. In works involving oxaliplatin, free platinum levels in whole blood and PUF diminished by 80% over a period of 24 h at room temperature.

1177

R. R. Barefoot

36.2.5 Ormaplatin Ormaplatin, or trans-D ,L -1,2-diaminocyclohexanetetrachloroplatinum(IV), has shown unpredictable neurotoxicity in trials. However, in speciation studies of ormaplatin, its major biotransformation products have been compared with those of oxaliplatin by Luo et al. [19]. RBC and plasma are prepared by a centrifugation procedure similar to that described in Section 36.2.2. The preparation of PUF is found in Table 36.2 [19]. RBC cytosol and RBC cytosol ultraﬁltrate are obtained from RBC pellets. The pellets are washed, sonicated on ice and then subjected to centrifugation for 10 min, 48C and 13,000g. The same procedures are used in the preparation of comparison samples containing oxaliplatin. In other works, platinum concentrations in plasma and PUF were determined by GF-AAS [20]. Plasma is decomposed with nitric acid and 3% hydrogen peroxide, while PUF does not require additional treatment prior to analysis. 36.2.6 Lobaplatin Lobaplatin, or 1,2-diamminomethylcyclobutaneplatinum(II), is a water soluble compound which is a 1:1 mixture of two diasterioisomers. They are designated as LP-D1 ¼ RSS conﬁguration and LP-D2 ¼ SSS conﬁguration. A HPLC procedure was described by Welink et al. [21] for the determination of the two isomers in PUF. Plasma is obtained from whole blood as described in the Section 36.2.2, and PUF is prepared as shown in Table 36.2. Solid phase extraction (SPE) by means of a C18 Sep Pak cartridge is employed for the separation of lobaplatin from PUF. Prior to the application of a sample diluted 1:1 in water, the cartridge is treated with 5.0 ml methanol, 1.0 ml acetonitrile and then 5.0 ml water. Lobaplatin retained on the cartridge is washed with 1 ml water and the eluted with 1 ml methanol. The eluate is evaporated under a stream of nitrogen and the residue is dissolved in 0.3 ml water for HPLC analysis. 36.2.7 JM216 JM216, or bis(acetato)amminedichloro(cyclohexylamine)platinum(IV), is an anticancer drug that can be administered orally. Advantages of oral administration include convenience for patients and lower delivery costs than those associated with injections. A procedure for preparation of samples for speciation analyses of biotransformation products is shown in Table 36.2 [22]. This involves an extraction into methanol. An alternative procedure for the preparation of PUF from plasma requires centrifugation for 45 min at 48C and 1400g with a 10,000 MW cut-off ﬁlter. PUF is ﬂash frozen with liquid nitrogen and stored at 2 208C [23]. JM216 and several of its biotransformation products were stable when stored for 28 d at 2 208C.

1178

Sample preparation for metal-based drugs

36.2.8 Validated methods of analysis Validations of analytical methods constitute important parts of research reports. Many of the reports cited here contain such information. In addition, a number of research papers have been published concerning validations of designated methods for the determination of platinum drugs and platinum concentrations in biological samples. The reports included details of sample preparation procedures as they relate to the methods being studied. The methods themselves have applications in pharmacokinetic studies of the particular drugs. Five validation reports are summarized in Table 36.3. Blood is collected in heparinized test tubes on ice. Samples are processed within one hour in order to minimize changes in speciation of platinum compounds. Plasma is separated from whole blood at 48C by centrifugation for periods of 5 min or longer at 1500 – 3000g. PUF is prepared by centrifugation of plasma. Conditions range from 1500g to 3000g at temperatures of 4 or 158C for 10 min. Samples are stored at 2 208C. Urine samples are stored at 2 208C. Saliva samples are collected in dental cotton role [24]. The saliva is recovered by centrifugation at 1500g and then frozen (2208C). TABLE 36.3 Validation of analytical methods for platinum and ruthenium-containing drugs Drug Carboplatin

Carboplatin Cisplatin Oxaliplatin

JM216

ZD0473 NAMI-A

Sample PUF Plasma Saliva Urine PUF PUF PUF Blood Plasma PUF Plasma Urine Plasma PUF Plasma Urine

Concentration 21

0.2 –242 mmol Cp l 0.8 –242 mmol Cp l21 0.11 –3.6 mmol Cp l21 2 –12.1 mmol Cp l21 0.05–40 mg Cp ml21 0.1 –1.25 mmol Pt l21 1 –250 ng Pt ml21 0.1 –10 mg Pt ml21 0.1 –10 mg Pt ml21 5 –150 ng Pt ml21 10– 150 ng Pt ml21 50– 150 ng Pt ml21 10– 5000 ng ml21 (as ZD0473) 0.2 –44 mmol 1 2 220 mmol 1 –220 mmol (as NAMI-A)

Detection/method

Reference

GF-AAS

[24]

HPLC-UV, HPLC-PC GF-AAS USN-ICP-MS

[25] [26] [27]

GF-AAS

[28]

HPLC-MS

[29]

GF-AAS

[30]

Cp ¼ carboplatin; PC ¼ postcolumn derivatization; USN ¼ ultrasonic nebulization.

1179

R. R. Barefoot

The determination of carboplatin in PUF by means of the HPLC-UV method described by Burns and Embree [25] required an SPE procedure to separate interferences which absorbed in the same spectral region as carboplatin. A Supelclean amino extraction cartridge (3 ml) is conditioned with 2 ml of acetonitrile –water (95:5) and a 200 ml sample diluted with 2 ml of acetonitrile is pulled through the cartridge. The cartridge is washed with 2 ml of acetonitrile –water (90:10). Carboplatin retained on the cartridge is eluted in 2 ml of acetonitrile –methanol –water (50:25:25). The sample is evaporated under nitrogen and then reconstituted in water for HPLC analysis. GF-AAS was used for the determination of platinum contents of biological samples obtained from cancer patients treated with JM216 [28]. A matrix modiﬁer, 5% Triton X-100, is applied to the platform prior to the introduction of a sample of plasma. PUF samples are preconcentrated on the platform prior to ashing, no modiﬁer is necessary. Urine is diluted with 10% hydrochloric acid. The drug ZD0473, or cis-amminedichloro(2-methylpyridine)platinum(II), is a relatively new antitumor agent undergoing tests. ZD0473 has properties that overcome drug resistance induced by cisplatin. The dichloro form of the drug exists in equilibrium with two aquated forms in plasma. A validated method [29] was developed for the conversion of the aquated species back to the dichloro form of the parent drug. A single molecular species in plasma is analyzed by MS. Components of a 50 ml sample are separated by HPLC prior to MS determination. Ruthenium complexes are being studied as possible antitumor drugs. NAMI-A, or ImH[trans-RuCl4(DMSO)Im], (Im ¼ imidazole, DMSO ¼ dimethyl sulfoxide) is a new drug which is receiving clinical attention as an alternative to cisplatin for treatment of several types of tumors. A rapid, sensitive method was developed and validated [30] for the determination of NAMI-A in plasma, PUF and urine by GF-AAS. Samples are diluted with an appropriate hydrochloric acid buffer solution.

36.3

GOLD-BASED DRUGS

Several gold-containing compounds have been employed for treating rheumatoid arthritis (chrysotherapy). These compounds are known as antiarthritic drugs. Products in current use include (1) auranoﬁn or triethylphosphine gold(I) tetraacetylthioglucose; (2) myochrysine or sodium gold(I) thiomalate; and (3) solganol or gold(I) thioglucose. Information on the modes of action of the drugs has been obtained by means of measurements of concentrations of total gold, and of intact species and metabolites in body ﬂuids. Sensitive analytical techniques, such as GF-AAS and ICP-MS, have been used for determination of total gold concentrations [3]. Separations of intact drugs and their metabolites have been accomplished by HPLC. Both ICP-MS and AAS as well as formation of

1180

Sample preparation for metal-based drugs

postcolumn derivatives have provided the means of detection of gold compounds eluted from HPLC columns. Table 36.4 contains a summary of applications. 36.3.1 Sample preparation Procedures for collection and preparation of samples involving gold-based drugs are similar to those described for cisplatin in Section 36.2.2. It is important to collect samples in cold test tubes. They are either analyzed immediately or stored at 2208C. 36.4

MERCURY

Some mercury compounds possess antimicrobial properties. One such compound, thimersal, is used in contact lens soaking and cleaning solutions and in vaccines. Thimersal is analyzed in the presence of mercury(II) chloride by HPLC coupled with cold vapor ICP-MS [36]. 36.5

VANADIUM

A number of vanadium compounds appear to have therapeutic effects. Insulinlike effects of vanadium have been observed. This has led to investigations of the relationship between vanadium and diabetes mellitus. Vanadium mimics most of the biological effects of insulin in various types of cells. Among vanadium compounds, VO(malolate)2, VO(picolinate)2, and VO(6-Me-picolinate)2 show potency and high efﬁciency in decreasing blood glucose levels. Biospeciations of these compounds in blood serum were assessed [37], and speciation curves were calculated using stability constants taken from the literature. Sample preparations involved additions of vanadium compounds to serum. TABLE 36.4 Determination of gold species Compounds

Sample

Detection/method

Reference

Auranoﬁn Myochrysine Metabolites Auranoﬁn Auranoﬁn Myochrysine Metabolites

Urine Urine Urine Urine Blood Plasma Plasma

HPLC; ICP-MS

[31,32] [31] [31] [32] [34] [35]

HPLC-PC ICP-MS HPLC; AAS

PC ¼ postcolumn derivatization.

1181

R. R. Barefoot

36.6

LEAD

There have been many studies on the effects of lead in occupational and environmental medicine. The direct bonding of lead to proteins is important in an understanding of the toxic effects of this element. Protein-bound lead in human erythrocytes has been studied [37]. Red blood cells are separated from plasma by centrifugation. An aliquot containing 1 g of cells is frozen and thawed three times in order to lyse the cells. The cells are then dissolved in 9 g of a 0.05 mol l21 solution of ammonium hydrogen carbonate. The lead compounds are analyzed by means of gel permeation chromatography-ICE-MS. Three chromatographic peaks were linked to the presence of lead containing compounds.

36.7

CONCLUSIONS

A knowledge of speciation of metals in samples obtained in clinical studies of metal-containing drugs is important in understanding the activities of drugs in treatment of patients. Proper preparation and preservation of samples such as blood plasma, PUF, erythrocytes and urine are necessary in maintaining the species present in their original concentrations. Measurements of concentrations of intact drugs, degradation and biotransformation products, and other selected reaction products are very useful. Most of the reports of investigations in this ﬁeld are concerned with drugs based on platinum and gold. As laboratory equipment such as chromatographs and ICP-MS and MS instruments become available in more laboratories, the number of studies applied to metal-containing drugs will increase.

REFERENCES 1 2 3 4 5

6 7 8

1182

E. Wong and C.M. Giandomenico, Chem. Rev., 99 (1999) 2451. R.R. Barefoot and J.C. Van Loon, Anal. Chim. Acta., 334 (1996) 5. R.R. Barefoot, J. Chromatogr. B, 751 (2001) 205. K.J. Barnham, U. Frey, P. del S. Murdoch, J.D. Ranford and P.J. Sadler, J. Am. Chem. Soc., 116 (1994) 11175. N. Nagai, M. Kinoshita, H. Ogata, D. Tsujino, Y. Wada, K. Someya, T. Ohno, K. Masuhara, Y. Tanaka, K. Kato, H. Nagai, A. Yokoyama and Y. Kurita, Cancer Chemother. Pharmacol., 39 (1996) 131. A. Bonetti, P. Apostoli, M. Zaninelli, F. Pavanel, M. Colombatti, G.L. Cetto, T. Franceschi, L. Sperotto and R. Leone, Clin. Cancer Res., 2 (1996) 1829. A. Andersson, J. Fagerberg, R. Lewensohn and H. Ehrsson, J. Pharm. Sci., 85 (1996) 824. A. Johnsson, H. Bjork, A. Schutz and T. Skarby, Cancer Chemother. Pharmacol., 41 (1998) 248.

Sample preparation for metal-based drugs 9

10 11 12 13 14 15 16 17

18 19 20

21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37

M. Kinoshita, N. Yoshimura, H. Ogata, D. Tsujino, T. Takahashi, S. Takahashi, Y. Wada, K. Someya, T. Ohno, K. Masuhara and Y. Tanaka, J. Chromatogr., 529 (1990) 462. X. Tang, J.W. Hayes II, L. Schroder, W. Cacini, J. Dorsey, R.C. Elder and K. Tepperman, Met. Based Drugs, 4 (1997) 97. N. Nagai and H. Ogata, Cancer Chemother. Pharmacol., 40 (1997) 11. R. Milacic, M. Cemazar and G. Sersa, J. Pharm. Biomed. Anal., 16 (1997) 343. A. Andersson and H. Ehrsson, J. Pharm. Biomed. Anal., 13 (1995) 639. M. El-Khateeb, T.G. Appleton, B.G. Charles and L.R. Gahan, J. Pharm. Sci., 88 (1999) 319. M. Pujol, V. Girona, J. Prat, M. Munoz and J. De Bolos, Int. J. Pharm., 146 (1997) 263. R. Gust and B. Schnurr, Montash. Chem., 130 (1999) 637. C. Massari, S. Brienza, M. Rotarski, J. Gastiaburu, J.-L. Misset, D. Cupissol, E. Alafaci, H. Dutertre-Catella and G. Bastian, Cancer Chemother. Pharmacol., 45 (2000) 157. P. Allain, O. Heudi, A. Cailleux, A. Le Bouil, F. Larra, M. Boisdron-Celle and E. Gamelin, Drug Metab. Dispos., 28 (2000) 1379. F.R. Luo, S.D. Wyrick and S.G. Chaney, Cancer Chemother. Pharmacol., 44 (1999) 19. W.D. Figg, M.C. Christian, R. Lush, C.J. Link, P. Davis, E. Kohn, G. Sarosy, M.L. Rothenberg, R.B. Weiss, N. Ryan, J. Jacobs and E. Reed, Biopharm. Drug Dispos., 18 (1997) 347. J. Welink, B. Pechstein and W.J.F. van der Vijgh, J. Chromatogr. B, 675 (1996) 107. P. Galetis, J.L. Carr, J.W. Paxton and M.J. McKeage, J. Anal. At. Spectrom., 14 (1999) 953. W.R.L. Cairns, L. Ebdon and S.J. Hill, Fresenius J. Anal. Chem., 355 (1996) 202. L.J.C. Van Warmerdam, O. Van Tellingen, R.A.A. Maes and J.H. Beijnen, Fresenius J. Anal. Chem., 351 (1995) 377. R.B. Burns and L. Embree, J. Chromatogr. B, 744 (2000) 367. J.M.M. Terwogt, M.M. Tibben, H. Welbank, J.H.M. Schellens and J.H. Beijnen, Fresenius J. Anal. Chem., 366 (2000) 298. J.G. Morrison, P. White, S. McDougall, J.W. Firth, S.G. Woolfrey, M.A. Graham and D. Greenslade, J. Pharm. Biomed. Anal., 24 (2000) 1. S. Vouillamoz-Lorenz, J. Bauer, F. Lejeune and L.A. Descosterd, J. Pharm. Biomed. Anal., 25 (2001) 465. T. Oe, Y. Tian, P.J. O’Dwyer, D.W. Roberts, M.D. Malone, C.J. Bailey and I.A. Blair, Anal. Chem., 74 (2002) 591. M. Crul, H.J.G.D. van den Bongard, M.M. Tibben, O. van Telligen, G. Sava, J.H.M. Schellens and J.H. Beijnen, Fresenius J. Anal. Chem., 369 (2001) 442. Z. Zhao, W.B. Jones, K. Tepperman, J.G. Dorsey and R.C. Elder, J. Pharm. Biomed. Anal., 10 (1992) 279. R.C. Elder, K. Tepperman, M.L. Tarver, S. Matz, W.B. Jones and E.V. Heiss, J. Liq. Chromatogr., 13 (1990) 1191. S.G. Matz, R.C. Elder and K. Tepperman, J. Anal. At. Spectrom., 4 (1989) 767. M.H. Rayner, M. Grootveld and P.J. Sadler, Int. J. Clin. Pharmacol. Res., 9 (1989) 377. D.S. Bushee, Analyst, 114 (1989) 1167. T. Kiss, E. Kiss, E. Garribba and H. Sakurai, J. Inorg. Biochem., 80 (2000) 65. I.A. Bergdahl, A. Schutz and A. Grubb, J. Anal. At. Spectrom., 11 (1996) 735.

1183

This page intentionally left blank

Chapter 37

Sample preparation for speciation analysis for metallobiomolecules Joanna Szpunar, Brice Bouyssiere and Ryszard Lobinski

37.1

INTRODUCTION

Classical speciation analysis has targeted well-deﬁned analytes, usually anthropogenic organometallic compounds and the products of their environmental degradation, such as methylmercury, alkyllead, butyl- and phenyltin compounds and simple organoarsenic and organoselenium species [1]. Calibration standards were either available or could be readily synthesized. The presence of a metal(loid) –carbon covalent bond assured a reasonable stability of the analyte(s) during sample preparation. The volatility of the species allowed the use of gas chromatography with its inherent advantages, such as the high separation efﬁciency and the absence of the condensed mobile phase that enabled a sensitive (down to the femtogram levels) element-speciﬁc detection by atomic spectroscopy [2]. A totally different situation is faced by the analyst interested in endogenous metal species in biological systems [3– 7]. Millions of years of evolution have resulted in a great variety of biological ligands with a signiﬁcant coordinating potential for trace elements. They include small organic ligands (e.g. citrate, tartrate, oxalate, or phytate, aminoacids, oligopeptides), macrocyclic chelating molecules and macromolecules, such as proteins, DNA restriction fragments or polysaccharides. The complexity and the usually poor understanding of the system (the majority of trace element species with biological ligands have not yet been discovered!) often make even the deﬁnition of the target analyte problematic. The generally poor volatility of the metal coordination complexes with biological ligands by comparison with organometallic species calls for separation techniques with a condensed mobile phase that negatively affects the separation efﬁciency and the detection limits. The combination of a chromatographic separation technique, which ensures that the analyte compound leaves the column separately from other species of the analyte element, with atomic spectrometry, permitting a sensitive and speciﬁc detection of the target element, has become a fundamental tool for Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

1185

J. Szpunar et al.

speciation analysis, as discussed in many review publications [1,6,8 –11]. The recent impressive progress towards lower detection limits in ICP-MS, towards higher resolution in separation techniques, especially capillary electrophoresis and electrochromatography and towards higher sensitivity in electrospray mass spectrometry for molecule-speciﬁc detection at trace levels in complex matrices allows new frontiers to be crossed. More often the limiting factor to a valid speciation analysis is an adequate sample preparation that should allow the quantitative recovery of an intact analyte species from the sample, taking care of avoiding contamination. This chapter overviews methods for sample preparation for the three major categories of metal(metalloid) compounds of biological signiﬁcance. They include: (i) products of the biological metabolisms of arsenic and selenium resulting in the formation of a covalent bond between the metalloid and the carbon atom, (ii) metal complexes with endogenous biomolecules (e.g. peptides, proteins, carbohydrates) and (iii) metabolites of metal probes, e.g. metallodrugs. The discussion of the principles of the common sample preparation techniques used in bioinorganic speciation analysis (homogenization and analyte recovery from solid samples, analyte puriﬁcation by ultra- and gel ﬁltration and multidimensional chromatography and freeze-drying preconcentration) is followed by a critical review of sample preparation methods applied for speciation studies. 37.2

ELEMENTAL SPECIES IN BIOLOGICAL SYSTEMS: METALLOBIOMOLECULES

Figure 37.1 overviews trace elements with an identiﬁed role in biological systems and their molecular species. Some metals have the notoriety of showing either acute (e.g. Hg) or chronic (e.g. Pb) toxicity, whereas others (e.g. Mo, Mn, Fe, Co, Cu, Zn), referred to as essential, are needed for the accomplishment of life processes [12]. Some elements (e.g. V, Cr, Ni) are recognized as beneﬁcial to life although the borderline between being essential or beneﬁcial is vague. A number of elements show a dual character: they are essential in one oxidation state, e.g. Cr(III) or Se(IV), and yet toxic in the other state, e.g. Cr(VI) or Se(VI). Some elements, e.g. cobalt, can be considered essential only if present in a particular organic form, e.g. as cyanocobalamin (vitamin B12) and are toxic when in other forms. On the other hand, arsenic, a notoriously toxic element, becomes harmless when present in the form of arsenobetaine. A separate class consists of the metals used as pharmacological probes. Platinum (cisplatin, carboplatin), Ru3þ ( fac-RuCl3(NH3)3) compounds are used in cancer therapy whereas some Au compounds (aurithiomalate, aurothioglucose) are important antiarthritic drugs [13]. A wide range of Tc compounds (e.g. Tc-labelled antibodies, Tc-mercaptoacetyl glycine complex) are used for diagnostic imaging of renal, cardiac and cerebral functions and various forms of cancer [14].

1186

Sample preparation for speciation analysis for metallobiomolecules

Fig. 37.1. Trace elements with an identiﬁed role in biological systems and their most common species.

The occurrence of a free metal ion, especially of a transition element, in a biological cell rich in ligands with a signiﬁcant coordinating potential is highly improbable. Metalloids (As, Se) are known to be metabolized by living organisms in a way that leads to the formation of a covalent bond between the heteroatom and the carbon atom and to the biosynthesis of larger biomolecules (e.g. arsenosugars, selenoproteins). Metals are usually present in the form of coordination complexes, of which some, e.g. cyanocobalamine, are remarkably stable. Metal complexation by proteins via nitrogen or oxygen confers the activity to several enzymes, whereas the coordination via a sulfur atom is usually associated with the detoxiﬁcation of heavy metals. Metals that activate an enzyme, e.g. nicotianamine synthase (Fe, Ni) or phytochelatin synthase (Cd), are usually found to be complexed by the metabolite, nicotianamine or phytochelatin, respectively. The quantity of the synthesized metabolite regulates the concentration of the element available for the enzymatic reaction. Relatively little is known about the relevance of metal coordination to lipids and carbohydrates, although the potentially negatively charged oxygen functions and polyhydroxy groups can bind cations electrostatically and by chelation, respectively. A number of organic acids such as citric acid, oxalic acid or succinic acid as well as some amino acids and members of the mugineic acid family occur in plant tissue and are possible ligands for metal complexation

1187

J. Szpunar et al.

[15]. Complexation of divalent cations by the carboxylic acid groups of uronic acids from plant cell walls is well established. The thermodynamic stability of the metal complexes strongly depends on their equilibrium constants, thus not each and every possible complex can be isolated by chromatography. The analytical challenges faced in the area of metal probes include both the identiﬁcation of the products of metallodrug metabolism and the understanding of the binding of metallodrugs to transport proteins and DNA fragments [16]. Arsenic and selenium are two metalloids with a very rich biochemistry. As a consequence, a number of species containing As–C and Se–C bonds are naturally formed by living organisms claiming for species-speciﬁc analytical methods. 37.3

TAILORING SAMPLE PREPARATION: DEFINITION OF THE ANALYTE MOIETY

The deﬁnition of a target species (moiety) in the speciation analysis for biomolecules is often problematic because of the complexity of real-world systems. For example, speciation of mercury in biological tissues is usually understood as the differentiation between CH3Hgþ and Hg2þ. In reality, however, these species hardly exist as ions. Both methylmercury and Hg2þ can form covalent compounds with a counterion, coordination complexes with –SH aminoacids, and bind strongly to cystein residues of metallothioneins or larger proteins. In turn, metallothioneins can polymerize to form dimers and higher oligomers each of which can bind (methyl)mercury. This is why the analytical target should be deﬁned a priori in terms of the needs, the latter usually resulting from eco- or clinical toxicology. The target species should also be considered in terms of thermodynamic stability and kinetic inertness. The time scale of the analytical method should match the lifetime of the target. Although metal ions can interact with many organic ligands, labile coordination compounds cannot be separated or analysed directly. In this case, calculation based on equilibrium constants is sometimes the only feasible approach to the identiﬁcation of the species. A change in the ambient physicochemical conditions often disturbs the existing acid –base, redox and complexation equilibria in such a way that the species ﬁnally determined may not reﬂect those that had originally existed in the sample. These are important considerations that should be addressed during the sample preparation procedure.

37.4

HOMOGENIZATION AND ANALYTE RECOVERY USING UNREACTIVE BUFFERS

Plant tissues are either simply cut in pieces and extracted as fresh tissues, frozen in liquid nitrogen and ground, or freeze-dried and ground prior to leaching with a cell lysis buffer.

1188

Sample preparation for speciation analysis for metallobiomolecules

Mammalian tissues are divided into two groups: soft tissues (e.g. liver) and hard tissues (e.g. muscle) and routinely the types of homogenizer used to disrupt these tissues are liquid shear (Potter-Elvehjem or Dounce) or mechanical shear (e.g. Polytron), respectively. Whatever technique is used, it is good practice to facilitate the homogenization by an initial coarse mincing of the tissue with scissors, scalpels or (for large masses of tissue) a mincer. The methods require no addition of reagents but ﬁne cellular debris can be formed if mechanical lysis is too vigorous. Hard tissues may be rendered susceptible to liquid shear homogenization by treatment with hydrolytic enzymes at the expense of rendering the matrix more complex but an advantage to isolate the tissue fraction containing the species of interest. Vascular tissues, such as rat liver, may require some form of perfusion to remove blood prior to homogenization. Perfusion can be accomplished by a trained and licensed operator by injection of buffered saline or homogenization medium through the portal vein after cutting the blood vessels above the liver. In the most common procedures, soft tissues and cultured cells are homogenized in 0.25 M sucrose and buffered with low concentrations of an organic buffer such as TRIS, HEPES or Tricine at a pH between 7.4 and 8.6. The use of buffers at the physiological pH is required in order to avoid dissociation of the complexes. Typical components of a homogenization mixture include: b-mercaptoethanol (antioxidant), NaN3 (an antibacterial agent) and phenylmethanesulfonylﬂuoride (proteases inhibitor). Often, 1 mM EDTA is included to reduce aggregation. Brain tissues are frequently disrupted in 0.32 M sucrose rather than 0.25 M. Hypoosmotic media (e.g. 10 mM Tris–HCl, pH 7.5 or 5 mM EDTA, pH 7.4) were often used. Media selected for muscle homogenization were quite variable; KCl was often included (up to 180 mM) to solubilize some of the protein and prevent the formation of gels. After coarse mincing of muscle tissue, it is commonly softened by incubating with Nagarse at 5–50 mg per 100 ml at 48C for about 5 min. Dilution factors (proportion of tissue to the homogenizing buffer) up to 10 have been used, whereas for those with endogenous non-induced levels of biomolecules equal amounts of tissue and buffer were found suitable. The homogenization step is followed by centrifugation. The use of a refrigerated ultracentrifuge (100,000g) is strongly recommended. As a result, two fractions: a soluble one (cell supernatant, cytosol) and a particulate one (cell membranes and organelles) are obtained. Only the supernatant is usually analysed for biomacromolecules thus limiting the number of species of concern to those being cytosoluble. It is recommended that the supernatant be stored at 2208C under nitrogen prior to analysis. Filtration of the cytosol (0.2 mm ﬁlter) before introducing it onto the chromatographic column is strongly advised. A guard column should be inserted to protect the analytical column, particularly from effects of lipids, that otherwise degrades the separation. The drawback of leaching procedures using the simple aqueous buffers is their generally poor recovery of element species. The typical efﬁciencies are

1189

J. Szpunar et al.

about 10 –20% with the highest values reported of 50 –85 and 30 –57% in the case of protein-bound Zn and Cd, respectively. The low yields of the aqueous leaching procedure for some species and samples promoted more aggressive leaching media to be used by some workers. The increase in this recovery can be achieved by destruction (at least partly) of the sample matrix while the preservation of the original identity is ensured. A trade-off is always necessary between the recovery of the element from a solid matrix and the preservation of the original elemental species. The two major categories include the use of protein denaturating reagents, such as sodium dodecyl sulfonate (SDS) or 3[(3-cholamidopropyl)dimethylammonio]propanesulfonic acid (CHAPS), and the use of enzymes. 37.5

ANALYTE RECOVERY THROUGH PARTIAL DEGRADATION OF SAMPLE MATRIX

37.5.1 Extraction with protein denaturating reagents The use of SDS or CHAPS is particularly recommended in the case when the element is incorporated or strongly complexed by the moiety of interest. The most widely investigated example is the recovery of the selenoprotein fraction from yeast and animal tissues. The addition of SDS to a leaching mixture increases the yield of Se by releasing selenoaminoacids bound in selenoproteins [17]. The use of concentrated surfactants and dithiotreitol is important for the solubilization of high-molecular weight proteins and metalloenzymes [18]. 37.5.2 Enzymatic extraction of organometallic compounds The recovery of selenoaminoacids can be increased to above 95% by degrading the species originally present with a mixture of proteolytic enzymes [19]. The enzymes used in most digestion protocols are protease [19], proteinase K [20,21] or a mixture of proteolytic enzymes [17]. The use of protease XIV and proteinase K provided extraction yields of ca. 70% in plant tissues [22]. The addition of cellulase to proteinase K was found not to offer any advantages, which was attributed to the fact that the initial liquid nitrogen treatment destroyed the cellulose walls of the tissue [22]. Proteinase K extraction is considered to be adequate only when the species are not very labile. Indeed, decomposition of Se-methylselenocysteine during extraction from Arabidopsis thaliana leaves was observed [21]. Different enzymatic protocols were compared to elucidate the possible role of the cell wall digesting enzymes, such as Lysing enzyme and Driselase, for the improvement of extraction efﬁciency of selenium in edible mushrooms with and without inhibiting proteolysis during cell wall digestion [23]. A three-step procedure applying Lysing enzyme and pronase gave the highest extraction

1190

Sample preparation for speciation analysis for metallobiomolecules

efﬁciency (89%) [23]. A sequential sample preparation process was described for the same Se-enriched edible mushroom (Agaricus bisporus) after comparing ﬁve different sample extraction methods. The most efﬁcient was a three-step protocol using water extraction and two proteolytic enzymes (pepsin and trypsin) with an extraction efﬁciency of 75% [24]. The hydrolysis, carried out by boiling the sample with 4 M methanesulfonic acid under reﬂux for 8 h, was claimed to offer higher recovery of selenomethionine in comparison with proteinase K –protease XIV procedure [25]. In the analysis of enzymatic digests, the recoveries from the column were not complete (40–90%), which was attributed to the presence of strongly hydrophobic peptides resulting for the incomplete decomposition of proteins to amino acids [20,26,27]. Enzymatic procedures for the recovery of organoarsenic species were also reported. A two-step procedure for extraction of arsenic from freeze-dried apples using overnight treatment with a-amylase enzyme followed by sonication for 6 h with 40% acetonitrile was found to provide good extraction efﬁciency [28].

37.5.3 Controlled enzymatic degradation prior to speciation of metal complexes In the case of metal complexes, alkaline or acid hydrolysis cannot be accepted because it would affect the complexation equilibria. Selective enzymatic degradation of the matrix at pH natural for the sample investigated is often the only possibility. Cellulose and complex water-insoluble pectic polysaccharides are the main matrix of the water-insoluble residue after centrifugation of fruit and vegetable homogenates. The use of pectinolytic enzymes is therefore necessary to solubilize the solid sample. Pectinolysis is known to efﬁciently degrade large pectic polysaccharides but some of them, e.g. rhamnogalacturonan-II, are considered enzyme resistant [29]. A mixture of commercial preparations: Rapidase LIQe and Pectinex Ultra-SPLe was reported for release of metal complexes from the solid parts of edible plant, fruits and vegetables [29]. Water-soluble polysaccharide species with higher molecular weights can be readily decomposed by enzymic hydrolysis with a mixture of pectinase and hemi-cellulase to release the dRG-II complex [29]. The same mixture was found efﬁcient to extract the dRG-II–metal complexes from water-insoluble residue of vegetables owing to the destruction of the pectic structure [29]. Figure 37.2 shows two size-exclusion chromatograms demonstrating the release of a Pb–rhamnogalacturonan moiety from a larger molecular ediﬁce by action of a mixture of pectinolytic enzymes.

1191

J. Szpunar et al.

Pb-dRG-II complex

Intensity, cps

0.18 kDa

5.8 kDa

12.2 kDa

1.00

23.7 kDa

48.0 kDa

1.25 105

0.75

0.50 enzymatic digestion 0.25

hot water extraction

0 0

5

10 15 Elution time, min

20

25

Fig. 37.2. HPLC-ICP-MS chromatograms of apple juice before and after enzymatic treatment showing the liberation of a Pb– dRG-II complex which itself survives the enzymatic digestion.

37.5.4 Sequential enzymatic extractions for the evaluation of the bioaccessibility of metals in foodstuffs Some attention has been paid to the analysis of enzymic digests of foodstuffs in the quest for molecular information to contribute to the knowledge on the bioavailability of some elements. Enzymolysis in simulated gastric and gastrointestinal juice was proposed for meat samples [30] and for estimation of bioavailability of Pb and Cd species from cocoa [31,32]. The soluble fraction of the stomach and upper intestinal contents of a guinea pig on different diets were investigated for the species of Al, Cu, Zn, Mn, Sr and Rb. The effect of citrate on each of these elements was also assessed [32]. An enzymic digest of bovine thyroglobulin was analysed for iodine species [33]. Figure 37.3 shows the principle of an in vitro model simulating enzymatic activity in the gastrointestinal tract of monogastric species developed for the assessment of the potential bioavailability of Cd and Pb in cocoa powder and liquor of different geographical origins. The model was based on the sequential extraction with simulated gastric and intestinal juices [34]; the residue after the latter extraction was further investigated by using, in parallel, solutions of phytase and cellulase [34].

1192

Sample preparation for speciation analysis for metallobiomolecules

Sample in powder

+ Gastric juice

+ Gastric juice

(37 C, 4h)

(37 C, 4h)

+ Intestinal juice (37 C, 4h)

Insoluble residue Centrifugation + Phytase

Centrifugation Supernatant

Supernatant

Supernatant

1) 55 C, 6 h 2) 37 C, 16h

+ Cellulase (37 C, 16h)

Centrifugation Supernatant

Centrifugation

Total Analysis Fig. 37.3. Scheme for sequential enzymolysis protocol used for the evaluation of the bioaccessibility of Cd and Pb in cocoa.

37.6

FRACTIONATION OF METAL SPECIES ACCORDING TO THE MOLECULAR WEIGHT PRIOR TO ANALYTICAL CHROMATOGRAPHY OR CAPILLARY ELECTROPHORESIS

The complexity of biological matrices often makes it necessary to isolate the molecular mass fraction containing the species of interest in order to separate it from matrix components such as high-molecular weight polysaccharides or inorganic salts that would otherwise foul the analytical column used in the subsequent analysis. The simpliﬁcation of the matrix often allows the preconcentration of the analytes in the isolated fraction by means of lyophilization [35– 40]. The principle of lyophilization is to “dry” (remove water or solvents) material through the process of sublimation. Care should be taken to avoid possible losses of volatile compounds or species degrading to volatile species [41]. The two most common techniques for the fractionation of extracts of biological tissues include ultraﬁltration and gel ﬁltration.

1193

J. Szpunar et al.

37.6.1 Ultraﬁltration Ultraﬁltration is a membrane separation technique used to fractionate substances according to molecular weight and size. It is ideally suited to separate salts and other low-molecular weight solutes from high-molecular weight species. Unlike dialysis, ultraﬁltration is based on a pressure differential across the semipermeable membrane to drive permeable materials through the membrane. The centrifugal force is used to create a pressure differential on the sample to facilitate the separation process. For this reason, ultraﬁltration separates solutes and concentrates retained materials more rapidly. Membranes used in molecular ﬁltration have pore diameters ranging from 1 to ˚ and usually separate particles ranging from 100 to 106 Da. Particles with 1000 A molecular weight or size less than the membrane molecular weight cut-off pass through the membrane and emerge as permeate. Solutes with greater molecular weight or size are retained by the membrane as retentate and are concentrated during the molecular ﬁltration process. Ultraﬁltration offers a fast separation and minimal denaturation of molecules compared to precipitation methods. The most frequently used is ﬁltration using 10 kDa cut-off ﬁlters. It was used prior to the determination of free selenoaminoacids [42] and metal complexes [43] in milk, Al-complexes in blood plasma [44] and metallodrug metabolites in blood plasma prior to RP HPLC with ICP-MS [45–48] or ESI-MS [49]. Note that some smaller molecules such as metallothioneins (MTs) and phytochelatins (PCs) do not pass this cut-off because their apparent molecular masses are higher than the theoretically calculated ones. Successive ultraﬁltration through membranes with molecular weight cut-offs of 500, 5000 and 30,000 was widely used to study the distribution of metal species as a function of molecular weight is recommended for the simpliﬁcation of the matrix loaded on an HPLC column [50–52]. 37.6.2 Gel ﬁltration A ﬁner resolution can be obtained by using low-pressure size-exclusion chromatography (SEC). The latter was routinely used in earlier studies of speciation of metal complexes prior to graphite furnace AAS [53 –55]. In order to avoid the degraded chromatographic resolution, reduction of the column lifetime and interference with ES-MS when the co-eluted species are preconcentrated by freeze-drying of the fractions, a sample clean-up step by gel-permeation chromatography was introduced prior to fractionation of arsenic compounds by anion exchange (AE) [56]. It allowed the elimination of high molecular mass biopolymers (polysaccharides, proteins) and other compounds that might be adsorbed on the chromatographic stationary phase or co-elute with arsenic compounds during speciation analysis of edible algae. The recent advance in the size-exclusion gel manufacturing technology allows a relatively high peak capacity of the columns. Figure 37.4 shows the fractionation of selenium species using a Superdex Peptide gel (exclusion

1194

Sample preparation for speciation analysis for metallobiomolecules

500

Intensity

104, cps

400

300

200

100

60

80

100 120 Number of fraction

140

160

Fig. 37.4. Fractionation of selenium species by high-resolution SEC.

volume of 10 kDa) that allows the recognition of 7–8 organoselenium fractions in a selenized-yeast hot water extract. It should be noted that other than size-exclusion mechanisms often play a role in the fractionation process and the calibration of the column with the molecular weight markers may be problematic. 37.7

MULTIDIMENSIONAL LC CLEAN-UP PROCEDURES PRIOR TO CHARACTERIZATION OF METAL SPECIES BY ELECTROSPRAY MS

The non-availability of retention time standards for most of metallobiomolecules makes electrospray MS an important technique for the bioinorganic speciation analysis. However, electrospray ionization suffers from the signal suppression effect by the matrix and the generally poor compatibility of the chromatographic mobile phase with the ionization conditions. The chromatographic clean-up protocol to be developed should (i) assure the isolation of each of the arsenic species present in the sample in chromatographically pure form (one species in each fraction) and (ii) achieve the maximum simplicity of the matrix in which an arsenic species would have been isolated. It is important especially for organoarsenic species because arsenic is monoisotopic and the attribution of peaks in an MS spectrum to arsenic compounds may not be straightforward. The achievement of the second objective implies the choice of a

1195

J. Szpunar et al.

plasmaspiked with a metallodrug metal bound to plasma proteins

free-circulating metal

Metal reversibly bound Pharmacologically active!!! Metal irreversibly bound

metabolites reversibly bound

metallodrug reversibly bound

metallodrug free-circulating

metabolites free-circulating

ultrafiltration

deproteinisation

Fig. 37.5. Possible fates of a metallodrug in blood plasma after administration.

ready-to-be-removed (volatile) mobile phase buffer and an efﬁcient elimination matrix components by sequentially applying chromatography with orthogonal (different principle) separation mechanisms. For two-dimensional chromatography, it is convenient that the ﬁrst separation to be employed can handle a larger quantity of sample to provide sufﬁcient analyte quantities in individual fractions collected for further chromatography. Figure 37.5 demonstrates the need for multidimensional chromatographic puriﬁcation of organoarsenic species prior to ES-MS analysis using the example of the identiﬁcation of an arsinoylriboside (arsenosugar B). A puriﬁcation solely by SEC is insufﬁcient to assure a detectable signal at the m/z 329 where the arsenosugar is expected. The combination of SEC with anion exchange allows the identiﬁcation of a small peak at the m/z 329 that gives a correct collision-induced dissociation fragmentation pattern (not shown). However, in order to attribute this particular peak to the arsenic compound of interest, a prior knowledge on its molecular mass is necessary. Only after a third dimensional separation, by cation-exchange HPLC, the purity of the arsenosugar is sufﬁcient to assure its intensity to be dominant among other peaks in the mass spectrum. Similar multidimensional puriﬁcation protocols were used for the identiﬁcation of selenium species in yeast [36,57] and garlic [58], and metallothioneins in rat tissues [59,60]. 37.8

SAMPLE PREPARATION PRIOR TO SPECIATION ANALYSIS OF BIOLOGICAL FLUIDS

37.8.1 Arsenic in urine Urine samples can be injected on a chromatographic column directly (ﬁltered) or after dilution with acid [61 – 64]. Urine samples were collected in

1196

Sample preparation for speciation analysis for metallobiomolecules

polycarbonate bottles and ﬁltered through a 0.45 mm ﬁlter [65]. The storage procedure implies the use of acid-washed polyethylene bottles at 2 108C [66] but storage at 208C [65] was also reported. A clean-up step (C18 cartridge extraction, dilution or freezing) was judged necessary to eliminate proteins and salts from the matrix [67]. The effects of the following storage conditions on the stability of arsenic species in human urine: temperature (25, 4 and 2208C), storage time (1, 2, 4 and 8 months) and the use of additives (HCl, sodium azide, benzoic acid, benzyltrimethylammonium chloride and cetylpyridinium chloride) were compared [68]. As(III), As(V), monomethylarsinic acid (MMA), dimethylarsonic acid (DMA) and arsenobetaine were stable for up to 2 months when urine samples were stored at 4 and 2208C without any additives [68]. No need for any sample pretreatment was claimed prior to ion pair HPLC separation followed by hydride generation atomic ﬂuorescence detection for speciation of MMA(III), DMA(III), As(III), As(V), MMA and DMA in urine samples [69]. A pre-oxidation procedure that converts arsenite into arsenate was used in urinary arsenic speciation prior to on-line photo-oxidation hydride generation with ICP-MS detection in order to eliminate As(III) and As(V) preservation concerns and simplify the chromatographic separation [70].

37.8.2 Selenium in urine Due to the complexity of urine matrix and low concentration of selenium, analytical methods for urinary speciation of Se are scarce. Urine samples were injected following 1:1 dilution with distilled water [71] or directly after ﬁltration only [72]. Cation-exchange HPLC-ICP-MS analysis of urine diluted 1 þ 1 suffered from a shift in retention times; tetramethyl selenonium could be separated from other species, but the signal from selenomethionine co-eluted with the selenate signal [73]. The use of crown ethers to eliminate salts from urine samples prior to HPLC or capillary zone electrophoresis (CZE) of organoselenium compounds was proposed [74]. The stability of ﬁve selenium compounds (selenate, selenourea, trimethylselenonium ion, selenomethionine and selenoethionine) in a pooled human urine, stored in dark at 2 20, 48C or ambient temperature without addition of any stabilizing reagent was evaluated. The general trend was the lower the temperature used for storage, the higher the stability of Se species, when other conditions such as light, acidity and container material are kept constant. On the basis of these results, it was considered that the storage of urine samples at 2208C for a short term (within one month) did not affect the selenium speciation. Long-term storage of urine samples for speciation analysis should, however, be undertaken with caution [75].

1197

J. Szpunar et al.

37.8.3 Metal complexes in biological ﬂuids The most common body ﬂuids of interest include: blood (subdivided by centrifugation into plasma (serum) and red cells (erythrocytes), amniotic ﬂuid, breast milk and urine. The major bioligands include proteins and small anions with speciﬁc functions, such as citrate, ATP, porphyrins or cobalamins. The largest interest is generated by essential elements that include some transition metals such as iron, copper and zinc and toxic metals such as Al, Cr, Pb, Cd and Hg. Metals can be an integral part of metalloproteins and metalloenzymes such as ferritin (Fe, Cu, Zn), b-amylase (Cu), alcohol dehydrogenase (Cd, Zn) and carbonic anhydrase (Cu, Zn) or bound less ﬁrmly to transport proteins (albumin, transferrin). Sampling, sample preservation and preparation prior to chromatography are particularly critical in clinical chemistry because of the low concentrations involved (risk of contamination) the thermodynamic instability of some species and the complexity of the matrix [4,5,76]. Sample preparation of serum prior to HPLC includes ﬁltration of sample through a 0.45 or 0.2 mm ﬁlter [77]. Erythrocytes (packed cells, red blood cells) need to be lysed to free their content prior to chromatographic separation. The need for three freeze–thaw cycles to lyse the cells followed by a 10-fold dilution with a buffer and centrifugation at 18,000g to remove fragments of membranes was reported [78,79]. An alternative procedure recommended by Cornelis et al. [4] was based on mixing one part of packed cells with one part of toluene and 40 parts of ice-cold water, followed by centrifugation and 0.45 mm ﬁltration of the lysate. When low-molecular weight compounds are of interest, ultraﬁltration on a 10 kDa ﬁlter is carried out [46,47,49]. The ﬁltrate is protein free and can be analysed for metallodrug metabolites or porphyrins, for example. In order to obtain milk whey, a sample aliquot of a size sufﬁcient for subsequent chromatographic analysis (typically 500 ml) was centrifuged at 50,000 rpm at 48C for 15 min (infant formulas and breast milk) or for 60 min (cow and goat milk), in order to remove fat. The fat (upper layer) and the insoluble residue at the bottom were discarded. The middle fraction was ﬁltered through a 0.45 mm syringe ﬁlter prior to subjecting it to SEC [81]. Precipitation of casein with 1 M acetate is optional [80]. Dialysis and puriﬁcation by SEC was required prior to reversed-phase HPLC [80]. Amniotic ﬂuid is a urine-like ﬂuid inhaled and swallowed by the human foetus. Some heavy metals, such as Pb, can cross the placenta and end up in amniotic ﬂuid. Metal-binding ligands are important in amniotic ﬂuid because of the potential of serving as transporters to the neurological system [81]. The amniotic ﬂuid sample was centrifuged and the supernatant was stored frozen at 2 208C prior to analysis [82,83].

1198

Sample preparation for speciation analysis for metallobiomolecules

37.8.4 Metallodrug metabolites in biological ﬂuids Figure 37.6 shows schematically the different fates possible for a metallodrug upon incubation with blood plasma. In addition to being covalently bound to plasma proteins, a drug may undergo degradation, resulting in the formation of a number of metabolites that need to be identiﬁed and determined. A point of interest is the determination of the pharmacologically active drug that is deﬁned as the intact drug that is either free or non-covalently bound to plasma proteins. Earlier studies for the determination of complexes of metallodrugs with proteins in serum were based on ultraﬁltration techniques employing cut-off ﬁlters that enabled the fractionation of serum into fractions that differed in terms of molecular mass, followed by the determination of the metal concentration in the separated fraction off-line by GF-AAS or ICP-OES [84]. The limited selectivity in terms of the molar mass of the separated complexes can be overcome by the use of SEC that can be directly coupled with ICP-MS [83].

Fig. 37.6. Puriﬁcation of elemental species by multidimensional chromatography prior to electrospray mass spectrometry.

1199

J. Szpunar et al.

Protein precipitation was employed to isolate a fraction of the intact drug together with its metabolites that are reversibly bound to serum proteins. Usually, dilute HClO4 was used for this purpose [85]. The deﬁnition of the total metal found in ultraﬁltrate or in the supernatant after protein precipitation as corresponding to the pharmacologically active fraction of the drug is misleading because several metabolites can be present. Hence, the need for a ﬁne characterization of the unbound fraction by a hyphenated technique. 37.9

SAMPLE PREPARATION PRIOR TO SPECIATION ANALYSIS IN SOLID MATRICES

37.9.1 Organoarsenic species in marine biota and foodstuffs For solid materials, the recovery of organic arsenic from the matrix and separation from the matrix are the prerequisite for a chromatographic analysis. The species of interest include As(III), As(V), monomethylarsinic and dimethylarsonic acids, arsenobetaine, arsenocholine and a number of arsenosugars. An overview of analytical techniques used can be found in two extensive reports [86, 87]. A freezing procedure has been commonly used to preserve biosamples; arsenic speciation results in fresh and defrosted samples were compared [88]. Arsenobetaine in sample extracts stored at 48C for 9 months was decomposed to trimethylarsine oxide and two other unidentiﬁed arsenic species [88,89]. Defatting, by leaching with acetone, for example, was recommended as the ﬁrst step of the analytical procedure [90]. Removal of lipids by shaking with diethylether or petroleum was proposed [91]. It is necessary to avoid generating an emulsion with the fat, which would make the subsequent clean-up more difﬁcult [92]. A mass balance needs to be performed on a routine basis because losses of the fat-soluble arsenic are common. The efﬁciency of the subsequent methanol extraction step is apparently higher for defatted samples than for non-defatted ones [90]. Because some seafood products are prepared in oil and generally tend to have a high salt content, an additional clean-up step, using a strong cation exchanger, for example, is required to eliminate the remains of liposoluble compounds not extracted with acetone [90]. The clean-up also avoids the pressure buildup due to accumulation of material on the column. The uncleaned samples generate problems associated with the ICP-MS cones and electrospray ion source. Extraction of arsenobetaine, arsenocholine and arsenoribosides has usually been performed with methanol, methanol–chloroform–water or methanol– water. The different methods were compared [93]. A methanol–water mixture is recommended for the dry tissues, whereas fresh samples can be efﬁciently leached with pure methanol. A precipitate of fatty aspect during CH3OH–H2O extraction was sometimes observed [93]. No degradation of arsenobetaine was observed when an enzymic (trypsin) digestion procedure was applied to ﬁsh samples [94,

1200

Sample preparation for speciation analysis for metallobiomolecules

95]. The methanolic extraction is typically repeated 2–3 times followed by preconcentration of the extract by evaporation of methanol using a rotoevaporator. The typical recoveries are of 90% for ﬁsh and 80% for mussel samples. There has been a surge of interest in the use of microwave-assisted procedures for the recovery of organoarsenic compounds from biological tissues [92,96,97]. An alternative is accelerated solvent extraction (ASE) that is based on performing static extractions at elevated temperatures and pressures. The pressure can be programmed without the use of elevated solvent temperatures that could lead to decomposition of thermally unstable compounds. The optimization of extraction of organoarsenic compounds from seaweed by ASE was discussed [98,99]. A mixture of methanol/water (1:1) was used to extract arsenic compounds from the plant tissue. Recoveries of 85 to 100% were obtained for most parts of the plant except for roots, for which extraction efﬁciency was approximately 60% [100]. 37.9.2 Low-molecular weight organoselenium species in yeast and plants Because selenoaminoacids are water soluble, leaching with hot water has been judged sufﬁcient to recover selenium species not incorporated in larger molecules. The sample is homogenized with water, sonicated or heated, and ultracentrifuged. The typical recovery of selenium extracted in this way is approximately 10–20% [17,19,26,101,102]. Selenocysteine and some other selenoaminoacids are highly susceptible to oxidative degradation, because the selenol group has a signiﬁcantly lower oxidation potential than its sulfur counterpart. The carboxymethyl derivative was synthesized (by addition of iodoacetic acid) to stabilize selenocysteine and prevent its degradation [103]. After breaking the cell walls with liquid nitrogen and grinding, the plant tissues were leached with 1 M HCl or Tris– HCl buffer (pH 8) allowing liberation of the free or weakly bound selenium (inorganic and selenoamino acids). Recoveries of about 50% were obtained for Brassica juncea leaves [21] and Brazil nuts [104]. Extraction with diethyl ether was used to remove fatsoluble material from extracts of nutritional supplements [105]. An example of the effect of sample preparation procedure on the recovery and speciation of a selenized microalgae (Spirulina) sample is shown in Fig. 37.7 [106]. The approach is composed of fours steps: extraction of watersoluble species, extraction of species bound to cell walls, solubilization of the protein fraction and proteolytic attack aimed at the liberation of aminoacids remained in the residue. A similar protocol was successfully used for studying selenium speciation in yeast [17] and garlic [58]. Care is advised in the interpretation of literature data because the results depend on the sample preparation procedure. This applies in particular to the frequently used statement “the majority of Se is present as selenomethionine”, describing the result of a procedure involving an enzymic digestion. Actually,

1201

3000

selenomethionine

Signal int., cps

4000

Se(IV) Se(VI)

exclusion volume

J. Szpunar et al.

leaching with hot water

2000 0

40

20

60

5000

Signal, cps

4000 enzymolysis with driselase 3000 2000 1000 0 0

20

40

60

Signal, cps

100000 80000 60000

solubilization with SDS

40000 20000 0 0

10

20

30

40

50

2500 Signal, cps

2000 1500 Enzymolysis with lipase-protease

1000 500 0 0

20 40 Time, min

60

Fig. 37.7. Effect of sample preparation procedure on the recovery and speciation for a selenized microalgae (Spirulina) sample.

1202

Sample preparation for speciation analysis for metallobiomolecules

selenomethionine usually constitutes a part of a larger stable selenoprotein that was destroyed during the sample preparation procedure. A low molecular fraction was obtained from yeast cell lysates [107] and nuts [104] by precipitation of proteins with 0.4 M perchloric acid. The proteins were isolated from nut samples by dissolution in 0.1 M sodium hydroxide and subsequent precipitation with acetone [104]. Because selenium was not detected in any of the lipid extracts obtained from the different types of nuts, lipids were extracted (chloroform–methanol, 2:1) and discarded prior to analysis [104]. 37.9.3 High-molecular weight selenium species in animal tissues and yeast In animal tissues, speciation of selenium is mostly concerned with the determination of the different Se-containing proteins [108–115]. The most important seems to be selenoprotein P, major protein that is sometimes used as a biochemical marker of selenium status [110], selenoenzymes, such as several glutathione peroxidases and type 1 iodothyronine de-iodinase [110,113,115], and albumin [109,113,114]. More than 25 Se-containing proteins or protein sub-units were detected in rat tissues labelled in vivo with Se-75 [115]. The sample preparation procedures include equilibration with a physiological buffer and ultracentrifugation. The recoveries are low unless SDS is added [17]. Frozen, pulverized in liquid nitrogen and lyophilized tissues were cleared of lipids with methylene chloride extraction and extracted with SDS-containing buffer, boiled for 5 min and centrifuged to recover extracted and denaturated proteins (including selenoproteins) [116]. 37.9.4 Metal complexes with metallothioneins MTs are a group of non-enzymatic low molecular mass (6 –7 kDa), cysteine-rich metal-binding proteins, resistant to thermocoagulation and acid precipitation [117–119]. Liver and kidney have been the most widely studied organs because of their crucial function in the metal metabolism. Guidelines for the preparation of biological samples prior to quantiﬁcation of MTs were discussed with particular attention given to the care necessary to avoid oxidation [120]. Figure 37.8 shows a scheme for the sample preparation prior to analysis for metal–metallothionein complexes. Sample preparation prior to analysis of metallothioneins by coupled techniques was reviewed [121,122]. Storage of fresh liver tissues led to a reduction in the MT content depending on duration and storage temperature. When liver samples were stored for 1 month at 2 808C, total MT losses relative to fresh liver were on average three times greater than after storage at 2208C. After 3 months’ storage the reductions of MT in livers stored at 280 and 2 208C were almost identical, the maximum loss being 44% MT [123].

1203

J. Szpunar et al.

Fig. 37.8. Sample preparation prior to analysis of biological samples for metal complexes with metallothionein. Adapted from Ref. [135].

Washing cells with a Tris–HCl buffer (pH 8) containing 1 M EDTA was recommended to remove metal ions reversibly bound to the cell wall [103]. The use of non-acidic buffers for tissue homogenization is mandatory because Zn starts to dissociate from protein complexes at pH 5 [127]. Cd and Cu are removed at lower pH values. For cytosols containing Cd-induced MTs, dilution

1204

Sample preparation for speciation analysis for metallobiomolecules

factors up to 10 were used, whereas for those with natural MT levels equal amounts of tissue and buffer were found suitable. Five extraction buffers were examined to maximize the MT recovery from rat liver [123]. The amounts of extracted MT from liver were highest for 10 mM Tris –HCl or 10 mM NaH2PO4 buffer. The use of a higher buffer concentration of 50 mM gave slightly lower MT yields. A mixture of 10 mM Tris –HCl and 0.25 M sucrose provided less MT than extraction with 10 mM Tris –HCl buffer. The highest MT concentrations were achieved with a liver/buffer ratio of 1:3 (w/v). Up to 50–80% of cadmium in rat and mouse tissue could be solubilized using a similar extraction procedures [30]. Heat treatment had a considerable impact on MT1 and MT2 concentrations in the sample; the highest MT levels were obtained after 1 or 2 min of incubation in a 808C water bath while lower temperatures or longer incubation periods decreased the total MT content [123]. Because MTs may be oxidized by oxygen, Cu(I) or heme components, the homogenization of tissues and subsequent isolation of MT should normally be performed in deoxygenated buffers and/or in the presence of a thiolic reducing agent [124]. b-Mercaptoethanol was added as antioxidant that additionally prevents formation of dimeric forms of MT [125]. As a result of centrifugation, two fractions: a dissolved one (supernatant, cytosol) and a particulate one (cell membranes and organelles) are obtained. Only the supernatant is analysed for metallothioneins. It is recommended that it be stored at 2208C under nitrogen prior to analysis [125]. The heating of the supernatant at 608C for 15 min allows precipitation (and removal) of the high-molecular weight proteins, leaving the MTs (which are heat stable) in the supernatant. This procedure allows one to decrease the protein load on the HPLC column, not only improving the separation of MT isoforms, but also prolonging the column lifetime. Several workers, however, preferred gel ﬁltration [126–130] to heat treatment [30,131,132] for the isolation of the metallothionein fraction from the tissue cytosol [133]. 37.10 SOURCES OF ERROR During species-selective analysis by hyphenated techniques, the major obstacles are contamination, break-up of the original metal –protein bond during the separation process and insufﬁcient detection limits for the element ion in the eluate, requiring preconcentration. Control materials, even when selected to have the appropriate matrix, undergo a substantial number of manipulations during their production which can alter the properties of the matrix. These alterations include human and non-human additives for achieving speciﬁc concentrations and/or stability, as well as physical changes to the material, such as lyophilization. These alterations may, in turn, cause interferences in the testing process which may not be present in fresh human samples.

1205

J. Szpunar et al.

Contamination risk. The risk of contamination at the level of sampling is as acute as in trace analysis for total metals and the same precautions should be applied. The ubiquitous presence in the laboratory environment of iron and zinc, and to a lesser extent copper, makes the contamination problem particularly acute. Protocols for sampling of milk and standard speciﬁcations for cleaning the material used have been discussed in detail [80]. The procedure seems likely to be necessary to be extended for other bioﬂuids. Any organic species that adhere to the column can also bind inorganic species, giving rise to anomalous peaks in subsequent runs. To avoid contamination of the analytical column by trace elements, buffers should be cleaned by cation exchange on Chelex-100. Stability of species and acid – base equilibria. Biomolecules are easily subject to deterioration. Because the complexation equilibria between complexes of metals with biomacromolecules and “free” metals are strongly pH dependent, the control of pH of the buffers used for leaching and puriﬁcation steps is crucial. Acidic pH is responsible for the depletion of metals, gradually leading to apo forms. For instance, Zn is lost from MT at pH 5, at pH 3.0 Cd4 adducts of isoform are present, at pH 2 copper still remains attached. Various intermediate partially metallated forms occur at various pH ranges. A possible Zn-transfer from one protein to another was examined by combining a Zn-containing protein with a Zn-free one. No change of the Znstatus of the protein was detected, indicating a stable protein metal complex under the experimental conditions [134]. REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18

1206

R. Lobinski, Appl. Spectrosc., 51 (1997) 260A. B. Bouyssiere, J. Szpunar and R. Lobinski, Spectrochim. Acta B, 57 (2002) 805. R. Cornelis and J. De Kimpe, J. Anal. At. Spectrom., 9 (1994) 945. R. Cornelis, J. De Kimpe and X. Zhang, Spectrochim. Acta B, 53 (1998) 187. A. Sanz Medel, Spectrochim. Acta B, 53 (1998) 197. J. Szpunar, Analyst, 125 (2000) 963. J. Szpunar and R. Lobinski, Pure Appl. Chem., 71 (1999) 899. M.J. Tomlinson, L. Lin and J.A. Caruso, Analyst, 120 (1995) 583. G.K. Zoorob, J.W. McKiernan and J.A. Caruso, Mikrochim. Acta, 128 (1998) 145. J.A. Caruso, K.L. Sutton and K.L. Ackley, Elemental Speciation. New Approaches for Trace Element Analysis. Elsevier, Amsterdam, 2000. R. Cornelis, Handbook of Speciation Analysis. Wiley, Chichester, 2002. D.M. Taylor and D.R. Williams, Trace Element Medicine and Chelation Therapy. Royal Society of Chemistry, Cambridge, 1995. B.K. Keppler, Metal Complexes in Cancer Chemotheraphy. VCH, New York, 1993. O.K. Hjelstuen, Analyst, 120 (1995) 863. R. Lobinski (Ed.), Metals and Biomolecules – Foreword, Analysis, 26 (1998) M20. A. Mazzucotelli, V. Bavastello, E. Magi, P. Rivaro and C. Tomba, Anal. Proc., 32 (1995) 165. C. Casiot, J. Szpunar, R. Lobinski and M. Potin Gautier, J. Anal. At. Spectrom., 14 (1999) 645. I. Leopold and B. Fricke, Anal. Biochem., 252 (1997) 277.

Sample preparation for speciation analysis for metallobiomolecules 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51

N. Gilon, A. Astruc, M. Astruc and M. Potin-Gautier, Appl. Organomet. Chem., 9 (1995) 623. C. B’Hymer and J.A. Caruso, J. Anal. At. Spectrom., 15 (2000) 1531. M. Montes-Bayon, T.D. Grant, J. Meija and J.A. Caruso, J. Anal. At. Spectrom., 17 (2002) 105. M. Montes-Bayon, E.G. Yanes, C. Ponce de Leon, K. Jayasimhulu, A. Stalcup, J. Shann and J.A. Caruso, Anal. Chem., 74 (2002) 107. M. Dernovics, Z. Stefanka and P. Fodor, Anal. Bioanal. Chem., 372 (2002) 473. Z. Stefa´nka, I. Ipolyi, M. Dernovics and M. Fodor, Talanta, 55 (2002) 437. K. Wrobel, S.S. Kannamkumarath, K. Wrobel and J.A. Caruso, Anal. Bioanal. Chem., 375 (2003) 133. S.M. Bird, P.C. Uden, J.F. Tyson, E. Block and E. Denoyer, J. Anal. At. Spectrom., 12 (1997) 785. M. Kotrebai, M. Birringer, J.F. Tyson, E. Block and P.C. Uden, Analyst, 125 (2000) 71. J.A. Caruso, D.T. Heitkemper and C. B’Hymer, Analyst, 126 (2001) 136. J. Szpunar, P. Pellerin, A. Makarov, T. Doco, P. Williams and R. Lobinski, J. Anal. At. Spectrom., 14 (1999) 639. H.M. Crews, J.R. Dean, L. Ebdon and R.C. Massey, Analyst, 114 (1989) 895. S. Mounicou, J. Szpunar, R. Lobinski, D. Andrey and C.J. Blake, J. Anal. At. Spectrom., 17 (2002) 880. L.M.W. Owen, H.M. Crews, R.C. Massey and N.J. Bishop, Analyst, 120 (1995) 705. K. Takatera and T. Watanabe, Anal. Chem., 65 (1993) 759. S. Mounicou, J. Szpunar, D. Andrey, C. Blake and R. Lobinski, Analyst, 127 (2002) 1638. S. McSheehy, J. Szpunar, R. Lobinski, V. Haldys, J. Tortajada and J. Edmonds, Anal. Chem., 74 (2002) 2370. S. McSheehy, P. Pohl, J. Szpunar, M. Potin-Gautier and R. Lobinski, J. Anal. At. Spectrom., 16 (2001) 68. S. McSheehy, P. Pohl, R. Lobinski and J. Szpunar, Analyst, 126 (2001) 1055. S. McSheehy, P. Pohl, R. Lobinski and J. Szpunar, Anal. Chim. Acta, 440 (2001) 3. S. McSheehy, F. Pannier, J. Szpunar, M. Potin-Gautier and R. Lobinski, Analyst, 127 (2002) 223. S. McSheehy, V. Haldys, J. Tortajada and J. Szpunar, J. Anal. At. Spectrom., 17 (2002) 507. Y. Ogra, Personal communication. B. Michalke and P. Schramel, J. Chromatogr. A, 807 (1998) 71. A. Theobald and L. Dunemann, J. High Resolut. Chromatogr., 19 (1996) 608. H.B. Roellin and C. Nogueira, Eur. J. Clin. Chem. Clin. Biochem., 35 (1997) 215. W.R.L. Cairns, L. Ebdon and S.J. Hill, Fresenius J. Anal. Chem., 355 (1996) 202. W.A.J. De Waal, F.J.M.J. Maessen and J.C. Kraak, J. Chromatogr., 407 (1987) 253. S.G. Matz, R.C. Elder and K. Tepperman, J. Anal. At. Spectrom., 4 (1989) 767. Z. Zhao, W.B. Jones, K. Tepperman, J.G. Dorsey and R.C. Elder, J. Pharm. Biomed. Anal., 10 (1992) 279. G.K. Poon, F.I. Raynaud, P. Mistry, D.E. Odell, L.R. Kelland, K.R. Harrap, C.F.J. Barnard and B.A. Murrer, J. Chromatogr. A, 712 (1995) 61. K. Lange Hesse, L. Dunemann and G. Schwedt, Fresenius J. Anal. Chem., 339 (1991) 240. K. Lange Hesse, Fresenius J. Anal. Chem., 350 (1994) 68.

1207

J. Szpunar et al. 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87

1208

K. Lange Hesse, L. Dunemann and G. Schwedt, Fresenius J. Anal. Chem., 349 (1994) 460. K. Guenther and A. Von Bohlen, Spectrochim. Acta B, 46 (1991) 1413. K. Guenther and H. Waldner, Anal. Chim. Acta, 259 (1992) 165. K. Guenther, A. von Bohlen and C. Strompen, Anal. Chim. Acta, 309 (1995) 327. S. McSheehy, P. Pohl, D. Velez Pacios and J. Szpunar, Anal. Bioanal. Chem., 372 (2002) 457. T. Lindemann and H. Hintelmann, Anal. Chem., 74 (2002) 4602. S. McSheehy, W. Yang, F. Pannier, J. Szpunar, R. Lobinski, J. Auger and M. PotinGautier, Anal. Chim. Acta, 421 (2000) 147. K. Polec, M. Pere´z-Calvo, O. Garcia-Arribas, J. Szpunar, B. Ribas-Ozonas and R. Lobinski, J. Inorg. Biochem., 88 (2001) 197. K. Polec, M. Pere´z-Calvo, O. Garcı´a-Arribas, J. Szpunar, B. Ribas-Ozonas and R. Lobinski, J. Anal. At. Spectrom., 15 (2000) 1363. D.L. Tsalev, M. Sperling and B. Welz, Analyst, 123 (1998) 1703. E.H. Larsen, G. Pritzl and S.H. Hansen, J. Anal. At. Spectrom., 8 (1993) 557. X.R. Zhang, R. Cornelis, J. De Kimpe, L. Mees and N. Lameire, Clin. Chem., 43 (1997) 406. M. Ma and X.C. Le, Clin. Chem., 44 (1998) 539. B.S. Chana and N.J. Smith, Anal. Chim. Acta, 197 (1987) 177. D. Heitkemper, J. Creed, J. Caruso and F.L. Fricke, J. Anal. At. Spectrom., 4 (1989) 279. L. Benramdane, F. Bressolle and J.J. Vallon, J. Chromatogr. Sci., 37 (1999) 330. J. Feldmann, V.W. Lai, W.R. Cullen, M. Ma, X. Lu and X.C. Le, Clin. Chem., 45 (1999) 1988. X.C. Le, X. Lu, M. Ma, W.R. Cullen, H.V. Aposhian and B. Zheng, Anal. Chem., 72 (2000) 5172. X. Wei, C.A. Brockhoff-Schwegel and J.T. Creed, Analyst, 125 (2000) 1215. J.M. Marchante-Gayon, I. Feldmann, C. Thomas and N. Jakubowski, J. Anal. At. Spectrom., 16 (2001) 457. J. Zheng, M. Ohata and N. Furuta, J. Anal. At. Spectrom., 17 (2002) 730. B. Gammelgaard, K.D. Jessen, F.H. Kristensen and O. Jøns, Anal. Chim. Acta, 404 (2000) 47. B. Gammelgaard, O. Joens and L. Bendahl, J. Anal. At. Spectrom., 16 (2001) 339. J. Zheng, Y. Shibata and A. Tanaka, Anal. Bioanal. Chem., 374 (2002) 348. D. Behne, Analyst, 117 (1992) 555. G.F. Van Landeghem, P.C. D’Haese, L.V. Lamberts and M.E. De Broe, Anal. Chem., 66 (1994) 216. I.A. Bergdahl, A. Schuetz and A. Grubb, J. Anal. At. Spectrom., 11 (1996) 735. B. Gercken and R.M. Barnes, Anal. Chem., 63 (1991) 283. Y. Makino and S. Nishimura, J. Chromatogr., Biomed. Appl., 117 (1992) 346. G.S. Hall, E.G. Zhu and E.G. Martin, Anal. Commun., 26 (1999) 93. L. Fernandez Sanchez and J. Szpunar, J. Anal. At. Spectrom., 14 (1999) 1697. J. Szpunar, A. Makarov, T. Pieper, B.K. Keppler and R. Lobinski, Anal. Chim. Acta, 387 (1999) 135. R.J. Einha¨user, M. Galanski and B.K. Keppler, J. Anal. At. Spectrom., 11 (1996) 747. V. Vacchina, Personal communication. M. Burguera and J.L. Burguera, Talanta, 44 (1997) 1581. T. Guerin, A. Astruc and M. Astruc, Talanta, 50 (1999) 1.

Sample preparation for speciation analysis for metallobiomolecules 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109 110 111 112 113 114 115 116 117 118

S.X.C. Le, W.R. Cullen and K.J. Reimer, Environ. Sci. Technol., 28 (1994) 1598. A.J.L. Muerer, A. Abildtrup, O.M. Poulsen and J.M. Christensen, Analyst, 117 (1992) 677. N. Ybanez, D. Velez, W. Tejedor and R. Montoro, J. Anal. At. Spectrom., 10 (1995) 459. K. Shiomi, Y. Sugiyama, K. Shimakura and Y. Nagashima, Appl. Organomet. Chem., 9 (1995) 105. T. Dagnac, A. Padro, R. Rubio and G. Rauret, Talanta, 48 (1999) 763. J. Alberti, R. Rubio and G. Rauret, Fresenius J. Anal. Chem., 351 (1995) 420. S. Branch, L. Ebdon and P. O’Neill, J. Anal. At. Spectrom., 9 (1994) 33. K.J. Lamble and S.J. Hill, Anal. Chim. Acta, 334 (1996) 261. K.L. Ackley, C. B’Hymer, K.L. Sutton and J.A. Caruso, J. Anal. At. Spectrom., 14 (1999) 845. H. Helgesen and E.H. Larsen, Analyst, 123 (1998) 791. J.W. McKiernan, J.T. Creed, C.A. Brockhoff, J.A. Caruso and R.M. Lorenzana, J. Anal. At. Spectrom., 14 (1999) 607. P.A. Gallagher, J.A. Shoemaker, X.N. Wei, C.A. Brockhoff-Schwegel and J.T. Creed, Fresenius J. Anal. Chem., 369 (2001) 71. W. Zhang, Y. Cai, C. Tu and L.Q. Ma, Sci. Total Environ., 300 (2002) 167. S.M. Bird, H. Ge, P.C. Uden, J.F. Tyson, E. Block and E. Denoyer, J. Chromatogr. A, 789 (1997) 349. J. Zheng, W. Goessler and W. Kosmus, Trace Elem. Electrol., 15 (1998) 70. K. Takatera, N. Osaki, H. Yamaguchi and T. Watanabe, Anal. Sci., 10 (1994) 567. S.S. Kannamkumarath, K. Wrobel, K. Wrobel, A. Vonderheide and J.A. Caruso, Anal. Bioanal. Chem., 373 (2002) 454. G. Matni, R. Azani, M.R. Van Calsteren, M.C. Bissonnette and J.S. Blais, Analyst, 120 (1995) 395. J. Cases, I.A. Wysocka, B. Caporiccio, N. Jouy, P. Besanc¸on, J. Szpunar and J.M. Rouanet, J. Agric. Food Chem., 50 (2002) 3867. K. Wrobel, K. Wrobel and J. Caruso, J. Anal. At. Spectrom., 17 (2002) 1048. C.D. Thomson, Analyst, 123 (1998) 827. T. Plecko, S. Nordmann, M. Ru¨kgauer and J.D. Kruse-Jarres, Fresenius J. Anal. Chem., 363 (1999) 517. M. Persson Moschos, W. Huang, T.S. Srikumar, B. Akesson and S. Lindeberg, Analyst, 120 (1995) 833. H. Koyama, Y. Kasanuma, C. Kim, A. Ejima, C. Watanabe, H. Nakatsuka and H. Satoh, Tohoku J. Exp. Med., 178 (1996) 17. H. Koyama, K. Omura, A. Ejima, Y. Kasanuma, C. Watanabe and H. Satoh, Anal. Biochem., 267 (1999) 84. I. Harrison, D. Littlejohn and G.S. Fell, Analyst, 121 (1996) 189. J.T. Deagen, J.A. Butler, B.A. Zachara and P.D. Whanger, Anal. Biochem., 208 (1993) 176. D. Behne, C. Weiss Nowak, M. Kalcklosch, C. Westphal, H. Gessner and A. Kyriakopoulos, Analyst, 120 (1995) 823. T.W.M. Fan, E. Pruszkowski and S. Shuttleworth, J. Anal. At. Spectrom., 17 (2002) 1621. M.J. Stillman, Coord. Chem. Rev., 144 (1995) 461. M.J. Stillman, C.F. Shaw and K.T. Suzuki (Eds.), Metallothioneins: Synthesis, Structure and Properties of Metallothioneins, Phytochelatins and Metal-Thiolate Complexes, Wiley, New York, 1992.

1209

J. Szpunar et al. 119 120 121 122 123 124 125 126 127 128 129 130 131 132 133 134 135

1210

J.F. Riodan and B.L. Valee (Eds.), Metallobiochemistry Part B. Metallothionein and Related Molecules. Academic Press, Orlando, 1991. K.T. Suzuki and M. Sato, Biomed. Res. Trace Elem., 6 (1995) 51. R. Lobinski, H. Chassaigne and J. Szpunar, Talanta, 46 (1998) 271. A. Prange and D. Schaumloffel, Anal. Bioanal. Chem., 373 (2002) 441. K. No¨stelbacher, M. Kirchgessner and G.I. Stangl, J. Chromatogr. B, 744 (2000) 273. K.A. High, B.A. Methven, J.W. McLaren, K.W.M. Siu, J. Wang, J.F. Klaverkamp and J.S. Blais, Fresenius J. Anal. Chem., 351 (1995) 393. H. Van Beek and A.J. Baars, J. Chromatogr., 442 (1988) 345. S.G. Ang and V.W.T. Wong, J. Liq. Chromatogr., 14 (1991) 2647. S.G. Ang and V.W.T. Wong, J. Chromatogr., 599 (1992) 21. M. Apostolova, P.R. Bontchev, C. Nachev and I. Sirakova, J. Chromatogr., Biomed. Appl., 131 (1993) 191. H. Van Beek and A.J. Baars, At. Spectrosc., 11 (1990) 70. O.M. Steinebach and H.T. Wolterbeek, J. Chromatogr., Biomed. Appl., 130 (1993) 199. A. Mazzucotelli and P. Rivaro, Microchem. J., 51 (1995) 231. A. Mazzucotelli, A. Viarengo, L. Canesi, E. Ponzano and P. Rivaro, Analyst, 116 (1991) 605. K. Polec, M. Pere´z-Calvo, O. Garcı´a-Arribas, J. Szpunar, B. Ribas-Ozonas and R. Lobinski, J. Anal. At. Spectrom., 15 (2000) 1363. B. Michalke and P. Schramel, J. Trace Elem. Electrolytes Health. Dis., 4 (1990) 163. K.T. Suzuki and M. Sato, Biomed. Res. Trace. Elem., 6 (1995) 51.

Chapter 38

Sample preparation for the analysis of volatile metal species Jo¨rg Feldmann

38.1

INTRODUCTION

The analysis of metal species in environmental and biological samples consists of four consecutive steps: sampling, sample preparation, separation and detection. Major attention has been focused, especially by the analytical community, on the separation and detection of the metal species. Clearly, the processes of sampling and sample preparation have a strong inﬂuence on the correctness and the quality of the analysis. Factors such as homogeneity and representativeness are unique for the sampling step, while stability is the major factor which determines sample preparation. Stoeppler [1] addressed this theme in a very useful reference book, however, it does not cover the analysis of volatile metal compounds (VMCs) in detail. For the analysis of metal species in different atmospheres, various strategies have been employed. All these methods have a preconcentration step in common, which entails the separation of the analyte from the major gas constituents. In special cases when whole gas samples are needed, balloons in their simplest form, or cylinders with inlets and exit valves, internally polished stainless steel canisters or plastic bags (Tedlar or Teﬂon) can be used. Sampling and sample preparation often becomes one step when volatile compounds are the focus of the analysis [2,3]. The formation of VMCs has been studied since the beginning of the 20th century, with the identiﬁcation by Gosio [4,5] and Challenger [6,7] of volatile arsenic compounds generated by a fungi (Scopulariopsis brevicaulis). Obviously, scientists in those days used other methods for the identiﬁcation of such arsenic species (Me3As), which today is a much easier task than 100 years ago. However, the instability of these VMCs is as pronounced now as it was in the past. Most volatile metal(loid) compounds (VMCs) are thermodynamically unstable, and thus prone to degradation of various kinds, which becomes important when sample preparation is to be considered. Sample preparation should, therefore, be designed to prevent oxidation, hydrolysis, Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

1211

J. Feldmann

photo-decomposition, irreversible ad- or absorption and heterogeneous surfacecatalyzed decomposition reactions. Since the sample preparation should be as gentle as possible, to guarantee the integrity of the metal species, or at least to preserve their molecular information, the preparation procedure is dependent on the species involved. In this chapter, the variety of metal species is discussed, followed by a section focused on the characterization of the different types of gas components in which the VMCs occur, only to ﬁnalize the chapter with concrete recipes for sample preparation for different types of analysis. 38.2

SPECIES OF INTEREST

VMCs have been identiﬁed in various anthropogenic gases, such as landﬁll gas and sewage sludge digester gas [8,9], and in the natural environment such as hot springs [10] and in the open ocean [11]. Only compounds having high vapor pressures at moderate environmental temperatures lead to low-temperature volatilization of metals and metalloids. Non-charged hydrides, alkylated compounds or halides meet these requirements and they are principally derived from main group elements of groups 12 –17. The simplest alkyl group is methyl, [CH3 – ] and methylmetal(loid) derivatives comprise the most frequently detected organometals in the natural environment, due to a common enzymatically catalyzed methylation process. Covalent metal hydrides and their methylated counterparts are gases or volatile liquids at ambient temperatures. Attachment of hydrogen or alkyl groups directly to metals and metalloids of relatively high charge usually substantially enhances volatility. The lack of strong intermolecular attractions, such as hydrogen bonding or dipole –dipole interactions, are the reasons behind this phenomenon. The mobility, and especially the stability, are heavily dependant on such fundamental properties such as boiling point, the strength of the metal –carbon bond (indicated by the bond dissociation energies) and their thermodynamic properties (mainly standard molar enthalpies of formation). From Table 38.1, it can be seen that most of these compounds have boiling points above temperatures encountered in normal environmental conditions, but have a vapor pressure higher than 0.01 mbar at 258C, and are therefore, considered to be highly volatile. The dissociation energies of most VMCs are very low, enthalpy of formation is negative for nearly all of the compounds under consideration, which is a sign of low stability. On the other hand, the metal– carbon bond energies do, in fact, lie within the normal range of chemical bond energies. So other scenarios must be considered as well, namely the kinetic stability. Although intrinsically unstable, some of the compounds might be kinetically stabilized if there are high energy barriers to overcome before decomposition to a thermodynamically favorable compound can occur. This has

1212

Sample preparation for the analysis of volatile metals species TABLE 38.1 Boiling points and dissociation energies of metal–hydrogen and metal carbon bonds in selected environmentally important volatile organometallic compounds Element

Species

As

AsH3 MeAsH2 Me2AsH Me3As AsCl3 SbH3 MeSbH2 MeSbH2 Me3Sb SbCl3 BiH3 MeBiH2 Me2BiH Me3Bi GeH4 Me4Ge SnH4 MeSnH3 Me2SnH2 Me3SnH Me4Sn BuSnH3 MeSnCl3 Me2SnCl2 Me3SnCl3 SnCl4 Me2Cd Me3Tl PbH4 Me4Pb Et4Pb SeH2 MeSeH

Sb

Bi

Ge Sn

Cd Tl Pb

Se

Dissociation energy (mean) M–H (kJ/mol) [65]

Dissociation energy (mean) M–C (kJ/mol) [66]

332

235 265

222 217

146 ,320 260 264

222

147 92 176 159 152 315

Boiling point (8C) [65] 255 2 36 50 217 41a 61a 82 283 17 72a 103a 109 288 43.4 252 14 35 57 78 100 171 189 154 114 106 147 213a 110 200a 241.5 25.5 continued

1213

J. Feldmann

TABLE 38.1 (continuation) Element

Te

Hg Ni Fe Mo W

Species

Dissociation energy (mean) M–H (kJ/mol) [65]

Me2Se Me2Se2 TeH2 MeTeH Me2Te Me2Hg Ni(CO)4 Fe(CO)5 Mo(CO)6 W(CO)6

Dissociation energy (mean) M–C (kJ/mol) [66]

Boiling point (8C) [65]

.210

55 152 22 57 94 92 43 105 156a 175a

268

126

a

Extrapolated boiling point.

often been observed when the compounds are studied in very dilute gas samples. For instance, Et4Pb and Me2Hg have high decomposition energies of 290 and 2 41 kcal/mol, respectively. Although an explosive decomposition of both compounds is predicted according to thermodynamic data, Me2Hg and Et4Pb have been shown to exist in signiﬁcant levels in the environment; Et4Pb in urban air [12] and Me2Hg in sea water [13]. Although permethylated derivatives of metals and metalloids have relatively high molecular weights, they occur as gases or volatile liquids at room temperature. Such hydrides enter the natural environment through various pathways. A simpliﬁed classiﬁcation in terms of types of sources may consist of three principal categories: two different sources of anthropogenic origin; accidental release as unwanted by-product or deliberate introduction (mostly as fungicides); and sources of biogenic origin, i.e. generation through biotransformation of metals. Previous studies have revealed that the VMCs can be found in anaerobic reducing environments, such as municipal waste deposits or anaerobic digesters of sewage sludge [14,17]. Compounds such as Me2Se, Me3As, Me2AsH, MeAsH2, AsH3, Me4Sn, Me2Et2Sn, Me3Sb, Me2Te, MeI, Me2Hg, Me4Pb and Me3Bi have been identiﬁed in concentrations ranging from ng/m3 to mg/m3. Natural environments can also produce VMCs, such as algae mats on hot spring pools [10] or contaminated sediments [15]. In addition to various arsines, Me3Sb and up to ﬁve different iodine species are released by algae mats [10], whereas sediments can release mixed butylated –methylated tin compounds into the water column [16]. Transition metals, such as Co and Ni, only bind to –CH3 when the d-orbitals of low energy are ﬁlled with electrons

1214

Sample preparation for the analysis of volatile metals species

donated by strong ligands, i.e. the tetrapyrrole ring in methyl-cobalamin. Thus, no volatile covalent permethylated transition metal compound is expected to play a signiﬁcant role in biological systems and/or the environment. However, due to the strong afﬁnity of CO to the transition metals, neutral covalent metal carbonyls of relatively low molecular weight can be formed. Ni(CO)4, W(CO)6 Mo(CO)6) have been detected in sewage gas and landﬁll gas [17]. The only other class of compounds rarely studied are the covalent species of halogenated metals and metalloids; for example, SnCl4 and AsCl3. In contrast to the covalent organometal compounds, however, these species are easy to hydrolyze, such that the M –Cl bond is cleaved and replaced by –OH. This results in a low volatility compound due to the formation of intermolecular hydrogen bonds. In the water phase, these halogenated metal species occur in thermodynamic equilibrium with their hydroxylated counterparts, so that the halogenated compounds might be released from an aquatic environment if the equilibrium is shifted by high levels of dissolved chloride, as illustrated in a study by Mester and Sturgeon [18]. All of these metal species are produced at ambient temperature mainly of biogenic origin in environments which are characterized by high microbial activity. In addition, VMCs can also be expected at ambient temperature in industrial processes. In workplace air, the following compounds can be encountered: – – –

Ni(CO)4 in nickel reﬁneries AsH3 and SbH3 in battery factories and electrolysis of metals Me3Ga, Me3In, tert-butylarsine in semi-conductor workplace air

VMCs can be studied not only in gas samples, they often show slight solubility in water. The water can be sampled and the VMCs liberated by purging with helium. For example, Pongratz and Heumann [13] determined the amount of Me2Hg in seawater, while Frankenberger and co-workers [19] studied the ﬂux of Me2Se from the soil by measuring the ﬂux directly with specially designed ﬂux chambers. In addition to all these volatile metal species which occur at ambient temperature, it should not be forgotten that there are high temperature processes in nature, such as volcanoes, or industrial processes such as incineration, to mention only a few, in which metals can be released into the gas phase. Such metals are usually sampled by ﬁltration, electrostatic condensation and nuclear condensation as well as by impaction and impinger techniques. Since their speciation changes with temperature and the density of the gas, only on-line methods are capable of identiﬁcation of such species in gas samples. This is the reason why such samples are not discussed in this chapter. The interested reader, however, is advised to obtain more information on this subject from a review by Tegheder and Khvostikov [20] for the collection and determination of metal contaminants in gases in which such VMCs are covered.

1215

J. Feldmann

38.3

CHARACTERIZATION OF GAS SAMPLES

In order to choose the most appropriate sampling/preconcentration procedure, it is important to consider the gas matrix components. In general, there are two different areas in which VMCs could occur; one is characterized by high temperature and is mainly of anthropogenic origin, i.e. incineration, welding processes, ore roasting. Here, stack or ﬂue gases can contain VMCs. Such gases have, in general, high levels of CO2, air, some CO, NOx, and SO2 in addition to many different VOCs, mainly semi-volatiles such as PAHs. In addition, natural high temperature processes, such as volcanic erruptions, also generate high levels of VMCs (in particular Hg0 and maybe AsH3). Signorelli and co-workers [21,22] report that gases from fumeroles are rich in CO2 and SO2 and sometimes CH4 and H2S. The second type of gas sample in which VMCs can occur are characterized by a high microbial activity and can be described as low temperature processes. Here, again, industrial processes, such as the fermentation of sewage sludge within sewage treatment plants and the land ﬁlling of biodegradable waste, are the main areas in which many VMCs have been identiﬁed by Feldmann and coworkers [23]. The generated gases often contain a mixture of hundreds of VOCs besides the main gases [CH4 (20– 60%) and CO2 (20– 50%)]. Soil gas or intraoral airare gases which are super-saturated with water and contain a signiﬁcant amount of CO2. Furthermore, work place air in semi-conductor plants in which trimethylated metal compounds such as (CH3)3Ga and (CH3)3Sb have been used, contain small amounts of CO2. In molecular organic compound vapor deposition (MOCVD) and molecular beam epitaxy (MBE) processes, vacuum oil in contact with the deposition chamber has to be exchanged and workers are exposed to VMCs. When the oil is checked for VMCs, the headspace obviously contains high levels of volatile hydrocarbons, which should be taken into consideration because of the enormous matrix effects, which arise under certain sampling and preconcentration procedures. Natural gas and gas condensates seem to contain large amounts of other alkanes besides CH4 [24], while headspace associated with any kind of microorganism media is characterized by the class of fungi or bacteria. Anaerobic cultures tend to contain mainly CH4 and CO2 besides enormous amounts of H2S, while aerobic cultures contain mainly air. 38.4

SAMPLE PREPARATION STRATEGIES

The sampling method has to be matched to the preconcentration, clean-up and analytical technique used for subsequent determination. For sampling, there are two different concepts to be followed; one is the on-site preconcentration and/or clean up of the sample, the other is the sampling of the entire gas sample—so-called canister sampling. While all strategies for the ﬁrst case are

1216

Sample preparation for the analysis of volatile metals species

identical with those described in Section 38.4.2, is it necessary to point out the applicability of canister sampling for VMCs. 38.4.1 Sampling Only very limited information is available concerning the stability of VMCs in canisters of any kind of material. Tedlarw bags provide a convenient method to sample gases. VMCs show surprisingly high recoveries for a number of volatile arsenic, antimony and tin compounds [25]. Haas et al. [26] sampled volumes of 5–10 l landﬁll or sewage gas in several German and Scottish sewage treatment plants and landﬁll sites. Tedlarw bags have been described in the literature to be useful for sampling of various volatile organic compounds (VOC) [27,28]. As well, they have been successfully used for the generation of standard gaseous mixtures and their storage for studying gaseous media containing traces of analytes (lower ppbv) [29]. The results indicate the excellent reliability of Tedlarw bags, whereas Teﬂon bags are seen to cause some problems. Tedlar bags can be easily ﬁlled by gases extracted from gas wells and from landﬁll sites and sewage sludge fermentation tanks. Stainless-steel containers are ofﬁcially used in the US Environmental Protection Agency canister method TO-14 [30] when VOCs are monitored in urban air. It has to be mentioned that this method describes the use of the canisters in the passive mode (i.e. vacuum ﬁlling) to eliminate any introduction of contaminants originating from the pump. On the other hand, critical reports highlight possible complications and constraints of using Tedlarw bags, especially the evidence of signiﬁcant VOC losses and adsorption. Whereas Lipari [31] found methanol losses of over 70% in 6 h, Andino and Butler [32] also studied methanol vapor storage in Tedlarw bags and found only about 8% losses occurring in 60 l bags in 6 h. Apart from all these more scientiﬁc and methodological considerations, aspects of ﬂexibility of sample handling and sub-sampling, as well as sample management, have to be taken into account. As far as cryotrapping is concerned, the use of liquid nitrogen on-site, carrying a pump as well as the necessary power supply is not very convenient and the large amounts of liquid nitrogen needed during a sampling trip represent a major hazard during the transport. Tedlarw bags are inexpensive and easy to use and are suitable for large screening studies. 38.4.2 Preconcentration According to Pellizzari et al. [33], the evaluation of the best method for study of volatile compounds should include the consideration of the following criteria: † † †

quantitative collection efﬁciency high recovery of trapped metal species high breakthrough volume

1217

J. Feldmann

† † †

minimal decomposition and polymerization low background and blank levels small afﬁnity for water and carbon dioxide.

Before particular protocols are listed for the different type of samples, it has to be established what a VMC is. In the past, and even today, metal species have been deﬁned in occupational hygiene by the analytical method used. If metal species can pass through a ﬁlter, then they are by deﬁnition non-ﬁlterable metal species. Ito and Shooter [34] ﬁltered urban air and trapped the metal species passing through a 0.45 mm membrane in nitric acid and determined VMCs in air. No speciation information was given here. This fraction is an operationally deﬁned fraction, which cannot describe the compounds on a molecular basis. However, without any ﬁrm evidence, a molecular well-deﬁned metal species is often assigned to the non-ﬁlterable fraction; e.g. Filkova and Ja¨ger [35] reported volatile nickel in urban air as Ni(CO)4 and Pederson [36] assigns AsH3 to the arsenic fraction which passes through ﬁlters. On the other hand, speciation analyses have been made by identifying different species of various elements by using a variety of different preconcentration and separation methods. However, with any manipulation process there is a risk of changing the species. Use of gas chromatography, for example, could prevent the detection of very unstable volatile compounds, which decompose on the column or do not elute from the column. Simple use of gas chromatography, therefore, restricts the determination of volatile metal species to an operationally deﬁned fraction of the air sample. This is illustrated in the latest study by Paveaux et al. [37], who were able to determine volatile copper and selenium species in the ﬂue gas of a coal-ﬁred power plant using cryotrapping GC coupled to ICP-MS. Their impinger measurements on the ﬂue gas, in which the “gaseous metals” were trapped in a solution of HNO3/K2Cr2O7 (for mercury) or HNO3/H2O2 (for all other metallic compounds), showed that 90% of the sample was lost during the trapping and chromatographic separation. They speculated that the lost metal compounds were microaerosols. This illustrates the problem, although a gentle non-selective sampling technique has been employed, only the fraction which is less reactive and present as a true volatile compound can elute from a chromatographic column; the rest is lost. In other words, if the non-ﬁlterable fraction is measured as total metal, no assignment to speciﬁc VMCs in this fraction is, therefore, possible and this fact excludes those techniques from consideration in this book. A variety of different collection and preconcentration devices has been employed for VMCs and VOCs in different atmospheres. The different methods can be categorized into three different groups. Methods are based on † † †

adsorptive absorptive and cryogenic trapping.

1218

Sample preparation for the analysis of volatile metals species

Solids or liquids coated on a solid supporting material can interact with volatile compounds such that these compounds can be removed from the gas phase and “stuck” to the surface. Although the chemical nature of the adsorbents is very important for the efﬁciency of the collection device, more important is the strength of the interaction. This method can be seen to be a broad-based collection system. The collected species adsorb to the surface with other compounds having a higher density than the gas phase, which can lead to reactions. This is often the reason for the occurrence of artifacts or irreversible reactions. Solvents or absorbing liquids cannot be used for the sampling of VMCs because the interactions needed for the analyte to be caught in the absorbent are far too high such that a change in the chemical form of the VMCs would be inevitable. It should be mentioned here, though, that the likelihood of a species transformation increases with the strength of the interaction between the solvent and the collected analyte. For the same reason, adsorbents have been considered unsuitable for unstable compounds such as VMCs. Irreversible adsorption, degradation during the desorption process and build-up of artifacts [38,39] are possible negative results associated with using materials such as Tenaxw and Porapakw, especially when they are stored at ambient temperatures. Tenaxw has often been used for trapping VOCs in air in both a passive and active way. A combination of Tenaxw and Porapakw was successfully applied to the determination of tetraalkyl lead species in urban air by Nerin and Pons [40] and Harrison and co-workers [41]. While they noted that such tetraalkyl lead compounds are stable on the solid cartridge when they were stored at 48C for less than 45 days [42], (CH3)3Sb was less stable, and they recommend trapping the organoantimony compounds anaerobically, so that no oxidation can occur on the substrate. However, it should be noted that species with low boiling points, and polar compounds, will not be trapped as efﬁciently, therefore, Tenaxw is ideal for semi-volatile species, which are not reactive and occur in water-saturated gas samples since this adsorbent has a low afﬁnity to water. Although US-EPA method TO-17 is based on sampling onto solid adsorbents [43], adsorbents might be tailored for a small number of VMCs, as for example, sequential sampling for the determination of volatile Hgspecies using a noble-metal trap in series with an activated-carbon trap [44]. For this purpose, the use of gold –platinum gauze, which binds the mercury in the vapor phase, is recommended. This approach was recently used for determination of mercury in cigarette smoke by Chang et al. [45]. Total mercury could be sampled on iodated charcoal or on gold traps [46], while Me2Hg can be adsorbed on Carbotraps [47]. Schroeder and co-workers [48] used different adsorption tubes in series, thereby binding different mercury species. Although information about the mercury speciation can be deduced by using these methods, only desorption from the Carbotrap actually gives species

1219

J. Feldmann

information, whereas the other methods only operationally deﬁne the amount of certain mercury fractions. Absorptive methods are very selective, since the collected species chemically bind very strongly to the stationary phase. Due to the reaction involved, this type of method is very selective and even species-speciﬁc. This has often been used as a selective chemofocusing method for the determination of volatile arsenic and antimony species. Cullen and co-workers [49] used a method developed by Challenger [7]. The headspace of a micro-organism culture was swept through a solution of HgCl2 in hydrochloric acid in order to trap (CH3)3As as an (CH3)3Asp2 HgCl2 adduct. The volatile arsine can be characterized as trimethylhydroxylarsonium nitrate or picrate. This method, very speciﬁc for volatile trivalent metalloids of group 15, cannot be used for most other volatile species such as (CH3)4Sn. However, this is one of the few absorptive methods for which the species information is conserved and traceable. In contrast to most methods based on absorption, this chemofoccusing makes it suitable for speciation analysis. Cryogenic methods are described and discussed in detail by Donard et al. [50]. Brieﬂy, compounds less volatile than the main matrix gases, such as oxygen and nitrogen, can be collected by freezing out the compounds which have a lower vapor pressure at a certain temperature. Tubes are submerged in cryogenic liquids, which have a characteristic minimum temperature (Table 38.2). By choosing the right cryogenic ﬂuid, the appropriate temperature can be set easily, when the liquid starts to freeze. Peltier cooled devices or devices TABLE 38.2 Cryogenic liquids and minimum temperature Chemical

Min. temperature (8C)

Comments

Liquid nitrogen

2196

Liquid argon

2186

n-pentane/liquid nitrogen slush Acetone/liquid nitrogen Ethanol/liquid nitrogen Acetone/dry ice

2130

Could accumulate oxygen and carbon dioxide Useful for air sampling since no risk that oxygen is accumulated (bp: 21838C). Carbon dioxide still preconcentrated, highly viscous, ﬂammable Flammable, limited CO2, low viscosity Flammable, high viscosity Fast and can be used for storage due to the ability to have a reservoir of dry ice in acetone Inexpensive, easy

Ice/salt mixture

1220

285 290 278

215

Sample preparation for the analysis of volatile metals species

which are cooled by liquid nitrogen and a temperature control unit were developed by Pecheyran et al. [8]. This method is particularly suited for lowmolecular weight species, which do not adsorb efﬁciently on coated or noncoated solids. Small amounts can easily be preconcentrated, so that the resulting detection limits can be extraordinarily small. In addition, degradation of the metal species is limited by the low temperature used, although the species are condensed to a small area in the tubing on which reaction (polymerization or oxidation) would take place at a faster rate than in the gas phase. The latter point makes this approach very suitable for volatile very reactive compounds such as trimethyl bismuth (Me3Bi). Thus, the low temperature collects the volatile metals and preserves the species against degradation. The sampling efﬁciency of the cryotrapping method can be enhanced by using an absorptive phase inside the cooled trap, e.g. PDMS coated on supporting material such as chromosorb. This allows lowering the vapor pressure of the collected compound considerable, such that the resulting sampling efﬁciency is strongly inﬂuenced. The major disadvantages are that, especially for real environmental samples, water is always present and is also cryogenically trapped, leading to the blockage of the sampling tube. Often, water removal columns have been employed to eradicate this problem. Solid CaCl2 is very effective and reacts quickly with water, while Mg(ClO4)2 is more efﬁcient but slower. Other possibilities include cryogenic water traps, which are cooled to a higher temperature than the collection tube, mostly 2 208C. Today, membrane tubes such as Naﬁonw are often used for drying gas streams [51]. These tubes consist of a membrane, which allows polar low-molecular weight molecules such as water to pass through into a dry nitrogen stream. However, especially reactive gases, such as volatile metals, can react with any scrubber. Although often used, no systematic recovery study on the use of Naﬁonw tubes for the analysis of particularly reactive VMCs in gas samples has been conducted. Namiesnik and Wardencki [52] have recently published a good review concerning removal of water vapor, discussing advantages and limitations of various desiccants, adsorbents, cryotrapping or permeation tubes for the analysis of VOCs. For gas samples containing signiﬁcant amounts of CO2, use of a cartridge ﬁlled with NaOH pellets is necessary in order to prevent blockage and the collection of CO2 in the cryotron when temperatures below 2 788C are used. While NaOH is very effective, care has to be taken since it has been shown that some VMCs, in particular SbH3, react irreversibly with the NaOH pellets, resulting in a low recovery (see Fig. 38.1 [53]). The choice of the technique and their parameters is dependent on the concentration of the metal species, their chemical nature and stability, as well as the physico-chemical matrix in which the species has to be analyzed. Additionally, the nature of the analytical procedure used following the sample preparation step is also signiﬁcant. It should be noted that, although some recipes for sampling and preconcentration of VMCs will be given, most

1221

J. Feldmann

Fig. 38.1. Recovery of volatile metal compounds using a cartridge ﬁlled with NaOH prior to cryotrapping in order to absorb CO2 from the gas sample. Data reproduced from Feldmann et al. [53].

parameters have to be optimized with the given set-up. This becomes even more evident when it is fully acknowledged that the performance of the sampling can even change during the sampling process, i.e. ﬂow rate of the sample, humidity, air temperature and CO2 content. For the different preconcentration methods, the following parameters have to be optimized for the speciﬁc analytical task: † † † †

ﬂow rate trapping temperature dimension of the trap and adsorbents (porosity, mesh size, etc.) adsorbents in the cryotraps

The type of analyte and its concentration, as well as the major constituents will determine the method used and their parameters.

38.5

SPECIFIC PROCEDURES

As noted above, the analytical procedure as a whole conclusive concept must match the sampling, sample preconcentration and instrumental analysis with the analytes’ properties. Several articles give a detailed overview of all possible methods described in the literature, under which some are fairly general whereas others are focused on light hydrocarbons [56–57]. Here, in this section, detailed descriptions are given for certain types of samples gathered from different sources in the literature.

1222

Sample preparation for the analysis of volatile metals species

38.5.1 Cryotrapping methods So far, the method mostly used for the sampling and preconcentration of VMCs has been cryotrapping. Feldmann et al. [9,14,17] as well as Pecheyran et al. [8,12], used a chromatographic packing (SP-2100 10% on Supelcoport (60/80 mesh) in a U-shaped glass tube immersed in liquid nitrogen as the cryogenic liquid, for the sampling of VMCs in human breath [58], in the headspace above a microbial culture [59], in landﬁll gas [9,14, 23], in sewage gas [17] and in urban air [12]. The cryotrapping served as the ﬁrst step within a packed column GC-ICP-MS method. The strategy was to stabilize the analytes, which are thermodynamically labile, as well as airand UV-sensitive. The cryotrapping tubes can easily be stored in liquid nitrogen and thus the sample is stabilized until the measurement is carried out. The transfer of analytes from the sampling point into an adequate analytical device must be achieved without loss, transformation or decomposition. Practical experience unfortunately highlights plugging problems when sampling atmospheres with high levels of humidity. Cryotrapping can be performed with or without adsorbents being used. Depending on the trapping temperature, the use of adsorbents may not be necessary, with the advantage that moderate volatilization temperatures can be applied, so that the labile VMCs do not suffer the risk of thermal degradation [60]. Of course, the choice of the trapping temperature has an intrinsic inﬂuence on the trapping efﬁciency. Pecheyran et al. [12] controlled the trapping temperature to 2 1758C during the sampling of urban air, thereby avoiding condensation of oxygen in the U-tubes (boiling point of oxygen is 2 1838C). Usually, a second cryotrap is needed (cryofocusing) in order to allow narrow bands to enter the GC columns and thus to enhance the resolution (Table 3).

38.5.1.1 Ambient air Large quantities of gases can be trapped in order to detect low concentrations of VMCs in urban air without blockage. The sampling set-up was described by Pecheyran et al. [12] and is shown in Fig. 38.2. Fifteen liters of gas are trapped at a ﬂow rate of 0.8 l/min through a 0.1 mm ﬁlter and a U-shaped drying tube cooled to 2 208C. The dried air is then sampled in a cooled trap ﬁlled with silanized glass wool and cooled down to 2 1758C. The tubes should be stored and transported to the lab under liquid nitrogen and then installed in a GC equipped with a cryofocusing device (column cooled to 2 1968C). The cryotrapped gas sample is heated electrothermally and ﬂushed by a helium ﬂow to an element-speciﬁc detector such as ICP-MS or AAS. The focussed analytes can be separated by gas chromatography and detected either by electron impact mass spectrometry or ICP-MS.

1223

1224 TABLE 38.3 Matrix dependent temperature for cryotrapping methods Main components

Trapping temperature

Trapping conditions

Reference

Landﬁll gas, sewage gas

CO2, CH4, saturated water

2808C or with CO2 trap (NaOH) at 21968C possible

Water trap (CaCl2, Mg(ClO4)2 or second pre-cryotrap at 2208C, and a CO2 trap (NaOH).

[9,14,17]

Landﬁll gas Natural gas Ambient air Incineration gas

CH4 Water, small amount of CO2 High amounts of water

21308C 21758C 2808C

Water sample

Water

21968C, with water trap

Working place air

Air, water, small amounts of CO2 (,0.1%)

21758C

No water or CO2 trap necessary 2208C water trap High capacity water traps are necessary (pre-cryotrap) Stripping vessel with water trap (K2CO3) and cryotap Due to low concentration, large volume sampling necessary, therefore, water trap (2208C)

[2] [67] [12] [37] [13] [8]

J. Feldmann

Sample

Sample preparation for the analysis of volatile metals species

Fig. 38.2. Set-up for the cryotrapping of analytes from ambient air according to Pecheyran et al. [8].

38.5.1.2 Landﬁll/sewage gas Since these gases contain large amounts of volatile organics as wellas high levels of VMCs, there are two types of sample preparation: one for the identiﬁcation of very volatile species, and one for semi-volatile species. (a) Very volatile species: The gases can be sampled directly from the gas wells into a Tedlar bag. The Tedlar bags, wrapped in a black bag to prevent decomposition by UV light, are transported to the lab and sub-samples taken using a gas-tight syringe. Between 10 –100 ml of gas can be injected through a cartridge ﬁlled with fresh NaOH pellets. The outlet of the cartridge is connected to the capillary cooled to 21968C. With a ﬂow rate of approximately 100 ml/min, good recovery of very volatile species such as AsH3 and SnH4 are guaranteed without any disturbance of the detector. (b) Semi-volatile species: The cryotrap can be cooled only to 2788C, at which point only a limited amount of CO2 is pre-concentrated. At a ﬂow rate of more than approximately 500 ml/min, the recovery of compounds having a boiling point of þ358C is quantitative. With this technique, CaCl2 or Mg(ClO4)2 can be substituted for the water trap (2208C) by in order to dry the water-saturated gas sample. A cryotrap ﬁlled with Chromosorb SP2100 (60/80 mesh) can separate volatile compounds very crudely by externally heating to 1808C in a helium ﬂow of 130 ml/min. If this column is connected to a syringe needle and a fraction collector equipped with 15 ml evacuated vials, the different gas fractions can be collected and individually injected into a capillary column with different properties. Thus, the gas samples are separated in an orthogonal manner,

1225

1226 J. Feldmann Fig. 38.3. Set-up for the identiﬁcation of VMCs in a very complex matrix (sewage and landﬁll gas). 2-dimensional GC separation and detection by ICP-MS and EI-MS. Taken from Ref. [9].

Sample preparation for the analysis of volatile metals species

which leads to cleaner samples. This makes the identiﬁcation of VMCs using EI-MS possible (Fig. 38.3 [9]). 38.5.1.3 Water samples VMCs dissolved in water can be determined by sampling the water into a vessel equipped with a frit at the bottom in order to purge the water with a low ﬂow of helium gas (700 ml/min). The gas is dried by a cooled U-trap (2 208C) [16] or by using K2CO3 [61] and then cryotrapped. The set shown in Fig. 38.4 is in accord with Amouroux et al. [16]. 38.5.1.4 Incineration gas Flue gas from incinerators cannot be directly cryotrapped since their temperature of about 2008C and above would have a negative inﬂuence on the trapping efﬁciency of the system used for CO2 rich atmospheres. Figure 38.5 shows a sampling unit used for the sampling of ﬂue gas from a coal-ﬁred power plant in which the gases were cooled prior to the water trap and the cryotrap [62]. 38.5.2 Solid phase microextraction A promising approach as a new sample introduction method for volatile metal species has been described only recently by Mester et al. [63] and Smith et al. [64]. Solid phase microextraction (SPME) is directly coupled to an ICP-MS using a thermal desorption interface directly at the base of the torch. Direct immersion and headspace extraction was studied for methylmercury providing a limit of detection of 0.19 ng/ml for the headspacesampling version, which should be applicable to landﬁll gas samples as well. This method is very convenient and does not need any cryogenic liquid. The disadvantage, however, is that gases with high levels of organic trace components, such as landﬁll gas, cannot be analyzed for VMCs due to competitive reactions on the solid phase adsorbent and the amount of gas cannot be varied. 38.5.3 Adsorption method 38.5.3.1 Carbosieve method Metal compounds present in the atmospheric, such as Hg0, MeHgCl and Me2Hg, can be collected on Carbosieve in order to separate these species from CO2 and water. When the Carbosieve is heated up to 4008C, the desorbed species can be cryotrapped again and subsequently determined by GC-AED or AFS [13]. 38.5.3.2 Tenax/Porapak method A ﬁne example of the applicability of an adsorbtion method is the use of a combination of Tenax and Porapak for the trapping of organolead compounds in urban air. According to Nerin and Pons [40], glass tubes (11 cm £ 0.6 cm) ﬁlled with Tenax and Porapak (35:65, w/w) can be used in series in order to collect

1227

1228 J. Feldmann

Fig. 38.4. Purge and trap GC set-up for determination of dissolved VMCs in water samples Amouroux et al. [16].

Sample preparation for the analysis of volatile metals species

Fig. 38.5. Set-up for the sampling of ﬂue or stack gas [36].

volatile and semi-volatile species at the same time. With a ﬂow rate of 1 l/min, up to 70 l of air can be trapped. The organolead compounds are extracted with 4 ml hexane with the aid of an ultrasonic bath. After reducing the ﬁnal volume under a nitrogen ﬂux, the crude extract was injected into a GC-MS. Due to the abundance of VOCs, the determination of organolead species is difﬁcult and could be easier if AAS or ICP-MS was used as the lead-speciﬁc detector (Table 4).

38.6

PROBLEMS AND FUTURE STUDIES

One of the major problems is the clear identiﬁcation of trace amounts of volatile organometallic compounds in very complex matrices. On the one hand, it is TABLE 38.4 Application of different adsorption methods Species

Matrix

Sample preparation

Analytical method

Reference

AsH3/SbH3/PH3

Workplace air

Total metal

[68]]

PbR4

Urban air

GC-MS

[40]

Me3Sb

Aerobic fungi cultures

Adsorption on AgNO3 impregnated pads Adsorption on Porapak/Tenax, hexane Anaerobic trapping on Tenax

GC-MS and GC-AAS

[42]

1229

J. Feldmann

necessary to use an element-speciﬁc detector in order to identify VMCs in a cocktail of more abundant VOCs. On the other hand, if the speciation method includes a chromatographic separation and is linked to a plasma source mass spectrometer, molecular information is lost. If the ionization is gentler, in order to retrieve molecular information using electron impact or chemical ionization methods, the molecular information can be retrieved. Due to the occurrence of a complex matrix of dozens or hundreds of compounds, high performance chromatographic separation is essential. Here, more innovative approaches are necessary to separate the mostly unstable VMCs from the more abundant VOCs in process gases or urban gas samples. Problems relating to the formation of artifacts during the sample preconcentration and manipulation have not been extensively studied, and hence very limited information is available. It is expected that experiments with isotopically labeled compounds will shed some light on this area in the near future.

REFERENCES 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23

1230

M. Stoeppler, in: M. Stoeppler (Ed.), Sampling and Sample Preparation: A Practical Guide for Analytical Chemists. Springer, Berlin, 1997. M. Schweigkoﬂer and R. Niessner, Environ. Sci. Technol., 33 (1999) 3680. Y. Wang, T.S. Raihala, A.P. Jackman and R.S. John, Environ. Sci. Technol., 30 (1996) 3115. B. Gosio, Science, 19 (1892) 104. B. Gosio, Arch. Ital. Biol. Science, 18 (1893) 253. F. Challenger, C. Higginbottom and L. Ellis, J. Chem. Soc., (1933) 95. F. Challenger, Chem. Rev., 36 (1945) 315. C. Pecheyran, C.R. Quetel, F.M.M. Lecuyer and O.F.X. Donard, Anal. Chem., 70 (1998) 2639. J. Feldmann, I. Koch and W.R. Cullen, Analyst, 123 (1998) 815. A.V. Hirner, J. Feldmann, E.M. Krupp, R. Gru¨mping, R. Goguel and W.R. Cullen, Org. Geochem., 29 (1998) 1765. R. Pongratz and K.G. Heumann, Chemosphere, 39 (1999) 89. C. Pecheyran, B. Lalere and O.F.X. Donard, Environ. Sci. Technol., 34 (2000) 27. R. Pongratz and K.G. Heumann, Chemosphere, 36 (1998) 1935. J. Feldmann and W.R. Cullen, Environ. Sci. Technol., 31 (1997) 2125. A.V. Hirner, U.M. Gru¨ter and J. Kresimon, Fresenius J. Anal. Chem., 368 (2000) 263. D. Amouroux, E. Tessier, C. Pecheyran and O.F.X. Donard, Anal. Chim. Acta, 377 (1998) 241. J. Feldmann, J. Environ. Monit., 1 (1999) 33. Z. Mester and R.E. Sturgeon, J. Anal. At. Spectrom., 16 (2001) 470. L. Guo, W.A. Jury and W.T. Frankenberger, J. Contam. Hydrol., 49 (2001) 67. U. Telgheder and V.A. Khvostikov, J. Anal. At. Spectrom., 12 (1997) 1. S. Signorelli, A. Bucciani, M. Martini and G. Piccardi, Geochem. J., 32 (1998) 367. S. Signorelli, Environ. Geol., 32 (1997) 239. J. Feldmann and A.V. Hirner, Int. J. Environ. Anal. Chem., 60 (1995) 339.

Sample preparation for the analysis of volatile metals species 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59

B. Bouyssiere, F. Baco, L. Savary, H. Garraud, D.L. Gallup and R. Lobinski, J. Anal. At. Spectrom., 16 (2001) 1329. K. Haas and J. Feldmann, Anal. Chem., 72 (2000) 4205. K. Haas, J. Feldmann, R. Wennrich and K.J. Sta¨rk, Fresenius J. Anal. Chem., 370 (2001) 587. J.C. Pau, J.E. Knoll and M.R. Midgett, J. Air Waste Mgmt Assoc., 41 (1991) 1095. Y. Hoshika, M. Nishikitani, K. Yokoyama and S. Araki, Anal. Sci., 13 (1997) 505. J.L. Fugit, L. Dutaur, V. Simon, M.L. Riba and L. Torres, Fresenius Environ. Bull., 5 (1996) 682. G.F. Evans, T.A. Lumpkin, D.L. Smith and M.C. Somerville, J. Air Waste Mgmt Assoc., 42 (1992) 1319. F. Lipari, J. Chromatogr., 503 (1990) 51. J.M. Andino and J.W. Butler, Environ. Sci. Technol., 25 (1991) 1644. E.D. Pellizzari, J.E. Bunch, B.H. Carpenter and E. Sawicki, Environ. Sci. Technol., 9 (1975) 552. M. Ito and D. Shooter, Atmos. Environ., 36 (2002) 1499. L. Filkova and J. Ja¨ger, C. Hygenia, 31 (1986) 255. B. Pedersen, Ann. Occup. Hyg., 32 (1988) 385. M.P. Pavageau, C. Pecheyran, E.M. Krupp, A. Morin and O.F.X. Donard, Environ. Sci. Technol., (2003) in press. R. Falter, Chemosphere, 39 (1999) 1037. X.L. Cao and C.N. Hewitt, J. Chromatogr. A, 688 (1994) 368. C. Nerin and B. Pons, Appl. Organmet. Chem., 8 (1994) 607. Y. Wang, A.B. Turnbull and R.M. Harrison, Appl. Organomet. Chem., 11 (1997) 889. R. Jenkins, P.J. Craig, W. Goessler, D. Miller, N. Ostah and K.J. Irgolic, Environ. Sci. Technol., 32 (1998) 882. W.A. McClenny and M. Colon, J. Chromatogr. A, 813 (1998) 101. J. Sommar, X.B. Feng and O. Lindqvist, Appl. Organomet. Chem., 13 (1999) 441. M.J. Chang, R.L. McDaniel, J.D. Naworal and D.A. Self, J. Anal. At. Spectrom., 17 (2002) 710. N.S. Bloom, E.M. Prestbo, B. Hall and E.J. van der Geest, Water, Air Soil Pollut., 80 (1995) 1315. S.E. Lindberg, D. Wallschla¨ger, E.M. Prestbo, N.S. Bloom, J. Price and D. Reinhart, Atmos. Environ., 35 (2001) 4011. W.H. Schroeder and R.A. Jackson, Int. J. Environ. Anal. Chem., 22 (1985) 1. W.R. Cullen and K.J. Reimer, Chem. Rev., 89 (1989) 713. O.F.X. Donard: in Chapter 16 N.G. Sundin, J.F. Tyson, C.P. Hanna and S.A. McIntosh, Spectrochim. Acta B, 50 (1995) 369. J. Namiesnik and K. Wardencki, Int. J. Environ. Anal. Chem., 73 (1999) 269. J. Feldmann, L. Nae¨ls and K. Haas, J. Anal. At. Spectrom., 16 (2001) 1040. J. Dewulf and H. Van Langenhove, J. Chromatogr. A, 843 (1999) 163. V. Camel and M. Caude, J. Chromatogr. A, 710 (1995) 3. D. Helmig, J. Chromatogr. A, 843 (1999) 129. K. Destombe, D.K. Verma, L. Stewart and E.B. Reczek, Am. Ind. Hyg. Assoc. J., 52 (1991) 136. J. Feldmann, T. Riechmann and A.V. Hirner, Fresenius J. Anal. Chem., 354 (1996) 620. P. Andrewes, W.R. Cullen, J. Feldmann, I. Koch, E. Polishuchuk and K.J. Reimer, Appl. Organomet. Chem., 12 (1998) 827.

1231

J. Feldmann 60 61 62 63 64 65 66 67 68

1232

N. Schmidbauer, J. High Resolut. Chromatogr., 8 (1985) 404. R. Pongratz and K.G. Heumann, Int. J. Environ. Anal. Chem., 71 (1998) 41. J. Feldmann, Abstract, Fourth International Nickel Symposium, Murmansk, Russia, 2002 Z. Mester, J. Lam, R. Sturgeon and J. Pawliszyn, J. Anal. At. Spectrom., 15 (2000) 837. L.M. Smith, W.A. Maher, P.J. Craig and R.O. Jenkins, Appl. Organmet. Chem., 16 (2002) 287. In: D.R. Lide (Ed.), Handbook of Chemistry and Physics, 72nd edn. CRC Press, Boca Raton, FL, 1992. B.I. Tel’noi and I.B. Rabinovich, Usp. Khim., 49 (1980) 1137. J. Feldmann, PhD Thesis, University of Essen, 1995 S. Hetland, I. Martinsen, B. Radziuk and Y. Thomassen, Anal. Sci., 7 (1991) 1029.

Chapter 39

Sequential extraction Angels Sahuquillo and Gemma Rauret

39.1

SEQUENTIAL EXTRACTION PROCEDURES: A SPECIAL CASE OF SAMPLE DISSOLUTION

39.1.1 A brief historical introduction Due to their effects on the environment, trace elements, especially heavy metals, are considered one of the main sources of pollution. Consequently, an increasing number of environmental samples are submitted to analysis in the frame of routine monitoring, and both risk and sustainability assessment studies are continually growing. For each type of matrix, the ﬁrst step of any monitoring action is to determine whether the total metal content lies within the range of background levels or above certain concentration limits established by international (e.g. European directives) or national legislation. The most common present-day methods used for the determination of heavy metal amounts in environmental samples involve highly sensitive spectroscopic techniques, such as atomic absorption spectroscopy with ﬂame or graphite furnace (F-AAS, GF-AAS) and inductively coupled plasma-optical emission and mass spectrometry (ICP-OES and ICP-MS). These techniques usually require the dissolution of the solid sample (sediment, soil, sewage sludge, ﬂy ash, waste, etc.) prior to metal content determination. Sample digestion is mainly accomplished using wet procedures based on an acid digestion, which have been widely discussed in Chapters 6 and 8 in this volume. The determination of total metal content is conducted with the aim of detecting polluted sites and target elements to be further investigated. This approach is on its way to becoming one more step available for the complete characterisation of a given matrix, together with other parameters, such as organic matter content, elemental analysis (C, H, N), major compounds, etc. At present, it is widely accepted that the total metal content is not sufﬁcient to obtain relevant information for environmental studies that are aimed at the evaluation of the impact either of anthropogenic or of natural contamination Comprehensive Analytical Chemistry XLI Mester and Sturgeon (Eds.) q 2003 Elsevier B.V. All rights reserved

1233

A. Sahuquillo and G. Rauret

sources, the assessment of potential element mobility and bioavailability. As stated by Kersten [1], the biogeochemical and ecotoxicological signiﬁcance of a pollutant is determined by its speciﬁc binding form and coupled reactivity rather than by its accumulation rate. Not all forms of a given metal have an equal impact on the environment; thus, the total concentration cannot be used as the only criterion to assess the potential effects of a pollutant from a sediment, soil or waste sample. In this sense, the use of partial dissolution techniques arose in the ﬁeld of trace element geochemical prospecting studies, as described by Pickering [2]. The aim is to identify anomalies, i.e. rocks, soils or sediments differing from expected background values. The analytical techniques used for this purpose should raise contrasting properties in order to enhance the potential of identifying a signiﬁcant geochemical pattern. With this aim, the so-called sequential selective extraction procedures started to become widely used. Tessier [3] highlighted the valuable information provided by these procedures in understanding the chemical and physical processes involved in the weathering of ore deposits from different experiments carried out since the late 1970s.

39.1.2 Deﬁnition of sequential extraction procedures The basics of the sequential extraction procedures (SEPs) involve the application of different reagents of increasing dissolution power in a sequential way, i.e. using the residue obtained from the previous step. Here, the aim is to understand the element distribution in the solid phase. Generally, from 3 to 8 extractants are used in a sequence where the earlier reagents are less aggressive and the most speciﬁc, the subsequent extractants being progressively more destructive and less speciﬁc. These procedures are intended to be a methodological approach for estimating the fractionation of trace elements according to the International Union of Pure and Applied Chemistry (IUPAC) deﬁnition [4]. The term fractionation is deﬁned as the process of classiﬁcation of an analyte or group of analytes from a certain sample according to physical (e.g. size, solubility) or chemical (e.g. bonding, reactivity) properties. Notably, there is a difference between these approaches and the term speciation in the sense of determination of chemical species, such as, e.g. Cr(III) and Cr(VI) speciation. However, the literature does not show uniformity of nomenclature and these approaches are referred to as chemical speciation procedures, speciation schemes, sequential chemical extractions, sequential extraction schemes, sequential analytical procedures, fractionation procedures and partitioning schemes. Here, we will refer to these methods as SEPs.

1234

Sequential extraction

39.2

TYPES, USES AND LIMITATIONS OF SEPs

39.2.1 Characteristics of the extraction agents As is well known, many different reagents are used as extractants in the existing SEPs. Many of them are chosen as representatives of the different processes than can induce the release of trace elements from an environmental solid sample, such as, e.g. salt-level effects, acidity changes, modiﬁcation in system reduction potentials or complex formation. Each of the extractants used leads to a liquid fraction wherein trace elements are measured. Each fraction tends to be associated with a solid phase and is correlated to possible changes of environmental conditions (pH, redox potential, etc.). Usually, the SEPs describe water soluble, weakly adsorbed and exchangeable fractions, weak acid soluble fractions or fractions bound to carbonates, easily reducible fractions or those bound to iron and manganese oxides, oxidisable fraction or fractions where trace elements are bound to organic matter and sulﬁdes, as well as residual fractions incorporated in the mineral phase. The principal extraction agents used in SEPs have been exhaustively reviewed and are summarised in Table 39.1. 39.2.2 SEPs more widely used There are many different SEPs that usually combine one extractant of each of the metal fractions indicated in Table 39.1. In some cases, the SEPs distinguish single fraction substances, such as easily reducible and moderately reducible oxides of Fe and Mn. In these cases, the SEPs can comprise up to eight steps. Moreover, the experimental conditions may require several days of work in the laboratory and lead to procedures that are too complicated to be applied routinely. On the other hand, there are shorter procedures wherein a few reagents of different chemical properties (acidic, reducing, oxidising) are selected and lead to schemes with up to three or four steps. In favour of the short SEPs is the fact that an increase in the number of steps implies further manipulation of the sample, and can have an effect on the overall precision of the applied procedure. This takes on even greater importance considering that the relative standard deviation will already be about 2–8% for short procedures in soils and sediments. It is beyond the scope of this chapter to give an overview of the existing SEPs, but two different schemes have been selected as representative of long and short ones. Nevertheless, there are obviously many different schemes, as can be seen from the possibilities of combining the extractant reagents summarised in Table 39.1. The classical ﬁve-step Tessier method [3,7,8] has been widely applied. The reagents used in this scheme and the fractions selected for partitioning of trace

1235

1236

TABLE 39.1 Extraction agents and associated solid phases in SEPs (from [2,5,6]) Metal fractions

Type of extractant

Extractants used

Water soluble fraction

Water (distilled or deionised)

Pore water or H2O extraction

Exchangeable and weakly adsorbed fraction

Salts of strong acids and bases or salts of weak acids

KNO3 1 M (pH 7) Mg(NO3)2 0.5 –1 M (pH 7)

Acid extractable or carbonate-bound fraction

Acid or buffer solutions

CH3COOH 25% or 1 M NaCH3COO 1 M/CH3COOH (pH 5) HCl or CH3COOH (unbuffered solutions) EDTA 0.2 M (pH 10– 12)

Reducing extractable or fractions bound to oxides of Fe, Mn and Al

Reducing solutions Other agents

0.2% hydroquinone in NH4CH3COO 1 M (pH 7) NH2OH·HCl 0.02– 1 M in CH3COOH or HNO3 (NH4)2C2O4 0.2 M (pH 3) (NH4)2C2O4 0.2 M/H2C2O4 0.2 M in ascorbic acid 0.1 M Na2S2O4/Na-citrate/citric acid Na2S2O4/Na-citrate/NaHCO3 (pH 7.3) Na2S2O4/K4P2O7

A. Sahuquillo and G. Rauret

CaCl2 0.01– 0.05 M MgCl2 or BaCl2 1 M (pH 7) NaCH3COO or NH4CH3COO 0.1– 1 M (pH 7 or 8.2)

TABLE 39.1 (continuation) Metal fractions

Type of extractant

Extractants used H2O2 10% in 0.0001 N HNO3 HCl 20 % EDTA 0.02–0.1 M (pH 8–10.5) Na2EDTA (buffered with NH4CH3COO 1 M) Hydrazine chloride (pH 4.5)

Oxidising reagents

H2O2 in HNO3 þ extraction with NH4CH3COO or MgCl2 NaClO (pH 9.5) Alkalipyrophosphate (Na4P2O7 or K4P2O7 0.1 M) H2O2/ascorbic acid HNO3/tartaric acid KClO3/HCl NaOH 0.1 M (pH 9.5) Na2B4O7 (addition of surfactant)

Residual fraction

Strong acids

Alkaline fusion HF/HClO4/HNO3 (different mixtures) Aqua regia HNO3/H2O2 HCl/HF/HNO3

Sequential extraction

Oxidising extractable or organically bound and sulﬁdic phase

1237

1238

S EDIMENT (1 g)

S EDIMENT (1 g)

STEP S TEP 1: 1: exchangeable e xcha nge a ble M (pH (pH == 7) 7) agent : Extracting agent: MgCl2211 M Extraction: m/v == 1:8; 1:8; 11 hh Extraction m/v Ce ntrifuga tion a nd wa s hing

S TEP 11 STEP Extracting Extra cting agent: a ge nt: HCH3COO 0.11 M Extraction: m/v Extra ction: m/v== 1:40; 1:40;16 16 hh

EXTRACT S TEP 1

RES IDUE S TEP 1

Ce ntrifuga tion a nd wa s hing

STEP S TEP 2: 2: bound bound to to carbonates ca rbona te s Extracting agent: Na CH3COO 1 M (pH5, HCH3COO) agent: Extraction Extraction: m/v = 1: 8 ; 5 h EXTRACT S TEP 2

RES IDUE S TEP 2

RES IDUE S TEP 1

Extracting agent: Extraction:

Ce ntrifuga tion a nd wa s hing

S TEP 3: STEP 3: bound bound to to Fe-Mn Fe-Mn oxides oxide s Extracting Extractingagent: agent: NH2OH.HCl 0.04 M (in 25 % HCH 3COO) Extraction: Extraction: m/v m/v == 1: 1: 20 20 ;; 66 hh aatt 96 96 ±± 33 ºC ºC Ce ntrifuga tion a nd wa s hing

S TEP 22 STEP NH22OH.HCl 0.5 M (pH 1.5, HNO 33 2 M fixe d vol.) m/v = 1:40; 16 h EXTRACT S TEP 2

RES IDUE S TEP 2

EXTRACT S TEP 3

RES IDUE S TEP 3 STEP 4: bound to organic matter H2O2 30 %; (2+3) h with (8+3) ml a t 85˚C Digestion: Extracting agent: NH4CH3COO 3.2 M (in 20 % HNO 3conc) Extraction: m/v = 1:5; 30 min Ce ntrifuga tion a nd wa s hing

EXTRACT S TEP 4

FINALRES FINAL RESIDUE IDUE HF:HClO HF:HClO445:1 5:1 (to (to dryne drynessss sseeve vera rall time timess)) A

Digestion: Extracting agent: Extraction:

S TEP 33 STEP H2O2 30 % (Room T a nd 85˚C) 10ml two time s NH4CH3COO 1 M (pH 2, HNO3 conc) m/v = 1:50; 16 h

Ce ntrifuga tion a nd wa s hing

EXTRACT S TEP 3

FINAL RES FINAL RESIDUE IDUE Aqua regia Aqua re gia dige diges stion tion 16 h room T + 2 h re flux 130 ºC B

Fig. 39.1. Experimental conditions for: (A) Tessier’s procedure (ﬁve steps); (B) modiﬁed BCR procedure (four steps) (m/v: mass/volume ratio).

A. Sahuquillo and G. Rauret

Ce ntrifuga tion a nd wa s hing

EXTRACT S TEP 1

Sequential extraction

metals affected by different environmental conditions are the following: MgCl2 1 M (pH 7) (exchangeable); NaCH3COO 1 M/HCH3COO (pH 5) (bound to carbonates); NH2OH·HCl 0.04 M in 25% HCH3COO (bound to Fe–Mn oxides), H2O2 8.8 M at pH 2 with HNO3 0.02 M and subsequent extraction with NH4CH3COO 3.2 M in HNO3 (bound to organic matter); a mixture of HF and HClO4 (residual phase). Under these experimental conditions, relative standard deviations of 10% or lower were typically obtained with sediments for trace metal concentrations amounting to ﬁve times or more the detection limit. A scheme outlining the experimental conditions for this SEP is shown in Fig. 39.1A. Different modiﬁcations to this procedure have been proposed in order to improve the lack of selectivity of the extraction agents towards the speciﬁc geochemical phases of the considered sediment [9,10]. Since its proposal in 1993, the European three-step procedure developed in the framework of the Standards, Measurement and Testing Programme of the European Commission (former BCR) has been also widely used. This SEP is a consensus scheme, within a group of leading European soil and sediment laboratories. It has been established with the aim of harmonising the existing SEPs in order to be able to compare data among laboratories and to provide proper certiﬁed reference materials (CRMs) for fractionation studies. The original BCR scheme was improved in 1998 after an exhaustive study of possible sources of uncertainty [11,12] using an existing CRM, BCR 601, a lake sediment, as a test sample. The reagents proposed in the modiﬁed version of the scheme are the following: HCH3COO 0.11 M (mainly carbonatic); NH2OH·HCl 0.5 M in HNO3 2 M (mainly oxides Fe/Mn); H2O2 30% and NH4CH3COO 1 M (pH 2) (mainly organic matter and sulﬁdes); a ﬁnal aqua regia digestion is recommended for estimating the residual phase. The experimental conditions used for this SEP are shown in Fig. 39.1B.

39.2.3 Types of matrices and elements analysed A survey of 140 published papers covering the period 1979 –2002 dealing with SEPs has been completed. The journals overviewed include the analytical chemistry and geological and environmental ﬁelds: Analytical Chemistry, Trends in Analytical Chemistry, The Analyst, Analytica Chimica Acta, Talanta, Fresenius Journal of Analytical Chemistry, Analytical and Bioanalytical Chemistry, Annali di Chimica, Spectrochimica Acta, Analytical Communications, International Journal of Environmental Analytical Chemistry, Microchemical Journal, Pure and Applied Chemistry, Journal of Analytical Atomic Spectrometry, Journal of Geochemical Exploration, Geochimica and Cosmochimica Acta, Ore Geology Reviews, Chemical Geology, Applied Geochemistry, Marine Geology, The Science of the Total Environment, Environmental Science

1239

A. Sahuquillo and G. Rauret

and Technology, Environmental Pollution, Environment International, Journal of Environmental Quality, Journal of Environmental Monitoring, Journal of Soils and Sediments, Communications on Soil Science and Plant Analysis, Marine Chemistry, Hydrobiologia, Environmental Geology and Water Science, Water Research, Water, Air and Soil Pollution, Netherlands Journal of Sea Research, Chemosphere, Water and Science Technology and Waste Management. The type of SEPs described in the papers considered is mainly three-, fourand ﬁve-step procedures; the longer procedures of up to eight steps are rarely used. Most of the methods are based on Tessier’s scheme and different modiﬁed versions of Gibbs or Fo¨rstner, and the BCR procedure in its original and modiﬁed versions. As far as the type of matrices studied is concerned, the survey has covered the distribution illustrated in Fig. 39.2. As can be seen, the principal subject of these papers (about 60%) refers to sediment studies, as this was the ﬁrst matrix that SEPs were specially designed for. In fact, all papers mentioned in the present review, until 1987, refer solely to sediment samples. Different origins have been described for these samples: lake, river, estuary, marine, lagoon and road sediments. Both bottom and surface samples have been studied, and depth proﬁles of sediments are also described for diagenesis studies. The second matrix where SEPs are widely applied is soils, their main provenance being polluted areas hosting industrial and mining activities, although agricultural and forest soils are also studied. With the generic name of solid wastes, different samples have been considered: ﬂy and bottom ashes from municipal incineration plants, municipal compost, different wastes from mining areas, road dust and blast furnace sludges.

7%

2%

5%

Sediments Soils Sewage sludges 27% 59%

Solid wastes Particulate matter

Fig. 39.2. Distribution of analysed matrices using fractionation procedures (1979– 2002).

1240

Sequential extraction

Other sample types include sewage sludges coming from urban wastewater treatment plants. A few studies on particulate matter have also been also found. The distribution obtained for the elements analysed in the reviewed environmental samples is shown in Fig. 39.3. Cadmium, Cr, Cu, Ni, Pb and Zn are the elements of most environmental concern and are studied in 57% of the reports. This is due to the fact that the procedures under discussion were designed for heavy metals, occurring mainly in cationic forms. In most cases, the information presented for these trace elements is also complemented by that obtained for some matrix elements such as Si, Al, Ti, Fe, Mn, Ca, Mg and K. There are some data available for other elements, such as Sc, V, Co, Y, Mo and Ag, and some speciﬁc studies deal with As and Hg. SEPs are in few cases applied to lanthanides and actinides, such as La, Ce, Th and U.

39.2.4 Use of the information obtained Even if SEPs have limitations, as will be discussed in Section 39.2.5, the information provided is considered highly useful in many ﬁelds of application. Chemical fractionation was mainly applied, in its early years of use in the ﬁeld of environmental geochemistry, to obtain distribution patterns of trace metals among speciﬁc geochemical phases. As the leachability of a metal from a certain sample will depend on the strength of its bonding, the identiﬁcation of geochemical phases, i.e. its different levels of bonding, can be related to the risk assessment of the presence of the metal in the environment. SEPs are valuable tools for providing information in this sense. In this case, only samples from sites with high heavy metal contents identiﬁed as a potential sources of

9%

1%

7%

26%

57%

Cd, Cr, Cu, Ni, Pb, Zn Major compounds As, Hg Other elements Lanthanids,actinids

Fig. 39.3. Distribution of analysed elements using fractionation procedures (1979– 2002).

1241

A. Sahuquillo and G. Rauret

pollutants are good candidates for use in further fractionation studies. As stated by Perin [13], a polluted sediment releasing less than 1% of total metal content in the ﬁrst steps of a SEP (exchangeable and easily extractable phases) can be considered safe for the environment, whereas extractable amounts higher than 50% in the same phases can be considered as highly hazardous. Estimation of the remobilisation of metals under changing environmental conditions and of the potential uptake by biota are two of the main aims of heavy metal fractionation on particle-bound trace metals. Direct relationships of empirical studies applying SEPs with bioavailability are highlighted as difﬁcult by different authors [14], due to the fact that many complex physical, chemical and biological processes are involved in the partitioning of trace metals in sediments and also to a lack of knowledge of the speciﬁc mechanisms by which organisms actively translocate trace element species [9]. Although the fractions obtained by applying SEPs are not connected directly with bioavailability, the provided information is useful to decision makers when it comes to assessing pollutant mobility at times of changing environmental conditions in different ﬁelds, such as waste management and agricultural practices. For example, SEPs are applied to bottom [15] and ﬂy ashes [16] from incineration of municipal wastes prior to being used in road construction or stored in monoﬁlls or in establishing proper procedures for treatment of wastes. The amendment of agricultural soils with sewage sludges and composts is a usual practice because these waste substances are a source of nutrients for plants. The application of SEPs in these samples aims at assessing the mobility of trace elements prior to soil disposal as fertilisers [17] and also at studying the effect on mobility of different disposal alternatives [18]. 39.2.5 Limitations of SEPs 39.2.5.1 Lack of selectivity The importance of the use of the right terms when speaking about fractionation is linked to the noted lack of selectivity and speciﬁcity of the reagents used in the SEPs. There are many references in the literature dealing with the selectivity problems of the reagents used in different SEPs. The unsatisfactory reproducibility obtained in exchangeable and organic fractions after the application of Tessier’s procedure was attributed by Accomasso [19] to the insufﬁcient selectivity of the extraction. Nirel [20] pointed out that different mineralogical compositions lead to different efﬁciencies of extraction. The geochemical phase speciﬁcity of the original BCR-SEP has been shown by Whalley [21] to be of varying quality on single substrates. For this procedure, sufﬁcient repeatability and reproducibility are reported for their application in fractionation studies [22,23].

1242

Sequential extraction

Furthermore, an evaluation of the selectivity of SEPs on Cd fractionation in soils [24] shows that the selectivity of extraction depends not only on the type of reagents used to extract each phase, but also on the order in which they are applied. Figure 39.4 shows the extractable amounts of Ca obtained following the modiﬁed BCR procedure applied to samples having different CaCO3 content: harbour sediment (18.3%), river sediment (14.2%), soil A (8.6%) and soil B (4%). As can be seen, extractable Ca in step 1 ranges from 68 to 92% of the total extractable amount but the extractability does not correlate with the total CaCO3 content. Moreover, signiﬁcant Ca amounts are still obtained in step 2 of the SEP, ranging from 10 to 20% of the total extractable amounts. Thus, the extractable amounts obtained by using a weak acid (HCH3COO 0.11 M) cannot be directly correlated with trace metals bound to carbonates, as was supposed in the early publications.

39.2.5.2 Operational deﬁnition SEPs have to be considered an operationally dependent methodology, as derived from their design. There are many factors affecting the extractability of metals following a SEP: type and reagent conditions (concentration, pH, acid used for adjusting pH), type and time of shaking, mass/volume ratio, extraction temperature, etc. [11].

Extractable Ca amount

100%

80%

60%

40%

20%

0% Harbour sediment`

River sediment Step 1

Soil A Step 2

Soil B

Step3

Fig. 39.4. Extractable Ca amounts following modiﬁed BCR-SEP in sediments and soils.

1243

A. Sahuquillo and G. Rauret

As an example, Cd, Cr and Cu extractable amounts in the second step following the original BCR procedure in a sediment are shown in Fig. 39.5 as a function of pH and the acid used in adjusting it. As can be seen, the observed effects depend on the elements considered. Whereas for the most mobile element (Cd), neither of the variables has a signiﬁcant effect on extractable amounts, for Cu and Cr an increase of 0.5 units of pH yields an increase of extractability of 70–80 and 15–25%, respectively. The use of HCl instead of HNO3 for adjusting the pH in the extraction agent resulted in an increase in the extractability of around 1.5– 3% for Cr and Cu. This fact makes comparison of results arising from different SEPs especially difﬁcult and justiﬁes efforts towards standardisation. The strategies followed by the Standards, Measurement and Testing Programme of the European Commission are very valuable in this sense [25,26], including the validation aspects of these procedures by providing suitable CRMs [27–29]. The operational deﬁnition of the procedures makes it necessary to keep in mind that the association of extracted fractions to mineralogical phases, such as carbonates, Fe and Mn oxides, cannot be interpreted as an absolute concept [6,19]. So, it is more accurate to speak of extracted trace elements in a certain reagent, following the given conditions of the SEP. 39.2.5.3 Readsorption and redistribution The importance of post-extraction readsorption of elements has been widely studied and discussed by some authors with notable controversy. For instance,

9,0 Cu (: 5)

8,0

mg.kg–1 (dry mass)

7,0 6,0 5,0 4,0 3,0 2,0 1,0 0,0 Cd pH 1,5 (HNO3)

Cr pH 2,0 (HNO3)

Cu pH 1,5 (HCl)

pH 2,0 (HCl)

Fig. 39.5. Extractable Cd, Cr and Cu amounts in the second step of the original BCR-SEP.

1244

Sequential extraction

Belzile [30] found no signiﬁcant degree of readsorption of different metals in oxic lake sediments during the extractions. However, substantial redistribution of Pb was observed in different procedures in synthetic soil model systems and spiked natural soils [31]. It is recommended that the effect of readsorption be assessed for each particular sediment to avoid serious misinterpretation of metal mobility [32]. Bermond [33] pointed out that the problem of readsorption is closely linked to the non-selectivity of reagents used in SEPs, an important factor being the initial pH. Readsorption phenomena are increased if compared with low pH values, when the ﬁnal acidity conditions are such as to allow selectivity of the reagents (pH 4 –6). To avoid or to correct for readsorption artefacts arising from the application of SEPs to the subsequent phase, different strategies have been proposed. One approach is the assessment of the binding capacity of the remaining solid residue instead of in the solubilised fractions [34]. Another is to add nitrilotriacetic acid (NTA) to reduce or completely eliminate sorption [35] or to use a radiotracer [36]. 39.2.5.4 Suitability of the methodology Most of the SEPs have been designed for the extraction of cationic elements and the feasibility of their application to elements that can exist as anionic species has to be questioned [37]. Thus, specially designed SEPs based on the nature of the association with solid-phase components of sediments have been proposed for As [38] and phosphorous forms [39]. Gruebel [40] pointed out that As and Se associated with amorphous Fe-oxides are usually not found in solution after reductive dissolution due to readsorption on other mineral phases. Recent work for assessing As and Hg mobility by the application of the modiﬁed BCR-SEP, designed for heavy metals, proposed the strategy of reversing the BCR procedure by measuring the As and Hg contents in the highconcentration residues rather than in the low-concentration extracts, using solid-sampling Zeeman effect ET-AAS [41,42]. Extremely low extractable amounts were found, especially for Hg in step 2 of the procedure, a fact that may indicate the necessity of using specially designed procedures for these types of elements. 39.2.5.5 Conclusions The limitations mentioned in SEPs require that: † † †

The fractions obtained are not considered as geochemical phases. A standardised procedure has to be applied and validated to obtain comparable results. There is a strict adherence to the procedure, including the extraction times and waiting times between steps, to minimise readsorption or redistribution processes.

1245

A. Sahuquillo and G. Rauret

†

Application of a speciﬁcally designed SEP is conducted for each element or group of elements.

In spite of all these limitations, SEPs are considered to be very valuable decision-making tools that present advantages, such as sufﬁcient sensitivity of reagents to quantify mg kg21 concentrations in the fractions, low specialisation requirements for instrumentation and favourable time investment/information ratios. The correct interpretation of the results and an increased awareness of the pitfalls noted above are the responsibility of the scientist in the ﬁeld.

39.3

SAMPLE PRE-TREATMENT FOR SEPs

As a part of the analytical process, sample pre-treatment is a step that must be carefully taken into consideration because it might inﬂuence the ﬁnal analytical data obtained. Many of the aspects considered in this section are common not only to fractionation procedures but also to other chemical analyses. Once the sampling strategy has been designed, including sites, number of replicates, sampling devices and procedure, the proper sample handling that takes place between sampling and analysis is of crucial importance to the quality of data (contamination, mixing, etc.) and to the interpretation of such data (fraction analysed, modiﬁcation of the original matrix) [43]. It is widely known that, prior to being submitted to a SEP, storage and preparation of samples may affect the extractability of elements [44]. Consequently, the information provided may differ from the environmental point of view. However, different steps are currently applied in the sample pre-treatment process, such as drying, grinding, sieving and homogenisation. Table 39.2 summarises the sample pre-treatment conditions described in the literature for natural samples (CRMs are not included). Under the section on wastes, sewage sludges, municipal solid wastes and ﬂy ashes are considered. Further comments on the different steps are included in the subsequent sections. 39.3.1 Drying Usually, the ﬁrst step in the sample pre-treatment described in the literature is a drying step. Many of the procedures applied use an air-dried sample or a sample dried at relatively low temperature (40 –608C). According to the literature, higher temperatures are used for wastes. Problems arising from the use of dried samples are especially important in regard to anoxic sediments, where major changes in relative proportions of the different fractions of heavy metals were observed [85]. Initially, anoxic canal sediments presented redistribution from residual phases into more mobile

1246

TABLE 39.2 Sample pre-treatment processes described in the literature Wet sieving

Drying conditions [References]

Fractions considered [References]

Observations [References]

Sediments

Yes

Air-drying [19] 528C [45] Evaporation at 808C [46] Freeze-drying [47] Air-drying [22,32,48–53] 40–608C [54– 57] 80–1108C [7,49,58– 61] Freeze-drying [38,54]

,850 mm [3] ,250 –200 mm [45,47] ,80 mm [46] ,63 mm [20,62,63] ,2 mm– 850 mm [32,50,52,58] ,250 –200 mm [55,56] ,125 –80 mm [48,51,57] ,63– 50 mm [22,52,53,59,60] Not speciﬁed [49]

Sampling under N2 [63] Sampling under Ar [45] Sampling and frozen [19]

No

Sampling under N2 [38] Sampling and frozen [54]

Soils

No

Air-drying [64– 73] 30–408C [57,74,75] 808C [49] Lyophilisation [76]

,2–1 mm [65,67– 69,71 –73,75,77] ,200 –170 mm [64,66,76] ,125 mm [57] ,63– 50 mm [70,74] Not speciﬁed [49]

Sampling under N2 [76]

Wastes

No

Air-drying [17,78]

,500 mm [17]

Working on wet samples without sieving [18]

40–608C [79,80] 100– 1108C [15,81 –84]

,170 –100 mm [80,83,84] ,70– 63 mm [78,81,82]

Sequential extraction

Type of sample

1247

A. Sahuquillo and G. Rauret

phases during air-drying and oxidising from 2 to 12 weeks, except for Cd and As [86]. This effect is not so important in superﬁcial samples collected in oxic conditions and drying is recommended to stop the microbiological activity and to avoid changes in speciation due to reduction reactions and pH evolution [87]. Instead of drying, freezing samples or keeping them at sufﬁciently low temperatures provides an alternative to limiting biological and chemical activities in marine sediments [54]. The effect of drying samples at 608C on easily exchangeable fractions was studied by Ra¨isanen in organic stream sediments [88]. Drying decreases the extractability of Fe and metals easily coprecipitated with Fe or organic complexes. A study of the effect of higher temperatures on dredged sediments showed that at 250 and 4508C the acid extractable fractions increased for many heavy metals [89]. Another risk involved in the drying step is the possible loss of volatile elements. Freeze-drying is the alternative method for minimising this risk and to obtain powdery materials instead of hard aggregates [54]. This increases the ﬁnal homogeneity of the sample, but it is a methodology that is used in very few cases. Even if it may alter chemical forms of metals, drying is a normal practice in laboratories due to several advantages: ease of handling of dried samples [22], difﬁculty in correcting for an indeterminate amount of water retained by a wet sample [19] and higher reproducibility of results obtained with dried samples [38]. We must bear in mind that all drying procedures disturb the original metalfractionation distribution [90] and, strictly speaking, environmentally relevant information can only be obtained when dealing with wet samples. The effect of different drying methods is particularly signiﬁcant in water-soluble, exchangeable and carbonate-bound fractions [91]. A compromise has to be taken between information usefulness and minimum reproducibility required for trusting the results obtained. 39.3.2 Grinding and sieving steps After drying, in many cases a grinding step is carried out as a way of increasing the homogeneity of natural samples that might be physically less homogeneous than a reference material [23]. SEPs usually require 0.2 –5 g of sample intake and are mainly applied to 0.5 –1 g of subsample. Even if little change in metal extractability and precision results from sample intake of soil (, 1 mm) in the range 1–5 g [92], problems may arise with more complex samples, such as solid wastes from different origins, where heterogeneity is inherent to this type of sample. The literature describes different procedures for grinding. The softest procedure is a disaggregation or gentle crushing step using a wooden-roller,

1248

Sequential extraction

and this is widely used in the sample pre-treatment of soils. In other cases, more aggressive techniques are described, such as milling in an agate mortar, in a planetary mortar or in a mechanical rotary mill. Even if grinding increases the homogeneity of samples, it also affects the speciﬁc surface of the material and the characteristics of these particles, such as organic matter–mineral phase interactions. Consequently, the extractable trace metal content may change. The increase or decrease in extractability depends on the nature and chemistry of the considered element and on the extraction agent used. A study performed on extractability of Cd, Cr, Ni, Pb and Zn in a given soil showed that a ground sample (, 63 mm) presented higher extractable amounts of Cd, Cr, Ni and Zn using a dilute acetic acid solution with respect to the original soil sample (, 2 mm), Ni and Cr being detected at up to 50 and 80%, respectively. Only when using an extraction agent capable of higher interaction with metals, such as EDTA, the grinding showed no effect on extractability (except for Cr) [93]. Grinding and sieving procedures can be performed on samples in different sequences. As it can be seen in Table 39.2, wet sieving is described in a few cases as the ﬁrst step in the sample pre-treatment of sediments, even if further grinding and sieving of samples also occur once the sediment is dried. For soils, a previous sieving at 2 mm is performed after drying and disaggregating the sample; then further milling and sieving is undertaken for chemical analysis. However, parameters for characterisation of soils, such as texture, pH, organic matter and cation exchange capacity, are run on the , 2 mm fraction [64,66]. There is no consensus in the literature on the ﬁnal fraction considered to apply SEPs. Thus, data arising from many different fractions can be found (see Table 39.2). Coarse-grained fractions (2000 –63 mm) with metal poor minerals usually act as a diluent for trace metal concentrations. Clay and silt particles (,63 mm) generally contain the highest concentrations of pollutants and are the most commonly transported in suspension, being an important fraction for environmental concern [54,62,94]. The effect of the considered fractions on extractability makes regional comparisons of pollutant concentrations possible only by using texturally equivalent samples and/or size fractions. Harmonisation of methodology in this sense is therefore just as necessary as is the use of properly standardised SEPs. 39.3.3 Use of inert atmosphere The initial steps of sample pre-treatment, including sampling, is performed in some cases under an inert atmosphere (N2 or Ar) in order to avoid oxidation of the material [38,78,95]. The manipulation of sample under an inert atmosphere is particularly important for anoxic sediments. Ngiam [45] showed that the nature of existing SEPs made it difﬁcult to maintain the natural redox state of anoxic sediments

1249

A. Sahuquillo and G. Rauret

during extraction. Despite the maintenance of an oxygen-free environment, three different SEPs oxidised the sediments for the reducible fraction. This artefact resulted in an over-estimation of the reducible fraction and an underestimation of the organic/sulﬁde fraction for Cd, Cu, Zn, Pb, Fe and Mn. 39.3.4 Recommendations As stated in the discussion of the previous sections, there is no sample pretreatment step that does not alter the sample in some way or another. Strictly speaking, valuable environmental information can only be obtained when dealing with wet samples with minimum manipulation. However, for samples from oxic environments, a drying step at low temperatures and a soft disaggregation process is recommended to increase homogeneity of samples and to obtain good reproducibility in the measurement step characterising the extracted fractions. Hard grinding of the samples is not advisable in order to maintain their speciﬁc surface as unaltered as possible. The application of SEPs is less recommendable for samples from anoxic environments where the nature of extraction agents can oxidise the sample, even under an inert atmosphere. Taking this fact into account, we conclude that SEPs can provide information of environmental concern, in any case. 39.4

APPLICATION OF OTHER EXTRACTION TECHNIQUES TO SEPs

The application of SEPs to routine analysis is handicapped by the timeconsuming laboratory work required with these types of procedures. With the aim of speeding up the extraction process and of improving accuracy in some cases, different approaches have been described as alternatives to conventional extraction systems. 39.4.1 Microwave Microwave heating has proved to be a very effective technique in reducing acidic digestion times for total heavy metal determination. Microwave power and time of heating are the two variables to be optimised for each matrix. In the case of its application to SEPs, the mass/volume ratio and the reagents used in the conventional procedures are maintained, and power and time are optimised in order to obtain the same yield of extraction as that obtained by conventional procedures. Having as target values the recoveries provided for Cd, Cr, Cu, Ni and Zn by the original BCR-SEP, an optimised microwave oven extraction yielded similar results only in 53% of the cases for an estuarine sediment [96]. A modiﬁed SEP derived from Tessier’s scheme has been developed for calcareous soil samples using microwave heating with a total time of 2 h. The validity of the method

1250

Sequential extraction

was veriﬁed by a mass balance in relation to the total metal contents for two CRM soils [74]. 39.4.2 Ultrasound Ultrasonic energy is also described for metal fractionation in sediments [51,96] and sewage sludges [84]. As expected, if considering the completely different extraction process, signiﬁcant differences with respect to conventional procedures are observed. However, because total extractable amounts of metals were practically the same, the proposed schemes are considered very useful for the information provided and the signiﬁcant decrease in sample preparation time (20 min to 1 day). For application in situations where rapid assessment of metal mobility is required, small-scale ultrasound-assisted single extractions performed on 25 mg test portions of sediments are proposed as an alternative to sequential extractions. The results obtained agreed with those of the original-BCR procedure for Cd, Cr, Cu and Pb. No problems of sample homogeneity are described at the low sample intake required by this method [60]. 39.4.3 Other alternatives A different approach for reducing time has come from the design of a SEP for heavy metals based on CO2 and water extractants using supercritical ﬂuid extraction instrumentation. This system provided similar chemical information to that obtained from the original BCR procedure for sediments. However, further investigation is needed to validate the proposed method [61]. Using a slightly modiﬁed original BCR-SEP, Shiowatana [97] tested a novel continuous ﬂow SEP in soils with a specially designed extraction chamber. With a sample intake of 0.25 g, the system is an exhaustive extraction approach because fresh extractant is continuously passing through the sample until the metal is completely leached, as shown in the so-called extractograms. The three-step SEP can be run in this way within 2–6 h. 39.4.4 Conclusions One single conclusion that can be drawn from all these alternatives proposed to conventional extraction procedures is that even if in general shortened times are generated for microwave, ultrasound and supercritical ﬂuid procedures, a previous optimisation of experimental conditions is still required. The optimisation to obtain the target values in agreement with conventional extractions is matrix- and element-dependent, and thus, their routine applicability remains somewhat uncertain.

1251

A. Sahuquillo and G. Rauret

39.5

QUALITY CONTROL FOR SEPs

The need for a good quality control of analyses has led the Standard, Measurements and Testing Programme (SM&T) of the European Commission to organise a series of interlaboratory projects, including the production of CRMs. The aim is the improvement and quality assurance of determinations of extractable trace elements in soils and sediments analysed over the last 15 years. One of the main problems encountered for the validation of SEP methodology is the lack of suitable CRMs. The great variety of matrices and possible SEPs to be applied make the difﬁculty obvious. Hence, a normal practice is to use as an internal check the mass balance obtained from the sum of extracted fractions and the residual phase with respect to the total trace metal content. The only CRMs available for Cd, Cr, Cu, Ni, Pb and Zn extractable amounts are those of the BCR that follow the original and modiﬁed BCR-SEPs, as a result of wide collaborative efforts of different expert laboratories in the framework of the Measurements and Testing Programme (M&T). Table 39.3 compiles the materials that are available from the Institute for Reference Materials and Measurements (IRMM) in Geel (Belgium). The quality control of sampling and sample pre-treatment is also of paramount importance for the ﬁnal validity of the data produced. Thus, the validation of drying, grinding and other sample pre-treatment steps also has to be considered [102]. The validation of oven-drying processes should be checked for each substance and each type of matrix and freeze-drying could be evaluated by placing a pure trapping substance to control losses, for example. Finally, as all possible sample pre-treatments change the original state of the samples, the availability of wet CRMs, especially for sediments, would be a very welcome way to help the validation of fresh sample analysis.

TABLE 39.3 CRMs available following SEPs Sample

Origin

Extractable amounts

BCR CRM 483 [98] Sewage sludge amended soil BCR CRM 601 [99,100] Lake sediment BCR CRM 701 [101] Lake sediment

Northampton (Great Britain) Lake Maggiore (Varese, Italy) Lake Orta (Piemonte, Italy)

Modiﬁed BCR-SEP (indicative values)

1252

Original BCR-SEP (certiﬁed values) Modiﬁed BCR-SEP (indicative values) Modiﬁed BCR-SEP (certiﬁed values)

Sequential extraction

REFERENCES 1

2 3 4 5

6 7 8 9 10

11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30

M. Kersten and U. Fo¨rstner, Speciation of trace elements in sediments. In: G.E. Batley (Ed.), Trace Element Speciation: Analytical Methods and Problems. CRC Press, Boca Raton, FL, 1989. W.F. Pickering, Ore Geol. Rev., 1 (1986) 83. A. Tessier, P.G.C. Campbell and M. Bisson, J. Geochem. Explor., 16 (1982) 77. D.G. Templeton, F. Ariese, R. Cornelis, L.-G. Danielsson, H. Muntau, H.P. van Leewen and R. Lobinski, Pure Appl. Chem., 72 (2000) 1453. A. Sahuquillo, J.F. Lo´pez-Sa´nchez, G. Rauret, A.M. Ure, H. Muntau and Ph. Quevauviller, Sequential extraction procedures for sediment analysis. In: Ph. Quevauviller (Ed.), Methodologies for Soil and Sediment Fractionation Studies. Royal Society of Chemistry, Cambridge, 2002. C. Gleyzes, S. Tellier and M. Astruc, Trends Anal. Chem., 21 (2002) 451. A. Tessier, P.G.C. Campbell and M. Bisson, Anal. Chem., 51 (1979) 844. A. Tessier, F. Rapin and R. Carignan, Geochim. Cosmochim. Acta, 49 (1985) 183. U. Fo¨rstner, Int. J. Environ. Anal. Chem., 51 (1993) 5. M. Meguellati, D. Robbe, P. Marchandise and M. Astruc, Proc. Int. Conf. Heavy Metals in the Environment, Heidelberg, CEP Consultants, Edinburgh, Vol. 2, 1983, p. 1090. A. Sahuquillo, J.F. Lo´pez-Sa´nchez, R. Rubio, G. Rauret, R.P. Thomas, C.M. Davidson and A.M. Ure, Anal. Chim. Acta, 382 (1999) 317. G. Rauret, J.F. Lo´pez-Sa´nchez, A. Sahuquillo, R. Rubio, C. Davidson, A. Ure and Ph. Quevauviller, J. Environ. Monit., 1 (1999) 57. G. Perin, L. Craboledda, M. Lucchese, R. Cirillo, L. Dotta, M.L. Zanette and A.A. Orio, Proc. 5th Int. Conf. Heavy Metals in the Environment, Vol. 2, 1985, p. 454. A. Tessier and P.G.C. Campbell, Hydrobiologia, 149 (1987) 43. V. Bruder-Hubscher, F. Lagarde, M.J.F. Leroy, C. Coughanowr and F. Enguehard, Anal. Chim. Acta, 451 (2002) 285. P. Van Herck and C. Vandercasteele, Waste Manage., 21 (2001) 685. J. Chwastowska and K. Skalmowski, Int. J. Environ. Anal. Chem., 68 (1997) 13. J. Scancar, R. Milacic, M. Strazar, O. Burica and P. Bukovec, J. Environ. Monit., 3 (2001) 226. G.M. Accomasso, V. Zelano, P.G. Daniele, D. Gastaldi, M. Ginepro and G. Ostacoli, Spectrochim. Acta A, 49 (1993) 1205. P.M.V. Nirel and F.M.M. Morel, Water Res., 24 (1990) 1055. C. Whalley and A. Grant, Anal. Chim. Acta, 291 (1994) 287. C.M. Davidson, R.P. Thomas, S.E. McVey, R. Perala, D. Littlejohn and A.M. Ure, Anal. Chim. Acta, 291 (1994) 277. B. Marin, M. Valladon, M. Polve and A. Monaco, Anal. Chim. Acta, 342 (1997) 91. I.N. Benı´tez and J.-P. Dubois, Int. J. Environ. Anal. Chem., 74 (1999) 289. A.M. Ure, Ph. Quevauviller, H. Muntau and B. Griepink, Int. J. Environ. Anal. Chem., 51 (1993) 135. Ph. Quevauviller, Trends Anal. Chem., 17 (1998) 289. Ph. Quevauviller, G. Rauret, J.F. Lo´pez-Sa´nchez, R. Rubio, A. Ure and H. Muntau, Sci. Total Environ., 205 (1997) 223. Ph. Quevauviller, Trends Anal. Chem., 17 (1998) 632. M. Pueyo, G. Rauret, D. Lu¨ck, M. Yli-Halla, H. Muntau, Ph. Quevauviller and J.F. Lo´pez-Sa´nchez, J. Environ. Monit., 3 (2001) 243. N. Belzile, P. Lecomte and A. Tessier, Environ. Sci. Technol., 23 (1989) 1015.

1253

A. Sahuquillo and G. Rauret 31 32 33 34 35 36 37

38 39

40 41 42 43

44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62

1254

M. Ratsasataya, A.G. Langdon and N.D. Kim, Anal. Chim. Acta, 332 (1996) 1. J.L. Go´mez-Ariza, I. Gira´ldez, D. Sa´nchez-Rodas and E. Morales, Anal. Chim. Acta, 399 (1999) 295. A. Bermond, Anal. Chim. Acta, 445 (2001) 79. J.R. Lead, J. Hamilton-Taylor and W. Davidson, Sci. Total Environ., 209 (1998) 193. J.L. Howard and W.J. Vandenbrink, Environ. Pollut., 106 (1999) 285. E.A. Gilmore, G.J. Evans and M.D. Ho, Anal. Chim. Acta, 439 (2001) 139. G. Gleyzes, S. Tellier and M. Astruc, Sequential extraction procedures for the characterisation of the fractionation of elements in industrially-contaminated soils, In: Ph. Quevauviller (Ed.), Methodologies for Soil and Sediment Fractionation Studies. Royal Society of Chemistry, Cambridge, 2002. N.E. Keon, C.H. Swartz, D.J. Brabander, C. Harvey and H.F. Hemond, Environ. Sci. Technol., 35 (2001) 2778. V. Ruban, J.F. Lo´pez-Sa´ nchez, P. Pardo, G. Rauret, H. Muntau and Ph. Quevauviller, Sequential extraction procedures phosphorous forms in lake sediments, In: Ph. Quevauviller (Ed.), Methodologies for Soil and Sediment Fractionation Studies. Royal Society of Chemistry, Cambridge, 2002. K.A. Gruebel, J.A. Davis and J.O. Leckie, Soil Sci. Soc. Am. J., 52 (1988) 390. A. Sahuquillo, G. Rauret, A. Rehnert and H. Muntau, Anal. Chim. Acta, 476 (2003) 15. A. Sahuquillo, G. Rauret, M. Bianchi, A. Rehnert and H. Muntau, Anal. Bioanal. Chem., 375 (2003) 578.. M. Perttila¨ and B. Pedersen, Quality assurance of sediment sampling. In: Ph. Quevauviller (Ed.), Quality Assurance in Environmental Monitoring. Sampling and Sample Pretreatment. VCH, Weinheim, 1995. A.K. Das, R. Chakraborty, M.L. Cervera and M. de la Guardia, Talanta, 42 (1995) 1007. L.-S. Ngiam and P.-E. Lim, Sci. Total Environ., 275 (2001) 53. P. Szefer, G.P. Glasby, J. Pempkowiak and R. Kaliszan, Chem. Geol., 120 (1995) 111. P. Svete, R. Milacic and B. Pihlar, J. Environ. Monit., 3 (2001) 586. B.G. Prusty, K.C. Sahu and G. Godgul, Chem. Geol., 112 (1994) 275. I.T. Urasa and S.F. Macha, Int. J. Environ. Anal. Chem., 64 (1996) 83. Z. Mester, C. Cremisini, E. Ghiara and R. Morabito, Anal. Chim. Acta, 359 (1998) 133. B. Pe´rez-Cid, I. Lavilla and C. Bendicho, Int. J. Environ. Anal. Chem., 73 (1999) 79. A.V. Filgueiras, I. Lavilla and C. Bendicho, Anal. Bioanal. Chem., 374 (2002) 103. A.V. Filgueiras, I. Lavilla and C. Bendicho, Anal. Chim. Acta, 466 (2002) 303. D.H. Loring and R.T.T. Rantala, Earth Sci. Rev., 32 (1992) 235. C. Banﬁ, R. Cenci, M. Bianchi and H. Muntau, EUR-Report 1992, Istituto dell’Ambiente, Centro comune di ricerca. M.B. Alva´rez, M.E. Malla and D.A. Batistoni, Fresenius J. Anal. Chem., 369 (2001) 81. R.A. Sutherland, Appl. Geochem., 17 (2002) 353. S.J. Hoffman and W.K. Fletcher, J. Geochem. Explor., 15 (1981) 549. R. Martı´n, D.M. Sa´nchez and A.M. Gutie´rrez, Talanta, 46 (1998) 1115. S. Tokalioglu, S. Kartal and L. Elc¸i, Anal. Chim. Acta, 413 (2000) 33. Gy. Heltai, K. Percsich, I. Fekete, B. Baraba´s and T. Jo´zsa, Microchem. J., 67 (2000) 43. S. Stamoulis, R.J. Gibbs and M.G. Menon, Environ. Int., 22 (1996) 185.

Sequential extraction 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87

88 89 90 91 92 93 94 95 96

Z. Borovec, Sci. Total Environ., 177 (1996) 237. X. Li, B.J. Coles, M.H. Ramsey and I. Thornton, Chem. Geol., 124 (1995) 109. D. Mc Grath, Sci. Total Environ., 178 (1996) 37. M.C. Jung and I. Thornton, Sci. Total Environ., 198 (1997) 105. C.M. Davidson, A.L. Duncan, D. Littlejohn, A.M. Ure and L.M. Garden, Anal. Chim. Acta, 363 (1998) 45. J. Tho¨ming and W. Calmano, Acta Hydrochim. Hydrobiol., 26 (1998) 338. I. Maiz, I. Arambarri, R. Garcı´a and E. Milla´n, Environ. Pollut., 110 (2000) 3. M.D. Ho and G.J. Evans, Environ. Sci. Technol., 34 (2000) 1030. W.W. Wenzel, N. Kirchbaumer, T. Prohaska, G. Stingeder, E. Lombi and D.C. Adriano, Anal. Chim. Acta, 436 (2001) 309. V.F. Melo, B. Singh, C.E.G.R. Schaefer, R.F. Novais and M.P.F. Fontes, Soil Sci. Soc. Am. J., 65 (2001) 1324. V.F. Melo, C.E.G.R. Schaefer, R.F. Novais, B. Singh and M.P.F. Fontes, Commun. Soil Sci. Plant Anal., 33 (2002) 2203. E. Campos, E. Barahona, M. Lachica and M.D. Mingorance, Anal. Chim. Acta, 369 (1998) 235. O. Schramel, B. Michalke and A. Kettrup, Sci. Total Environ., 263 (2000) 11. J. Arunachalam, H. Emons, B. Krasnodebska and C. Mohl, Sci. Total Environ., 181 (1996) 147. N. Pustisek, R. Milacic and M. Veber, J. Soils Sediments, 1 (2001) 25. E. Alonso Alva´rez, M. Callejo´n Mocho´n, J.C. Jime´nez Sa´nchez and M. Ternero Rodrı´guez, Chemosphere, 47 (2002) 765. S. Eichfeld, J.W. Einax and G. Knapp, Anal. Bioanal. Chem., 372 (2002) 801. R. Zuﬁaurre, A. Olivar, P. Chamorro, C. Nerı´n and A. Callizo, Analyst, 123 (1998) 255. B. Pe´rez-Cid, I. Lavilla and C. Bendicho, Analyst, 121 (1996) 1479. I. Lavilla, B. Pe´rez-Cid and C. Bendicho, Anal. Chim. Acta, 381 (1999) 297. B. Pe´rez-Cid, I. Lavilla and C. Bendicho, Fresenius J. Anal. Chem., 363 (1999) 667. J. Scancar, R. Milacic, M. Strazar and O. Burica, Sci. Total Environ., 250 (2000) 9. M. Kersten and U. Fo¨rstner, Water Sci. Technol., 18 (1986) 121. S.R. Stephens, B.j. Alloway, A. Parker, J.E. Carter and M.E. Hodson, Environ. Pollut., 114 (2001) 407. A.M. Ure, C.M. Davidson and R.P. Thomas, Single and sequential extraction schemes for trace metal speciation in soil and sediment, In: Ph. Quevauviller, E.A. Maier and B. Griepink (Eds.), Quality Assurance for Environmental Analysis. Elsevier, Amsterdam, 1995. M.L. Ra¨isa¨nen, L. Ha¨ma¨la¨inen and L.M. Westerberg, Analyst, 117 (1992) 623. F.M. Tack and M.G. Verloo, Sci. Total Environ., 178 (1996) 29. C.M. Davidson, L.E. Wilson and A.M. Ure, Fresenius J. Anal. Chem., 363 (1999) 134. S. Zhang, S. Wang and X.-Q. Shan, Chem. Spec. Bioavail., 13 (2001) 69. C.M. Davidson, P.C.S. Ferreira and A.M. Ure, Fresenius J. Anal. Chem., 363 (1999) 446. A. Sahuquillo, H. Bosch, G. Rauret and H. Muntau, Fresenius J. Anal. Chem., 360 (1998) 304. U. Fo¨rstner and W. Salomons, Environ. Technol. Lett., 1 (1980) 494. J.F. Lo´pez-Sa´nchez, R. Rubio, C. Samitier and G. Rauret, Water Res., 30 (1996) 153. I. Ipolyi, C. Brunori, C. Cremisini, P. Fodor, L. Macaluso and R. Morabito, J. Environ. Monit., 4 (2002) 541.

1255

A. Sahuquillo and G. Rauret 97 98

99 100

101 102

1256

J. Shiowatana, N. Tantidanai, S. Nookabkaew and D. Nacapricha, J. Environ. Qual., 30 (2001) 1195. G. Rauret, J.F. Lo´pez-Sa´nchez, A. Sahuquillo, E. Barahona, M. Lachica, A. Ure, H. Muntau and Ph. Quevauviller, (Addendum to EUR-Report 17127 EN), EUR Report 19503 EN, European Commission, 2000. Ph. Quevauviller, G. Rauret, J.F. Lo´pez-Sa´nchez, R. Rubio, A. Ure and H. Muntau, EUR Report 17554 EN, European Commission, 1997. G. Rauret, J.F. Lo´pez-Sa´nchez, A. Sahuquillo, H. Muntau and Ph. Quevauviller, (Addendum to EUR Report 17554 EN), EUR Report 19502 EN, European Commission, 2000. G. Rauret, J.F. Lo´pez-Sa´nchez, D. Lu¨ck, M. Yli-Halla, H. Muntau and Ph. Quevauviller, EUR Report 19775 EN, European Commission, 2001. Ph. Quevauviller, E.A. Maier and A. Boenke, Quim. Anal., 13 (1994) S124.

Index AAS see atomic absorption spectroscopy AB see arsenobetaine abiotic matrices 333– 7 ablated mass 602–3 ablation 598, 599– 601 see also laser ablation absolute sensitivity 169, 178 absorptive methods 1218– 20 accelerated solvent extraction (ASE) 343–52 applications 349– 51 basic principals 344 instrumentation 345– 6 methods development 336, 351 accelerating sample dissolution 659 –60 acceptor-controlled conditions 564 –5 accumulation process 538–41 accuracy 48, 309 acetic acid 982 acid digestion 199– 227, 735–50 acid emulsiﬁcation 710– 11 acid mineralization 697– 704 acid vapor steaming 26, 27 actinides 329–30 additive interference 48– 9, 122 adsorbents 514 adsorption losses/contamination 29, 31– 2 metal ions 474– 7 sequential extraction 1244– 5 solid phase extraction 395– 6 volatile metals 1218– 20, 1227–9 adsorptive stripping voltammetry 184– 5 aerobic 1216 aerosols 903– 32 chromium 1146 impactors 909–16 sampling 903–34

sequential extraction 924– 30 transport 904 –6, 919– 21 AES see atomic emission spectrometry AFS see atomic ﬂuorescence spectrometry agriculture 676, 1051, 1053–5, 1058 air 1214– 9, 1222– 3, 1227 see aerosols airborne impurities 28– 9, 31– 4, 969– 71 arsenic 1031– 3 contamination/losses 42 cryogenic trapping 504–9 selenium 1055–6 algae 535– 6, 546– 50, 1039–40 alginate 541– 2 aliquot 932 alkaline digestions 949– 50 alkyl-volatile metal(loid)s 497 alkylethoxy-mercury 1075 alumina 288 –9, 420– 1 aluminum 888 ambient air 510–1, 920 –1, 1223–4 ambient atmospheric particles 912– 3 ambient particulate matter 921– 2 ambient sampling 916–8 American Society for Testing and Materials (ASTM) 857 ammonium nitrate 982 ammonium pyrrolidine dithiocarbamate (APDC) 477, 486 anaerobic 1214, 1216 anaerobic reducing environments 1214– 5 analysis optimization 296–7 analyte moiety 1188 analyte recovery 1188– 93 animal tissue 237, 252–3, 1203 anion-exchange 1143–5

Comprehensive Analytical Chemistry XXXIX, pages 1257–1286 q 2003 Elsevier Science B.V. All rights reserved. ISSN: 0166-526X

1257

anodic stripping voltammetry (ASV) 183–4, 360 anthropogenic 1212, 1214, 1216 anthropogenic volatile species 497, 498–502 anti-cancer drugs 1173– 80 anti-knock agents 1081– 111 anti-reﬂective coatings (ARC) 983–4 antifouling paints 991 –1020 antimony 551, 877, 878, 890, 892 AOAC International 858 APDC see ammonium pyrrolidine dithiocarbamate apparatus see instrumentation aquatic samples 219, 991 –1020 aqueous systems chromium 1118– 19, 1121 derivatization 580– 3, 584 –5, 589 extraction 706 industrial waste 939– 42 organotins 333–4 soil-sediment extracts 1058–9 solution chemistry 1063– 5 supercritical ﬂuid extraction 329 ARC see anti-reﬂective coatings aromatic hydrocarbons 914– 5 arsenate 1034–5 arsenic biotrapping 551 compounds 339, 349– 50, 1029, 1033– 40 derivatization 577–92, 578 –581, 583–6 dry ashing 248– 53 occurrence/distribution 1028– 33 organoarsenic 1200– 1 solid phase microextraction 381–5 speciation 349–50, 1027– 41 urine 1196– 7 water 877, 878, 888, 892, 893 arsenic trioxide 1027 arsenite 1034– 5 arsenobetaine (AB) 1028–9, 1037 arsenocholine 1037 arsenosugars 1030, 1037– 8 arsphenamine 1027 arthritis 1180 –1 Artic 919 –21 1258

artifactual formation 1075–6 ASE see accelerated solvent extraction ash dissolution 695 ash solubilization 700 –1 ashing 250 –1, 693– 7 see also dry ashing ASTM see American Society for Testing and Materials Astragalus praleongus 1052 ASV see anodic stripping voltammetry atmosphere atmospheric budget 929 –30 gaseous constituents 495, 496 selenium 1055–6 sequential extraction 1248–9 atmospheric… fractionation 924–30 open vessel microwave extractions 283– 5 pressure microwave digestion 700–1 atom trapping 130 atomic absorption spectroscopy (AAS) see also electrothermal… chelation solvent extraction 488, 490– 1 chromium 1124–30, 1138–43 cold vapor 487, 938 ﬂame 118– 35, 235, 630–1, 938, 1142 ﬂow injection digestion 644– 5 geological samples 729 graphite furnace 486, 488, 938, 1138 hydride generation 249, 938 industrial waste 938 lead 1104 quantitation 164– 5 solid phase extraction 404 –9 vapor generation 638–41, 644–5 atomic emission spectrometry (AES) 118–35, 164–5, 1142 atomic ﬂuorescence spectrometry (AFS) 160– 5, 858 ﬁgures of merit 163 instrumentation 158–63 interferences 162 –6 sample introduction 127–35, 146– 54, 158– 9, 163 theory 162

autoclave digestions 701–2 automation 649 –77 glow discharge 173 high-volume aerosol samplers 918–9 liquid samples 653–9 membrane extraction 569 –70 robotics 660 –76 solid phase extraction 403– 4 solid phase microextraction 379 solid samples 659–60 wet acid digestion 220, 223 axial plasmas 144– 5 background correction 121– 3, 309 bacteria 536– 7, 547–8, 549– 50 balances 974 bar codes 652 –3 batch approaches 651–2, 656 –9 BCR procedure geological samples 757 organotin compounds 1006, 1008–10, 1017 sequential extraction 1238, 1239, 1251 Beer– Lambert Law 179 Berner impactors 909, 912 –3 BET isotherms 508 beverages 292– 3 bioabsorption 533–4 bioaccessibility of metals in foodstuffs 1192–3 bioavailability 1063 biogeochemical 496, 577, 659, 865, 905, 1092, 1234 biogeochemical cycles 1092 bioindicators 557 biological organotin compounds 582–3 biological samples chelate extraction 483–4, 490 –1 chelating resins 485, 490 –1 chromium 1153– 5 contamination/losses 36–7 ﬂuids chromium 1136– 45 metal complexes 1198

metal-based drugs 1199–200 metallodrug metabolites 1199– 200 selenium 1046– 9 ultrasonic extraction 363– 4, 365 food analysis 771– 839 laser ablation 605 mercury 1069 metallobiomolecules 1185 –206 organotin compound samples 995 –6 platinum-based drugs 1175 tissue 363– 4, 365, 1073– 4 biological substrates 535 –8, 546– 8, 549– 50 biomass 533– 57 biomethylation 1032– 4, 1119 biomonitors 557 bioremediation 556 biotechnology 676 biotic matrices 337– 9 biotransformation 540–1 biotrapping 533–57 analytical applications 546– 56 immobilization 541– 6 methodology 541– 6 speciation 6, 503, 728, 905 technical applications 556–7 uptake mechanisms 538– 41 birds 1050–1 bis(acetato)amminedichloro(cyclohezylamine)platinum(IV) 1174, 1175, 1178, 1179, 1180 bismuth 878, 888, 893 blood arsenic 1033 chelation solvent extraction 491 chromium 1136, 1139, 1142 –4 contamination/losses 34– 5 lead levels 1087–8 selenium 1046–9 boat hulls 991– 1020 boiling points metal carbon bonds 1213–4 metal-hydrogen bonds 1213– 4 volatile metals 1213–4 volatile species 502– 3 bombs 206, 207–8, 701 –2, 704– 5 1259

borates 301– 10, 580– 3, 589, 750–2 bottles 971– 2, 979–80 Brassica juncea 1052 Braunu¨len 35 Brazil nuts 1054 breast milk 1046–9 brines 866 broccoli 1053 buffer 1188– 9, 1196, 1198, 1201, 1203–5 bulk samples 176 butyl rubber 696– 7, 715– 8 butyltin compounds 991– 1020 butylation 1108 CU18u-bonded silica gel 415– 20 cadmium 534, 886, 888 –9, 892– 3, 1244 calcium 716, 1243 calibration 47– 90 approach selection 49– 52 curve technique 51 external calibration 51, 67– 74 internal standardization 51, 79–81, 82 isotope dilution mass spectrometry 51– 2, 81–90 laser ablation 594, 601 –3 linear regression 55– 67 matrix matching 51 method of standard additions 52, 74– 9, 978–9 recovery data statistics 52– 5 semiconductors 978–9 solid phase microextraction 378–9 standard additions 52, 74– 9, 978–9 standard preparation 979 terminology 48–9 traceability 99 cancer drugs 1173– 80 canister sampling 1216, 1217 capacitively coupled plasma 1105 capillary electrophoresis 1193– 5 capsules 209 carbohydrate 838, 842 carbohydrates 292–3, 1187 –8 carbon dioxide 314 –40, 510 –2 carbon sorbents 431– 2 carbonates 926 carbonyl-volatile metal(loid)s 497 1260

carboplatin 1174, 1177, 1179, 1180 carbosieve adsorption 1227 Carius tubes 204 cartridges 400, 401–2 cascade impactors (CIs) 909– 16 cathodic stripping voltammetry (CSV) 184– 5, 885, 888 –91 cations 397– 8 cavities 263, 267, 268 cellulose 432 centrifuge 667 certiﬁed reference materials (CRMs) 102, 957 homogenization 38– 9 industrial waste 957, 958– 9 methylmercury 1077–8 organotin compounds 1003–4, 1008, 1010, 1017 sequential extraction 1239, 1252 stated references 93– 114 traceability 94–114 chelates extraction 318–26, 330–1, 397 –8, 459– 92 hard-soft acid– base nature 463–5 supercritical ﬂuid extraction 318– 26, 330– 1 chelating resins 474– 7 biological samples 485 ion exchange 860, 863– 5 solid phase extraction 430 –1 chelation solvent extraction 459–94 distribution ratio 461 –3 enrichment factor 465–6, 564–5, 568– 9 equilibrium constants 460 extraction constants 459, 463, 467 masking 469 –72 preconcentration 459, 465–6 principals 677 sample preparation 477–92 separation factor 467, 471, 475 speciation 459, 766 stability constants 473– 4, 492 chemical... composition of aerosols 913 data traceability 93–114 fractionation 1241– 2

impurities 965, 968 interference 121–2 measurement traceability 94–5 principles in membrane extraction 566 semiconductors 965, 967 stated references 110– 3 structures 1029, 1030 vapor generation 127– 9, 140– 1, 150, 163 chemiluminescence (CL) chromium 1120, 1133– 4, 1143 quantitation 181 –3 water 880, 886–7 chromate ions 552– 3 chromatography see also gas chromatography; liquid chromatography electrothermal atomic absorption spectrometry 141 ﬂame spectrometry 131 –2 HPLC 131–2, 148, 1143– 5 inductively coupled plasma-atomic optical emission spectroscopy 148–50 metallobiomolecules 1193– 6 supercritical ﬂuids 313, 1103 chromites 745, 1117– 8 chromium biological ﬂuids 1136– 45 biotrapping 534, 551–3 chemical species 1118–20 industrial abundance 1117–8 methodology 1120–57 natural abundance 1117–8 reactivity 1115–6 sample preparation 1115– 58 sequential extraction 1244 solid phase extraction 420– 1, 439–43 solid samples 1145–57 speciation 1115–58 ultrasonic extraction 364, 365 water 886, 889, 893, 1120– 36 CII see Comite´ Inter-Instituts d’Etude des Techniques Analytiques cis-amminedichloro(2-methylpyridine)platinum(II) 1174, 1179, 1180

cis-diammine(1, 1-cyclobutanedicarboxylato)platinum(II) 1174, 1177, 1179, 1180 cis-diamminedichloroplatinum(II) 1174– 7, 1179, 1180 cisplatin 1174– 7, 1179, 1180 citrus leaves 280– 1 CL see chemiluminescence classical detection techniques 117– 8 classiﬁcation ﬂow injection 612– 13 food analysis 770, 771–2 solvent-free samples 371–2 clean area working 28– 9, 31– 4 clean chemistry 297 clean rooms 969 –71 clean-up contamination 26– 9 membrane extraction 567– 8 metallobiomolecules 1195 –6 cleaning samples 1066 cleanliness 713– 5 clinical laboratories 674 clinical samples 1153 –5 closed systems high pressure acid digestion 740–3 microwave extraction 273 –4, 282– 3 wet acid digestion 199, 204– 10, 214– 5, 221, 225 –6 co-precipitation 754–5 coatings 132, 375, 376, 983 –4 coaxial segmentors 618–9 cobalt 886, 888– 90, 893 coefﬁcients 53– 7, 314–6 coils 619 cold vapor atomic absorption spectroscopy (CV-AAS) 487, 938 collection strategies general principles 14–7 geochemical research 724– 6 mercury 1065–7 organotin compounds 995 –8 volatile metals 1218 collision cell technology 859 colloidal silica 985 columns 619, 622, 627–30 1261

combustion contamination/losses 40–1 ﬂue gas 521, 522 oxygen combustion 704 –5, 788–94, 835 Comite´ Inter-Instituts d’Etude des Techniques Analytiques (CII) 247– 8 commercial microwave extraction devices 257–8 complete dissolutions 287–8 complete trapping 562 –3 completely continuous ﬂow analysis 654, 656 complex organic extractions 295– 6 complex permittivity 261– 2 complex sequential extractions 290– 1 complex-supercritical ﬂuids 328 complexation 1129– 30 complexing agents 336 composite sampling/analysis 5, 937 composition inﬂuence 246 composition properties 685 –6 concentration external calibration 67–74 free metal ions 472–3 ligands 473–4, 492 microwave extraction 297– 8 soil arsenic 1031 condensation drying 514–5 conditioning parameters 433 conditioning solvents 433 containers 16–7, 34–7, 971– 2 contamination airborne 28– 9, 917 analyte losses 23–42 crude oils 691 levels 1121 materials 24–7 metallobiomolecules 1205– 6 petrochemistry 714 –5 reagent 24, 27–8, 200 semiconductors 965, 966, 969 –75 continuous automation 653– 6, 672– 4 continuous ﬂow analysis 654, 656 control automated sample preparation 668 materials 1205– 6 microwave extraction 275– 81 1262

controlled pore glass (CPG) 545– 6 conventional electrothermal atomic absorption spectrometry 136– 9 conventional heating microwave heating 260–1 wet acid digestion 199, 200– 1, 204–7, 210– 1, 221, 222 conventional nebulization 123 –7 cooling fusion beads 305 copolymers 423 copper 886, 888–9, 892 –3, 1244 costs, industrial waste 937 coupled separation/detection 1093–6 see also hyphenated techniques coverage factor 54 cracking 307 –8 crater images 594, 597 CRM 858, 880 CRMs see certiﬁed reference materials crucibles 303 crude oils 683– 719 crushing 726–7 cryofocusing 504– 9 cryogenic trapping 495– 529 analytical constraint 509–17 applications 520–8 aqueous derivatization 584– 5 instrumentation 520–8 sample preservation 517, 519 stability 517, 519 volatile metal(loid)s 495–529 volatile metals 1218, 1220– 2, 1223– 7 volatile organic compounds 495– 529 crysotherapy 1180– 1 crystallization 306– 7 CSV see cathodic stripping voltammetry CV-AAS see cold vapor atomic absorption spectroscopy cycles 348– 9, 1092 cysteine 1033 D2EHPA see di(2-ethylhexyl)phosphate DAL see dialkyllead data collection 13– 4 DBT see dibutyltin DC see direct current DCP see direct current plasma

DDDC see diethylammonium diethyldithiocarbamate DDTC extraction 488 decomposition contamination/losses 39–41 geological samples 750– 3 molten salt fusion 750 –3 resistant minerals 745 –50 ultraviolet on-line 211 wet digestion 199 –227 defatted dogﬁsh muscle 1038– 40 defects in semiconductors 965, 968 deﬁning goals 5 –6 deﬁnitions chemical measurement traceability 94– 5 fractionation 1234 sequential extraction 1234, 1243– 4 volatile species 502–3 degradation 999 –1000, 1190 –3 deionized water (DIW) 979 –81 Dekati cascade impactors 909– 12 DEL see diethyllead derivatization 577– 90 applications 585– 9 ethylation 380–6, 580– 1, 1107 Grignard reactions 578 –9, 582– 3, 587 hydride generation 579– 80, 584– 5, 588 lead 1097– 100 methodology 583–5 organoleads 1088–9 organotin compounds 336– 7, 1004– 5 pentylation 1108 phenylation 581– 3 propylation 581, 1107–8 solid phase microextraction 376, 381 –6 theory 578– 83 desiccants 514 desorption 29, 31– 2, 376, 387 detection techniques chromium 1123, 1138– 45 inductively coupled plasma-atomic optical emission spectroscopy 145–6 inductively coupled plasma-mass spectroscopy 155–6

organolead compounds 1104– 7 quantitation 117–86 detector dead time 48, 85–7 deuterium lamp correction 122–3 development guidelines 623, 629– 30, 636– 8 di(2-ethylhexyl)phosphate (D2EHPA) extraction 489 diabetes 1181 dialkyllead (DAL) 1081– 2 1,2-diamminomethylcyclobutaneplatimum(II) 1174, 1175, 1178 dibutyltin (DBT) 996 –7, 1006 dielectric constant 261– 2 dielectric loss factor 261 –2 diethylammonium diethyldithiocarbamate (DDDC) 477, 486 diethyllead (DEL) 1081 differential pulse anodic stripping voltammetry 183–4 diffusion coefﬁcient 314– 16 digestion acid 199–227, 735– 50 alkaline 949– 50 bombs 206, 207 –8, 701– 2, 704–5 chromium 1153 contamination/losses 39– 41 dry ashing 235 –54 ﬂow injection 643– 6 food analysis 794– 809, 812 –29, 836– 8, 843–6 industrial waste 938 –50 organic samples 199, 224, 294, 296 petrochemistry 698–9, 700 –2 semiconductors 976– 8 wet 193– 227 DIGITEL AUTOMATIC high-volume aerosol samplers 918 dilutions 39, 707 –9 dimethyl sulfoxide (DMSO) 983 dimethylarsonic acid 1036– 7 dimethylarsonous acid 1035– 6 dimethyllead (DML) 1081 dimethylmercury extraction 1074– 5 DIN 918, 948 diphenycarbazide 486 dipole rotation 261 1263

direct... current (DC) power 175–6 current plasma (DCP) 165 determinations 386– 7, 733, 857–63 extraction 350, 373– 5, 1074 sample insertion 151 solid sampling 660 discontinuous systems 656 –9 discs 400, 402 –3 dismutation 1075, 1083, 1097 disposable equipment 714– 5 disposable sorbent containers 400–3 dissociation energies 1213–4 dissolution automation 659 –60 contamination/losses 39–41 dry ashing 240– 4 ﬂow injection 643–6 food analysis 778 –95, 834 –6 geological samples 735– 50, 755– 8 microwave extraction 287– 8 petrochemistry 695, 707– 9 sequential extraction 1233– 4 wet digestion 199 –227 distribution ratios 461 –3 dithizone extraction 490– 1 divinylbenzene-based sorbents 422 –3, 424–9 DML see dimethyllead DMSO see dimethyl sulfoxide documented standards 93– 114, 1077– 8 see also certiﬁed reference materials dogﬁsh muscle 1038–40 door protection 273 DRAM 1/2 pitch 966 drinking water 868, 1031–3, 1121 drugs 1173–82, 1199– 200 dry ashing 235– 54 arsenic 248 –53 dissolution 240–4 food analysis 778 –95, 834 –6, 845 methodology 244–8 oxidation process 240 –4 petrochemistry 693 –4 residue dissolution 240 –4 selenium 248–53 1264

total analyte content determination 237 dry oxidations 240–4 drying accelerated solvent extraction 347 contamination/losses 37– 9 efﬁciency 514–5 food analysis 769 gas streams 512– 7, 518 sequential extraction 1246–8 duration, aerosol sampling 907 dust materials 1107 –9 ECD see electron capture detection economic factors 13–4 EDTA masking, lead 1093 EF see enrichment factors efﬁciency ablation 598 drying 514 –5 elution solvents 437 –8 wet acid digestion 216– 9 eggs 1050 EL see ethyl lactate electrical low-pressure impactors (ELPI) 912 electrochemical methods 183 –6 adsorptive voltammetry 184 differential pulse anodic stripping voltammetry 183 cathodic voltammetry 184 ion selective electrodes 48, 185 electrodes 185– 6 electromagnetic spectrum 259 electron capture detection (ECD) 1093 electrons 264–6 electrophoresis 1103– 4, 1193– 5 electrospray mass spectroscopy 1195– 6 electrothermal atomic absorption spectrometry (ET-AAS) chemical vapor generation 127, 140, 150, 158– 160, 163 chromium 1120 dry ashing 235, 249 ﬂow injection solid phase extractions 631– 5 food analysis 769

instrumentation 136, 139 multi-element 139– 40 quantitation 133 –41 speciation 141 vapor generation 487, 577– 92, 635, 636, 879 vapor trapping 639, 683 water 858– 9 electrothermal vaporization (ETV) 150– 2, 769, 859 elevated pressure digestion 701 elution ﬂow injection solid phase extractions 630 parameters 437 –9 solid phase extraction 395– 6, 400 emulsiﬁcation 709–10 emulsions 709– 11 energy dispersive spectrometry 176 energy transfer 258–63 enrichment factors (EF) 465 –6, 564–5, 568 –9 environmental... arsenic 1028–33, 1038– 40 contamination control 965, 966 health 1087– 8 organolead compounds 1083– 5 organotin compounds 582 –3, 991 –1020 samples arsenic compounds 1028– 33, 1038– 40 laser ablation 605 mercury 1069– 70 methylmercury 1063– 4 operational modes 246– 8 selenium 1055–9 ultrasonic extraction 358–60, 361 volatile metal(loid)s 520 –3 Environmental Protection Agency (EPA) 354, 857 –8, 939– 46, 949– 50 enzymatic degradation 1191– 2 enzymatic extraction 1190– 1, 1192–3 EP, industrial waste 956 EPA see Environmental Protection Agency equilibrium constants 460–1 equilibrium extraction 564–5

equipment cleaning 713–4 general principles 16 mercury 1066 microwaves 263– 9 semiconductor contamination 971– 5 wet acid digestion 220, 223 errors 48, 905–6, 931 –2, 1205–6 erythrocytes 1046– 9 Estonian soil samples 944 estuarine waters 866, 868, 872 ET-AAS see electrothermal atomic absorption spectrometry ethyl lactate (EL) 983 ethylation 380– 6, 580–1, 1107 ethylmercury 1075 ETV see electrothermal vaporization Europe, organotin measurements 1005–17, 1020 evaporation 28, 31, 297 –8 exchangeable elements 926 excimer lasers 594, 598–9 experimental set-ups 570– 3, 594– 9 external calibration introduction 51, 67–74 laser ablation 601– 3 semiconductors 978 traceability 99 extractable trace element determinations 108– 10 extraction accelerated solvent extraction 343– 52 agents 334, 1235, 1236–7 arsenic compounds 1038–40 chelates 318– 26, 330– 1, 397–8, 459– 92 chelation solvent 459 –92 chromium 1146–7 constants 461–5 inorganic compounds 343–52 membrane 559–74 metal ions 459 –92 microwaves 257– 99 modes 373 –5 modules 667 liquid–liquid 380, 1071, 1073, 1078 optimization 376 –8 1265

organometallic compounds 343–52 organotin compounds 1000–4 parameters 347 –9 petrochemistry 706 –7 protein denaturing agents 1190 recovery 1100–1 sequential 96, 108, 110, 755–8, 919, 924–31, 1233, 1234, 1251 solid phase 393–451 solid phase microextraction 371–90 solvent 332, 343– 51, 459– 94, 622, 706–7, 766, 1040, 1130 supercritical ﬂuids 313–40 time optimization 377 F –AAS see ﬂame atomic absorption spectrometry falling drop segmentors 618 –9 Fassel torches 144 fats 833, 842 feces 1153– 5 feedstuffs 771– 839 FEP, contamination 25 FI see ﬂow injection ﬁber coating selection 376 ﬁgures of merit atomic ﬂuorescence spectrometry 163 chromium speciation 1131, 1139– 41 ﬂame spectrometry 124 –5 inductively coupled plasma-atomic optical emission spectroscopy 152 inductively coupled plasma-mass spectroscopy 160, 161 ﬁlters 907 –9, 970– 1, 1095 ﬁltration 1189, 1194 –5 ﬁre assay 752 ﬁsh 584–5, 1050–1 ﬁtness-for-purpose 728 ﬂame atomic absorption spectrometry (F-AAS) 118– 35 chromium 1120, 1142 dry ashing 235 ﬂow injection solid phase extractions 630–1 industrial waste 938 ﬂame spectrometry 118– 35 background correction 121– 3 1266

chromatography 131 –2 ﬂow injection 129– 30 instrumentation 119– 21 interferences 121 –3 nebulizers 249, 284, 709 slotted tube atom trap 130 slurries 127, 132– 4, 829– 32, 838– 9 theory 118–9 vapor generation 487, 577 –92, 635–6, 879 ﬂow benches 29, 32, 33 ﬂow injection (FI) apparatus 617 –22 automated sample preparation 654– 5 chemiluminescence 1143 classiﬁcation 612– 3 ﬂame spectrometry 129–30 gas diffusion 641– 3 gas diffusion systems 641 liquid –iquid extractions 615– 26 on-line sample digestion 643– 6 on-line sample pretreatment 612 photo-oxidation 646, 1197 sample pretreatment 611– 46 solid phase extractions 626–35 vapor generation 635–41 ﬂow systems 199, 210– 3, 222, 570–1 ﬂow-rates 434, 439 ﬂue gases 521, 522 ﬂuffy ﬂuxes 302 ﬂuorimetry 408 ﬂuorinated polymers 25 ﬂuoropolymers 713– 4 ﬂuxes 301– 10 foam formation 526 focusing 167 food analysis 765 –847 arsenic 1031– 3 chromium 1155, 1156 classiﬁcation 770, 771– 2 dissolution 778– 95, 834– 6 dry ashing 778 –95, 834 –6, 845 elements 841– 2 examples 843 –6 literature 766 –8

low temperature ashing 788 –94, 836 microwave digestion 818– 29, 838, 845–6 on-line digestion 829–32 organoarsenic 1200– 1 oxidizing acids 794–809, 812– 7, 836–8 oxygen combustion 788– 94, 835 pressure wet digestion 812– 7, 836– 8 pretreatments 768– 832 sample preparation 839 –41 sequential enzymatic extractions 1192–3 slurry preparation 829– 32, 838– 9 wet digestion 794 –809, 812– 29, 836– 8, 843–6 forensics 605 formed metal complexes 473–4 Fourier transform ion cyclotron resonance mass spectrometry (FT–ICR –MS) 1095 fractionation aerosols 924–30 deﬁnition 1234 laser ablation 594, 603 –4 matrix analysis 1240 metallobiomolecules 1193– 5 sequential extraction 1234, 1240, 1241– 2 free metal ions 472– 3, 1187 freeze drying 38 frequency, aerosol sampling 907 fresh water 869, 873, 874, 876 fresh water sediments 1009, 1010 FT-ICR-MS see Fourier transform ion cyclotron resonance mass spectrometry fume cupboards 223 fungi 537– 8, 547–50, 1031, 1054– 5 furnace cleaning 715 fused silica capillaries 316 fusion 301 –10 beads 301, 305– 8, 310 borates 301– 5 contamination/losses 39 decomposition 750– 3 ﬂuxes 301–10 lithium borates 301– 5

gallium 893 gamma spectrometry 732 garlic 1053–4 gas chromatography (GC) ﬂame spectrometry 131–2 inductively coupled plasma-mass spectroscopy 159 lead 1092, 1097–100, 1101– 2, 1107– 8 metal(loid)s 526–8 solid phase microextraction 380– 7 volatile metals 1227, 1228 gas chromatography-mass spectroscopy (GC-MS) 1017–20 gas diffusion 641– 3 gas liquid separators 635– 6 gas phase reactions 199, 213– 6 gas scrubber sampling 711 –2 gas separators 635–6, 641 –2 gas streams 512 –7, 518 gases atmosphere 495, 496 ﬂue 521, 522 incineration 1224, 1227, 1229 landﬁll 522– 3, 524, 1224–7 sewage 522–3, 524, 1224–7 volatile metals 1212–4, 1216 gasoline 1081, 1085–88, 1102 gastropod populations 993 GC see gas chromatography GC-MS see gas chromatography-mass spectroscopy GD-MS see glow discharge mass spectrometry gel ﬁltration 1194– 5 geochemistry 605, 723 –58 geological samples chromium 1147–53 dissolutions 755 –8 molten salt fusion decomposition 750– 3 ores 290–1 preconcentration 753– 5 preparation 728–53 separation procedures 753–5 sequential extractions 755– 8 germanium 877, 892, 893 GF-AAS see graphite furnace atomic absorption spectroscopy 1267

Gilson’s solid phase extraction 658 glacial ice cores 920 glass disks 734–5 glass vessels 271 glassware cleaning 713 glassy carbon 25 glossaries 18–19, 931 –2 gloves 971 glow discharge mass spectrometry (GD-MS) 171–6 applications 175– 6, 660 atomization 173 ionization 173–5 processes 174 quantiﬁcation 173 –4 goals, deﬁning 5– 6 gold 730 gold-based drugs 1180– 1 grain 1051, 1054 graphite furnace atomic absorption spectroscopy (GF-AAS) 486, 488, 938, 1138 gravimetric method 913 gravity separators 619, 620 –1 Grignard reactions 578 –9, 582– 3, 587, 1004– 5 Grignard reagents 1097– 100 grinding 346– 7, 726–7, 769, 1248–9 ground water 869, 873, 876 hafnium 893 hair 1033, 1153–5 halogenated metal(loid)s 497, 1215 handling, general principles 14–7 hands 667 harbor sediment 1006– 8 hard-soft acid–base (HSAB) nature 463–5 headspace 1216, 1220, 1223, 1227 headspace extraction 373– 5 health effects 920 –1 heating 244, 260 –3, 282 heavy metals transport 919–21 herring gull eggs 1050 heterogeneity 931 –2 HG-AAS see hydride generation atomic absorption spectroscopy 1268

high... molecular mass (HMM) analysis 1048–9 molecular organoselenium 1203 performance liquid chromatography (HPLC) 131– 2, 148, 1143–5 pressure acid digestion 740 –3 pressure ashing (HPA) 206 –7 purity materials 481 –2, 488– 90 temperature dry ashing 694 temperature extractions 288– 9 volume aerosol samplers 918–22 historical overviews leaded gasoline 1085–8 microwave devices 257 –8 sequential extraction 1233–4 ultrasonic extraction 356– 8 wet digestion 193–4 HMM see high molecular mass homogeneity 931– 2 homogenization 38–9, 304– 5, 769, 1188– 90 homoscedasticity 61 hood cleaning 715 Hostaﬂon 25, 28 hot plate digestion 943 –4 HPA see high pressure ashing HPBI see 3-phenyl-4-benzoyl5-isooxazolone HPLC see high performance liquid chromatography HSAB see hard-soft acid– base hulls 991–1020 humans arsenic 1031– 3 leaded gasoline 1087–8 hybrid manifolds 658–9 hydride generation 386, 579 –80, 584–5, 588 hydride generation atomic absorption spectroscopy (HG-AAS) 249, 938 hydrides 497, 865, 877–9 hydrocarbons 711 –12, 914 –5 hydrochloric acid 196–7, 737, 981 hydroﬂuoric acid 736, 740, 742–3, 981 hydrogen peroxide 218–19, 981 hydroxyquinoline resins 864–5

8-hydroxyquinoline resins 864 –5 hygiene 360 –2, 520, 521, 1218 hyphenated techniques see also main names chromium 1134– 5, 1138, 1142–3 lead 1093– 6, 1109–11 ice cores 920 ICP see inductively coupled plasma ICP-AFS see inductively coupled plasmaatomic ﬂuorescence spectrometry ICP-MS see inductively coupled plasmamass spectrometry ICP-OES see inductively coupled plasma optical emission spectrometry ICP-QMS see inductively coupled plasma quadrupole mass spectrometry ID see isotope dilution IMH[trans-RuCLU4u(DMSO)Im] 1179, 1180 iminodiacetate-type chelating resins 430–1 immobilization 541–6 in aliginate 541, 542, 551, 555 on controlled pore glass 545–6 on silica gel 412, 445, 554, 1124 on sol-gel 554 impactors 909–16 in situ derivatization 1097– 100 in-torch vaporization (ITV) 151 INAA see instrumental neutron activation analysis incineration gases 1224, 1227, 1229 increments, statistics 8– 12 individual element preconcentration 865, 874–6 inductively coupled plasma-atomic emission spectroscopy (ICP-AES) 1142 inductively coupled plasma-atomic ﬂuorescence spectrometry (ICP-AFS) 858 inductively coupled plasma (ICP) 404 –9, 594, 596, 600–4 inductively coupled plasma-mass spectrometry (ICP-MS) chelation solvent extraction 487, 491

chromium 1120 dry ashing 235, 249 ﬂow injection solid phase extractions 630– 1 geological samples 728–9 industrial waste 938 quantitation 152– 60, 161 semiconductors 966, 968 water 858 inductively coupled plasma optical emission spectrometry (ICP-OES) chelation solvent extraction 477, 486, 489– 90 dry ashing 235, 249 ﬁgures of merit 152 ﬂow injection solid phase extractions 630– 1 geological sample preparation 729 industrial waste 938 instrumentation 143–52 interferences 142 –3 quantitation 142– 52 sample introduction 127–35, 146– 54, 158– 9, 163 theory 142 water 858 inductively coupled plasma quadrupole mass spectrometry (ICP-QMS) 966, 968 ﬁgures of merit 160, 161 instrumentation 154–6 interferences 156 –8 reaction cells 155 sample introduction 127–35, 146– 54, 158– 9, 163 industry ambient particulate matter 921– 2 chromium 1117–8, 1145– 7 hygiene 360–2, 521 –2 volatile metal(loid)s 520–3 waste 935–61 digestions 938– 50 leach procedures 950 –6 safety 936 sample preparation 935–61 1269

toxicity characteristic leaching procedure 951– 6 types 935–6 inert atmosphere 1248– 9 inhaled particles 917 –8 inorganic... arsenic 1032–4 based sorbents 411 –21 cations 397–8 compounds 343 –52 elements 914–5 oxides 411– 21 samples 285– 6 solid samples 481–2, 488 –90 instrumental neutron activation analysis (INAA) 729, 769 instrumentation accelerated solvent extraction 345 –6 atomic ﬂuorescence spectrometry 162 contamination/losses 34–5 cryogenic trapping 520 –8 ﬂame spectrometry 119– 21 ﬂow injection liquid– liquid extractions 617– 22 inductively coupled plasma-atomic optical emission spectroscopy 143–52 inductively coupled plasma –mass spectroscopy 154–6 lithium borate fusion 305 sample preparation 693 –713 solid phase microextraction 388 supercritical ﬂuid extraction 316–8 ultrasonic extraction 355–6 wet acid digestion 220, 223 X-ray ﬂuorescence spectrometry 177 interferences atomic ﬂuorescence spectrometry 163 electrothermal atomic absorption spectrometry 137– 8 ﬂame spectrometry 121 –3 inductively coupled plasma-atomic optical emission spectroscopy 142–3 inductively coupled plasma-mass spectroscopy 156–8 1270

ultraviolet/visible spectrophotometry 180 internal calibration 99, 603 internal standardization 51, 79– 81, 82 International Organization for Standardization (ISO) 94, 102, 110, 857, 917 –8 International Union of Pure and Applied Chemistry (IUPAC) 194, 459, 1234 Internet 960– 1 ion exchange 399 –400, 753–4 ion pairing 399 ion selective electrodes (ISE) 185– 6 ion traps (IT) 1095 ionic conduction 261 ionization 122, 173– 4, 175 IPA see isopropanol iron 443 –4, 553– 4, 886–90, 893 ISE see ion selective electrodes ISO see International Organization for Standardization isobaric interference 157–8 isolators 263, 268 isopropanol (IPA) 982 isotope dilution (ID) 51–2, 81–90, 389 isotopes chromium 1115 ratios 84–90 tin 1019 IT see ion traps ITV see in-torch vaporization Japanese National Institute for Environmental Studies (NIES) 1010, 1017 JM216 anti-cancer drug 1174–5, 1178–80 judgment samples 3 kinetics 331 Kjeldahl 836, 837, 844 Kjeldahl digestion 698– 9 knotted reactors 626, 628 Kraus-Callis process 1086 LA see laser ablation lab-on-valve (LOV) 655

laboratory reference materials (LRMs) 100, 104 lake waters 867 laminar ﬂow benches 29, 32, 33 landﬁll gases 522–3, 524, 1224–7 Langmuir isotherms 508 lanthanides 329–30 laser ablation (LA) 593–606 applications 604– 5 calibration 594, 601– 3 detection systems 599–601 experimental systems 594–9 fractionation 594, 603– 4 geological samples 732– 3 inductively coupled plasma-atomic optical emission spectroscopy 150–2 laser induced breakdown spectrometry (LIBS) 594–5, 599–601, 604 LC see liquid chromatography leaching automation 659 –60 contamination/losses 36 industrial waste 950– 6 metallobiomolecules 1189– 90 methylmercury extraction 1072–3 microwave extraction 286– 7 lead alloy reactions 1091 biogeochemical cycle 1092 compounds 339 drugs 1182 dust materials 1107–9 food 765– 856 industrial waste 946– 7 leaded gasoline 1085– 8 organoleads 339, 1083–5, 1088–111 in paints 364, 365 sample preparation 1081–111 solid phase microextraction 381–5 speciation 1081– 111 toxicity 1083– 5 water 877, 888, 889, 892, 893 leafy plants 1051–3 legislation 993, 1087

LIBS see laser induced breakdown spectrometry ligands concentrations 473– 4, 492 solid phase extraction 398, 436 supercritical ﬂuid extraction 318– 40 limits of detection (LODs) atomic ﬂuorescence spectrometry 163 chromium 1120 electrothermal atomic absorption spectrometry 138–9 ﬂame spectrometry 125–6, 128, 129 inductively coupled plasma-atomic optical emission spectroscopy 148– 9 inductively coupled plasma-mass spectroscopy 160, 161 ultraviolet/visible spectrophotometry 180 linear dynamic ranges 378 linear regression 55–67 lipids 1187– 8 liquid chromatography (LC) inductively coupled plasma-mass spectroscopy 159 lead 1092, 1102 –3 metallobiomolecules 1195 –6 solid phase extraction 404 –6 liquid –liquid extraction (LLE) 403, 615– 26 liquids automated sample preparation 653–9 industrial waste 937 semiconductor chemicals 966 volatile metals 1212 literature 766–8, 1239–40 lithium borates 301–10, 750–2 liver homogenates 1050 LLE see liquid– liquid extraction LMM see low molecular mass loading samples 433 –7, 629– 30 loam 1148 lobaplatin 1174, 1175, 1178 location, aerosol sampling 906–7 LODs see limits of detection losses of the analyte 23–42 1271

low... ﬂow rate samplers 923 molecular organoselenium 1201–3 pressure acid digestion 738– 9 volume microwave digestion 208 low molecular mass (LMM) analysis 1047– 8, 1050 low temperature ashing (LTA) 239, 694, 788–94, 836 LRMs see laboratory reference materials LTA see low temperature ashing lube oils 710–1 luminescence see chemiluminescence lung ﬂuids 1136, 1139, 1143– 4 lyophilized 1203 magnesium 892 magnetron tubes 263, 264–6 mammals 1046–50, 1189 manganese 887, 888, 890, 892, 893 manifolds automated sample preparation 658– 9 ﬂow injection gas diffusion 642 –3 ﬂow injection liquid-liquid extractions 624–6 ﬂow injection solid phase extractions 630–5 ﬂow injection vapor generation 638–41 microwave-heated ﬂow-through digestion 212 manipulators, robotics 663 –4 manure 946 marine ecosystems 991–1020, 1028–9, 1039– 40, 1200–1 masking 470– 2 mass ablated 602–3 mass discrimination 85– 7 mass ﬁltration 154–5 mass spectrometry (MS) chromium 1124– 30 lead 1106–7 organotin compounds 1017–20 materials see also certiﬁed reference...; reference... categories 99– 100 1272

characterization 102 losses/contamination/losses 24–7, 29 microwave dissipation factor 269 microwave laboratory equipment 269 vessels 271– 2 matrices aerosols 903– 32 calibration selection 51 contamination/losses 41– 2 food analysis 765– 847 fractionation 1240 geological samples 723–58 industrial waste 935 –61 laser ablation 602 petrochemistry 683–719 semiconductors 965– 86 sequential extraction 1239–41 solid phase extraction 436 –7 water 857 –94 X-ray ﬂuorescence spectrometry 177– 8 maximizing X-ray intensities 308–9 MBT see monobutyltin measurements contamination/losses 42 organotins 1005–17, 1020 traceability 94–5 mechanical ﬁlters 970 –1 melting 304–5 membrane extraction 559– 74 chemical principles 566 experimental set-ups 570– 3 properties 566– 70 techniques 559 –66 membrane phase separators 619, 621 –2, 623 memory effects 526, 573 mercury aqueous solution chemistry 1063– 5 biotrapping 534, 554– 5 collection strategies 1065– 7 cryogenic trapping 504–9, 517, 518 derivatization 577– 92, 578–581, 583– 6 drugs 1181 industrial waste 941 –2 preservation 1067–8 quality control 1075–8

sample preparation 1063–79 solid phase extraction 445, 446–7 solid phase microextraction 381–5 speciation 1063– 79 storage 1067– 8 supercritical ﬂuid extraction 338–9 water 878, 879–80, 881–5, 887, 892 metal... based drugs 1173– 82, 1199 –200 biotrapping 533– 57 carbon bonds 1213–14 complexes 473–4, 1198, 1203–5 coordination 1187–8 direct extraction 350 extraction kinetics 331 in foodstuffs 1192 –3 hydrogen bonds 1213– 4 industrial waste 939, 940, 946–9 ions adsorption 474– 7 chelate extraction 459–92 membrane extraction 566 –7, 571– 3 mutual separation 467– 72 natural waters 472– 4, 477–80, 486–8 preconcentration 465 –70 separation 459– 92 supercritical ﬂuid extraction 318–40 vessel materials 31–2 preconcentration alternative to biotrapping 533–57 retention 538 –41 supercritical ﬂuid extraction 318–40 metallic chromium 1117– 8 metallic impurities 969 –71 metallobiomolecules analyte moiety 1188 analyte recovery 1188–93 error sources 1205–6 fractionation 1193 –5 homogenization 1188– 90 multi-dimensional liquid chromatography 1195– 6 partial degradation analyte recovery 1190–3 sample preparation 1185 –206

solid matrices 1200–5 speciation 1185 –206 unreactive buffer analyte recovery 1188– 90 metallodrug metabolites 1199 –200 metalloenzymes 1190, 1198 metal(loid)s aqueous derivatization 589 Grignard reactions 587 hydride generation 588 metallobiomolecules 1185 –206 metallothioneins (MTs) 1203–5 metathesis polymerization-based polymers 431 method of standard additions 52, 74–9, 978– 9 method validation 51, 72, 93 see also QC methodology biotrapping 541 –6 chromium 1120–57 derivatization 583– 5 dry ashing 244 –8 food analysis 771– 841 geological samples 728–53 microwave extraction 281 –5 organolead compounds 1092–111 sequential extraction 1245 solid phase extraction 410 –39 solid phase microextraction 375– 9 sonication 354– 7 toxicity characteristic leaching procedure 952–3 ultrasonic extraction 354– 7 volatile metals 1218–30 methylarsonic acid 1036– 7 methylarsonous acid 1035–6 methyllead 1081–2 methylmercury aqueous solution chemistry 1063– 5 artifactual formation 1075–6 biological tissue 1073–4 extraction 338–9, 1071–4 particles 1072– 3 sediments 1072– 3 soils 1072– 3 speciation 1063–79 1273

supercritical ﬂuid extraction 338–9 water 1071– 2 micelle formation 331 micro-oriﬁce uniform deposit impactors (MOUDI) 912 microanalytical techniques 732 –3 microcolumns 401, 627– 8 microorganisms 538–41, 1055 microporous membrane liquid-liquid extraction (MMLLE) 559– 60, 565–6, 568 –70 microprobe techniques 732–3 microwave... acid digestion geological samples 743– 4 wet acid digestion 199, 201 –2, 207–13, 215, 217 –8, 221– 2, 224 concentration 217, 297– 8, 355, 948, 1057 digestion acid 743– 4 industrial waste 945– 9 petrochemistry 699– 704 wet 818–29, 838, 845 –6 wet acid 199, 201–2, 207 –13, 215, 217–8, 221 –2, 224 dissipation factor 269 dry ashing 253– 4, 694 drying 38 energy transfer 258 –63 evaporation 297– 8 extraction 257– 99 advanced applications 297–8 methodology 281–5 sample types 285–97 vessels 270–5 ﬂow systems 211 –13, 222 food 818– 28 laboratory equipment 261– 9 sequential extraction 1249– 50 theory 258– 63 wet acid digestion 199, 201–2, 207–13, 215, 217 –8, 221– 2, 224 wet digestion 818 –29, 838, 845– 6 1274

microwave induced plasma-atomic emission spectrometry (MIP-AES) 164–5 milk 1046– 9, 1136, 1139 mineralization 237– 5, 697– 704 minerals 735– 50 MIP –AES see microwave induced plasmaatomic emission spectrometry mixed solid/liquid industrial waste 937 MMLLE see microporous membrane liquid-liquid extraction mobility 1212– 4 mode stirrers 268– 9 model-based sampling 12–3 modes 168, 438 modules 665 –8 MOE see Ontario Ministry of the Environment moiety 1188 molecular... chromium speciation 1157 ﬂuorescence detection 181 –3 spectrometry 1123, 1143–5 spectrophotometry 1130–6 weight fractionation 1193–5 molten salt fusion decomposition 750– 3 molybdenum 890 monitoring objectives 904– 6 monobutyltin (MBT) 996– 7, 1006 moulds 303, 305 MS see mass spectroscopy MTs see metallothioneins MULSPOT project 1009–16 multi-element electrothermal atomic absorption spectrometry 139– 40 multi-dimensional liquid chromatography 1195– 6 multi-element preconcentration 860, 863– 5, 866–73 multiple coatings 132 multiplicative interference 49 mushrooms 1031, 1054– 5 mussels 993, 999, 1008–9 mutual separation 467–72 n-butyl acetate (NBA) 983 N-methyl pyrrolidine (NMP) 983

NAA see neutron activation analysis Naﬁon membranes 517, 518, 526 NAMI-A, ruthenium-based anti-cancer drugs 1179, 1180 naphthas 314– 6, 432, 709 –10 National Research Council of Canada (NRCC) 1010, 1017 natural abundances 1019, 1117– 8 natural volatile species 497, 498–502 natural waters 857– 94 chromium 1121, 1125– 6, 1131 metal ions 472– 4, 477–80, 486–8 NBA see n– butyl acetate Nd:YAG lasers 594, 598– 9 nebulizers 123– 7, 146–8 Nernst equation 185 neutrality, fusion beads 305– 6 neutron activation analysis (NAA) 1120 nickel 1215, 1218 NIES see Japanese National Institute for Environmental Studies nitrates 982 nitric acid food analysis 843 –5 geological samples 736– 40 semiconductors 975, 981 wet digestion 196 –7 NMP see N-methyl pyrrolidine nomenclature, wet digestion 194 non-biotic matrices 334–5 non-constant sensitivity 72– 4, 78–9 non-polymeric sorbents 421– 32 non-spectral interference 49 non-wetting agents (NWAs) 303– 5 normal-phase sorbents 410 NRCC see National Research Council of Canada nuclear reactor coolant water 873 nugget effect 727–8 nuts 1054 OES see optical emission spectroscopy off-line extractions 316–7, 411– 21 oils 277– 8, 279–80, 294– 7 on-line... coupling 403– 10 cryogenic trapping 524 –5, 526– 8

digestion 643– 6, 829–32 electrothermal atomic absorption spectrometry 141 extractions 317– 8 photo-oxidation 646 pretreatments 612 onions 1053–4 Ontario Ministry of the Environment (MOE) 941 open vessel acid digestion 199 –203, 221, 225–6, 738– 9 digestions 199– 203, 221, 225 –6, 698– 700, 738–9 extractions 274– 5, 283 –5 furnace dry ashing 693–4 microwave extraction 274 –5, 283– 5 wet acid digestion 199–203, 221, 225– 6 operational deﬁnitions 108–10, 1243– 4 operational modes 246–8 optical absorption spectrophotometry 859– 60 optical emission spectroscopy (OES) 488, 1105– 6 see also inductively coupled plasma... optimal ﬂuxes 306– 7 ordinary linear regression (OLR) 55–67 ores 290– 1, 364 –5, 1147– 8 organic... extractions 291– 2, 295–6, 421 –32, 623, 706– 7 food analysis 771– 839 fractions 926 halides 1091 microwave extraction 291 –2 sample dilutions 707– 9 sample dissolutions 707–9 solvents 623, 706– 7, 983 sorbents 421–32 organoarsenic 1200–1 photo-oxidation 646 supercritical ﬂuid extraction 313, 314, 404, 660, 765, 1097, 1251 organoleads 339, 1083– 5, 1088– 111 organomercury compounds 338– 9 organometallic compounds accelerated solvent extraction 343– 52 1275

aqueous derivatization 589 cryogenic trapping 517, 518, 522– 3, 524 determinations 1093–6 enzymatic extraction 1190– 1 Grignard reactions 587 hydride generation 588 semiconductors 985 supercritical ﬂuid extraction 332–9 organoselenium 1056, 1201–3 organotin compounds 332– 8, 582–5, 991–1020 ORL see ordinary linear regression ormaplatin 1175, 1177 ovens cleaning 715 digestions 701–2 drying 37–8 overall standard deviation 2 overspiking 88– 90 oxaliplatin 1174, 1175, 1177, 1179, 1180 oxidation chromium 1118– 20, 1135– 6 dry ashing 240– 4 states 1118– 20 oxide fractions 926 oxidizing acids 195– 8, 794–809, 812– 7, 836–8 oxidizing reagents 218– 9 oxygen combustion 704 –5, 788– 94, 835 oysters 363–5, 993, 999, 1009– 10 P-XRF see portable X-ray ﬂuorescence packed microcolumns 627 –8 paints 364– 5, 991–1020, 1118 partial acid attack 744– 5 partial degradation analyte recovery 1190–3 partial extractions 286– 7 particles 903 –32, 1072– 3 particulates 908, 915–30, 969–71 passive samplers 924 pentylation 1108 perchloric acid 196– 7, 736, 742 –3, 843– 5 percolation, solid phase extraction 433– 4 performance-based measurement system 857 1276

Periodic table 83 peripherals 665 –8 permeation drying 515–7 permittivity 261 –2 personal cascade impactor sampler (PCIS) 915– 6 pesticides 991 petrochemistry sample preparation 683– 719 acid mineralization 697–704 ashing 693– 7 cleanliness 713– 5 composition properties 685–6 physical properties 685–6 quality assurance 715 –8 trace element sensitivity 684, 687– 9 wet oxidation 697– 704 PGEs see platinum-group elements pH 434–6, 438 pharmaceutical ﬁelds 674, 676 phase segmentors 617–9 phase separation 613, 619– 22 3-phenyl-4-benzoyl-5-isooxazolone (HPBI) 489 phenyl-tin compounds 991– 1020 phenyltion 581– 3 phenylmercury 1075 phosphine 504– 9, 511 –3, 519 phosphoric acid 737–8, 982 phosphorus 350– 1, 943–4 photographic waste water 874 photolysis 203 photoresists 983–4 physical properties 685–6, 1088–90 physical separations 706 physico-chemical principles 504– 9 phytoremediation 556 pine needles 363– 4 pipettes 973–4 planning sampling operations 5 –8 plants arsenic 251–2 biotrapping 546 –8, 549– 50 chromium 1154–5 dry ashing 243 –4, 252 mercury 1068 metallobiomolecules 1188

organoselenium 1201–3 selenium 252, 1051–5 plasma 1046– 9, 1136, 1139, 1143 –4 plasma ultraﬁltrate (PUF) 1174–80 plastic containers 34– 5 platinum based drugs 1173– 80 group elements (PGEs) 730 vessels 245–6, 247 water 890 PM2.5 911, 921, 929 PM10 911, 921, 929 poisons 1027 see also toxicity polarography 1120 pollution 904 –6 poluacrylonitrile-based resins 431 poly-silicon 985 poly(acrylamidoxime) resins 489–90 polyacrylate polymers 423 polyamide polymers 430 polyatomic ions/isobars 83 polycyclic aromatic hydrocarbons 914– 5 poly(dithiocarbamate) resin 487, 489 –90 polyethylene 25, 430 poly(iminodiacetate) resin 487, 491 polymeric sorbents 421– 32 polymers cleaning procedures 26–7 dissolutions 709 ﬂuorinated polymers 25 ﬂuoropolymers 713–4 metathesis polymerization 431 polyamide 430 polytetraﬂuoroethylene 25, 27– 8, 430 polyurethane 423 sample preparation 683 –719 solid phase extraction 423, 431 vinylpyrrolidine 423 polypropylene 25 polystyrene 422–3, 424 –9, 430 polytetraﬂuoroethylene (PTFA), contamination 25, 27–8 polytetraﬂuoroethylene (PTFE) polymers 25, 27– 8, 430 polyurethane 423 ponds 868

Porapak adsorption 1227, 1229 porcelain vessels 245 portable X-ray ﬂuorescence (P-XRF) 730 –2 potable water 875 potassium 892 powder pellets 734 powders 733– 4 power density 355 microwave equipment 267, 275– 6 optimization feedback 280 –1 precipitation 754– 5, 1130 precision calibration 48, 70– 2 ﬂame spectrometry 125 internal standardization 80–1 isotope dilution mass spectrometry 88–90 method of standard additions 76– 8 solid phase microextraction 379 traceability 95 preconcentration chromium 1124, 1127 –8 contamination/losses 41– 2 ﬂow injection gas diffusion 642– 3 geological samples 753–5 metal ions 465 –70 volatile metals 1217–22 water 860 –77 preservation 1 –19, 403, 517 –9, 1067–8 pressure accelerated solvent extraction 348 digestion ﬂow systems 211– 3 food analysis 812–7, 836 –8 geological samples 738 –42 petrochemistry 700–4 wet acid 204 –10, 211– 3, 223– 4 liquid extraction 1038 microwave extraction 275 –8, 282– 3 supercritical ﬂuid extraction 336 pretreatments chromium 1123, 1137 –8, 1157 ﬂow injection 611– 46 food analysis 768– 832 1277

lead 1097– 100 sequential extraction 1246– 50 primary ions 167 primary methods 98, 104 process monitoring 605 proﬁciency testing 105–6 propylation 581, 1107–8 propylene glycol monomethyl ether acetate 983 propylenediaminetetraacetate chelating resins 431 proteins 834, 842 –3 protocol sampling 2 PTFE see polytetraﬂuoroethylene published papers 768, 1239– 40 pure water 974–5 QMF see quadrupole mass ﬁlters quadrupole ion traps 1095 quadrupole mass ﬁlters (QMF) 1095 quality assessment 17–8 quality assurance 17–8, 715 –8 quality control general principles 17 laboratory reference materials 100, 104 mercury 1075– 8 sequential extraction 1252 wet acid digestion 216–9 quality of data collection 13– 4 quantiﬁcation detection techniques 117– 86 glow discharge processes 173– 4 secondary ion mass spectrometry 168–70 quantitation adsorptive stripping voltammetry 184–5 anodic stripping voltammetry 183–4 atomic absorption spectrometry 164– 5 atomic emission spectrometry 164 –5 atomic ﬂuorescence spectrometry 160–5 cathodic stripping voltammetry 184 –5 chemiluminescence detection 181– 3 differential pulse anodic stripping voltammetry 183– 4 direct current plasma 165 1278

electrochemical methods 183 –6 electrothermal atomic absorption spectrometry 133–41 ﬂame spectrometry 118–35 glow discharge mass spectrometry 171– 6 inductively coupled plasma-atomic optical emission spectroscopy 142– 52 inductively coupled plasma-mass spectroscopy 152– 60, 161 ion selective electrodes 185– 6 microwave induced plasma-atomic emission spectrometry 164 –5 molecular ﬂuorescence detection 181– 3 secondary ion mass spectrometry 165– 70 ultraviolet/visible spectrophotometry 179– 81 voltammetry 183–5 x-ray ﬂuorescence spectrometry 176 –8 quartz 971, 972, 985 quartz vessels 271 racks 667 radial plasmas 144–5 radio frequency (RF) generators 143 radioactive isotopes 1115 radiotracers 251– 3 ramp 1053–4 random errors 48 random samples 4 rare earth elements 943– 4 rat liver homogenate 1050 ratios 84–90, 461–3 reaction cells 155, 859 reaction vessels 263 reactivity, chromium 1115– 6 reactor coolant water 873 readsorption 1244– 5 reagents contamination sources 27–8, 29, 30, 34 lead speciation 1109 semiconductors 974– 5 sequential extraction 1235, 1236–7 wet digestion 195–8

record keeping 937 recovery ashing 696–7 calibration 48– 52 data statistics 52–5 environmental organotins 1002– 4 metallobiomolecules 1188– 93 red cells 1046–9 redistribution, sequential extraction 1244– 5 redox reactions 1135– 6 reduction 1115– 7, 1123–4, 1127, 1129–30, 1133–5, 1136, 1138, 1155, 1157 reference materials (RMs) 93–114, 957, 1077– 8 see also certiﬁed... reference methods 94, 97– 9, 767 reﬂected energy 263, 267 –8 reﬂuxing digestion 698–9 regression 55– 67 relative permittivity 261– 2 relief sensors 273 renal tissue 1154 representative sample 3, 19 reseal vessels 274 residue dissolution 240–4 resistant mineral decomposition 745 –50 respirable fractions 916– 8 retention 1128 –9 reverse micelle formation 331 reversed-phase sorbents 410 reviews 767–8, 1239–40 RF see radio frequency rheumatoid arthritis 1180 –1 ring-opening metathesis polymerizationbased polymers 431 rivers 869, 872, 875 RMs see reference materials road dust 1108–9 robotics 660 –76 ruthenium 947 safety 223–4, 272– 3, 936 saline water 874 salt fusion decomposition 750– 3 Salvarsan (arsphenamine) 1027 sampler bias 937

sampler location 906–7 samples compatibility 707–9 component separations 706– 7 composite 5, 8, 18, 19 composition inﬂuence 246 industrial waste 936 –8 introduction 127–35, 146–54, 158–9, 163 judgment samples 2, 3 lithium borate fusion 302–4 loading 433– 7, 629–30 microwave extraction 285 –97 preparation accelerated solvent extraction 343– 52 aerosols 903– 32 arsenic 1027– 41 automation 649–77 chromium 1115– 58 crude oils 683–719 food analysis 765–847 geological samples 723 –58 industrial waste 935 –61 lead 1081–111 mercury 1063–79 metal-based drugs 1173–82 metallobiomolecules 1185– 206 petroleum products 683–719 polymers 683–719 sediments 723 –58 selenium 1045– 59 semiconductors 965– 86 soils 723–58 sonication 353– 66 volatile metals 1211– 30 preservation 517, 519 random samples 3, 4, 11, 12 subsamples 5, 6, 9, 10, 19 systematic 2 –4, 11 volume optimization 376–7 sampling aerosols 903– 34 contamination control 5, 39 containers 5, 6, 14, 16– 8 cups 129– 30 equipment 3, 6, 8, 14, 16 1279

handling 4, 13 model-based 12 organotin compounds 991– 1020 plans 5– 8 preservation 1, 14 sample quality 12, 17 scrubber 711, 712 statistical 2, 4, 8, 9, 11, 12, 17 storage 3, 6, 13, 16, 17 strategy 2, 11 temperatures 504– 9 transport 3, 5, 14, 16 uncertainty 2 variability 2 Sartorius 670 Saville, Russ 257 scrubber sampling 711 –2 sea water 859, 866–76 seafoods 1028–9 ﬁsh 584–5, 1050–1 mussels 993, 999, 1008– 9 oysters 363 –5, 993, 999, 1009– 10 seasonal ﬂuctuations 997–8 secondary ion mass spectrometry (SIMS) 165–70 practical principles 167 sensitivity and quantiﬁcation 168 yield 88 sediments aqueous derivatization 584–5 arsenic 1040 chromium 1147– 53 industrial waste 946– 7 mercury 1068 methylmercury 1072– 3 microwave extraction 287– 8 organotin compounds 995– 6, 1006– 10 sample preparation 723 –58 selenium 1058– 9 segregated populations 10– 1 selected ion monitoring (SIM) 1019–20, 1095 selective extractions 757–8 selectivity calibration 48– 52 elution solvents 438 membrane extraction 567 –8 1280

sequential extraction 1242–3 solid phase extraction 403 selenium biotrapping 555 –6 birds 1050–1 derivatization 577– 92, 578–581, 583– 6 dry ashing 248 –53 environmental samples 1055– 9 ﬁsh 1050–1 mammals 1046– 50 microorganisms 1055 organoselenium 1056, 1201– 3 plants 1051–5 sample preparation 1045– 59 solid phase extraction 445, 448 solid phase microextraction 381– 5 speciation 1045–59 urine 1197–8 water 878, 888, 893 semi-closed systems 214 semiconductors 176, 965– 86 sensitivity calibration 51–2, 72–4 coefﬁcients 53–7 factors 170 method of standard additions 78– 9 secondary ion mass spectrometry 168– 70 x-ray ﬂuorescence spectrometry 178 sensors, robotics 664 separation contamination/losses 41– 2 factors 467–70 ﬂow injection gas diffusion 641– 3 geological samples 753–5 lead 1101– 4 petrochemistry 706–7 sequential extraction aerosols 924– 30 BCR procedure 1238, 1239, 1251 certiﬁed reference materials 1239, 1252 deﬁnition 1234, 1243–4 enzymatic 1192– 3 extraction agents 1235, 1236– 7 fractionation 1234, 1240, 1241–2 geological samples 755–8 matrices 1239– 41

methodology 1245 microwave extraction 290– 1 operational deﬁnition 1243– 4 quality control 1252 sample dissolution 1233–4 speciation 1233– 52 supercritical ﬂuid extraction 1251 Tessier method 1235, 1238–9 toxicity characteristic leaching procedure 954– 6 ultrasound 1251 sequential injection 639 –41 serial automation 651–2 serum chelation solvent extraction 491 chromium 1136, 1139, 1143– 4 contamination/losses 36–7 selenium 1046– 9 sewage 522, 524, 1149, 1224– 7, 1241 SFC see supercritical ﬂuid chromatography SFE see supercritical ﬂuid extraction SGLP see synthetic groundwater leaching procedure sharp cut cyclones (SCC) 922 SI see Syste`me International sieving 1248–9 silica gel 411–2, 415 –21, 542 –3 silica vessels 245 silicates 287 –8, 926 silicon wafers 965, 968, 984 –5 silver 946– 7 SIM see selected ion monitoring SIMS see secondary ion mass spectrometry sintering 752 size exclusion chromatography (SEC) 1194–5 slags 290–1 SLM see supported liquid membranes slotted tube atom traps (STAT) 130 sludges 287– 8, 945, 1147 –53, 1241 slurries 127, 132– 5, 829–32, 838–9 small deposit area impactors 911– 2 Smith-Hieftje correction system 123 sodium... 892 borohydride 588 oxylate 751 tetraethylborate 580– 1

tetraphenylborate 581– 3 tetrapropylborate 581 software 668 soils arsenic 1031, 1040 chromium 1147–53 mercury 1068 methylmercury 1072–3 microwave extraction 287 –8 sample preparation 723–58 selenium 1058–9 sol-gel 543 –5 solid clinical samples 1153– 5 solid matrices 334 –5, 1058–9, 1200–5 solid phase extraction (SPE) 393– 451 applications 439–50 automation 658 chelating resins 474– 477 ﬂow injection 626– 35 methodology 410– 39 petrochemistry 707 sequential extraction 1235, 1236–7 theory 393–410 solid phase microextraction (SPME) 371– 90 coatings 334, 375–78, 388, 389, 926 extraction modes 373–5 gas chromatography 380– 7 investigative tools 388 isotope dilution calibration 389 limitations 388–9 methodology 375– 9 volatile metals 380 –7, 1227 solid samples automation 659–60 chromium 1145–57 digestion 976– 8 electrothermal atomic absorption spectrometry 138 ﬂow injection digestion 645 geological samples 733–5 industrial waste 942 –3 metal ions 329 solid sorbents 410– 32 solid wastes 937, 1240–1 solid-state lasers 598– 9 solubility 313–6, 318, 326– 8 1281

solvents compatibility 707 –9 consumption 570 extraction accelerated 347 –8 geological samples 754– 5 metal ions 459– 92 petrochemistry 706– 7 solid phase 433, 437 –9 polarity index 708 sensors 273 volumes 439 sonication 353–69 applications 358– 65 biological tissues 363– 4 environmental 358–60 industrial hygiene 360– 2 methodology 353–69 sorbents 395– 403, 410–32 sorption media 627–9 space charge interferences 157 spark source mass spectrometry (SS–MS) 175 SPE see solid phase extraction speciation aerosols 922–4 arsenic 349 –50, 1027–41 biotrapping 548, 551–6 chelation solvent extraction 459– 94 chromium 1115– 58 crude oils 686, 690– 1 cryogenic trapping 495 –529 electrothermal atomic absorption spectrometry 141 lead 1081– 111 mercury 1063– 79 metallobiomolecules 1185– 206 organotin compounds 991– 1020 reference materials 93, 94, 99– 105 selenium 1045– 59 sequential extraction 1233– 52 solid phase microextraction 373, 380 tin 991– 1020 volatile metals 1211– 30 specimen banking 37, 105 spectral interference 48– 9, 122 spectroﬂuorimetry 1133 1282

spectrophotometry 1120–1, 1123 –4, 1130, 1135, 1138, 1157, 1158 spectroscopic interference 157–8 spices 1053–4 spike recovery 976– 7, 1076– 8 spiked butyl rubber 696– 7 spiking isotope dilution mass spectrometry 83–4, 88– 90 organotin compounds 1002–4 recovery data statistics 52 SPME see solid phase microextraction spray chambers 146 –8 spring water 869 sputtered secondary ions 165 SRMs see standard reference materials SS –MS see spark source mass spectrometry stability arsenic compounds 1033–8 constants 473–4, 492 cryogenic trapping 517, 519 metallobiomolecules 1206 organotin compounds 998 –1000 volatile metals 1212–4 stable emulsions 709 –11 standard additions calibration 52, 74– 9, 978– 9 standard deviation 2 standard reference materials (SRMs) 957 standard uncertainties 53 standardized procedures 96– 7 standards fusion 309 semiconductors 974– 5 STAT see slotted tube atom traps stated references 93–114 stated uncertainty 95 static secondary ion mass spectrometry 165–6 statistics 8 –14, 52– 5 steaming 26– 9 steel industry 1118 storage contamination/losses 35– 7 general principles 14–7 mercury 1067–8

organotins 998–1000 solid phase extraction 403 stratiﬁed waste piles 937 streams 867 strontium 892, 893 sub-boiling distillation 27–8, 29, 30 subsamples 5 substrates 535–8, 546 –8, 549– 50 sugars 292– 3, 1030, 1037– 8 sulfated ashing 694–5 sulﬁdes 308 sulfur 308 sulfuric acid 737, 982 supercritical ﬂuid chromatography (SFC) 313, 1103 supercritical ﬂuid extraction (SFE) 313– 40 instrumentation 316– 18 organometallic compounds 332–9 processes 313– 4, 332 sequential extraction 1251 supercritical ﬂuids 313–6, 326 –8 supported liquid membranes (SLM) 559–65, 569–73 surface analysis 176 sweat 37 synergism 466 –70 synthesis, organoleads 1090–1 synthetic groundwater leaching procedure (SGLP) 956 syringe barrels 400, 401–2 systematic samples 4 Syste`me International (SI) units 94, 96 TAL see trialkyllead tap water 874, 875 TBT see tributyltin TCLP see toxicity characteristic leaching procedure TeAL see tetraalkyllead Technicon autoanalyzers 656–7 Tedlar bags 1217 Teﬂon 25, 27–8, 430 bombs 207 –8, 701– 2 microwave extraction 271– 2 TEL see triethyllead temperature accelerated solvent extraction 348 cryogenic trapping 510

dry ashing 239 –42 microwave extraction 275 –80 microwave heating 262–3 solid phase microextraction 377– 8 supercritical ﬂuid extraction 336 volatile metals 1215 Tenax adsorption 1227, 1229 terminology, calibration 48–9 terrestrial environment/plants, arsenic 251–2, 1031 Tessier method 756–8, 1235, 1238– 9 test portion selection 727–8 tetraalkyl(aryl)borates 580 –3, 589 tetraalkyllead (TeAl) compounds 1081–111 tetramethylammonium hydroxide (TMAH) 982 tetranethylarsonium 1037 thallium 944 theory atomic ﬂuorescence spectrometry 162 chelate extraction 460– 74 derivatization 578– 83 ﬂame spectrometry 118–9 ﬂow injection liquid–liquid extractions 616–7 inductively coupled plasma-atomic optical emission spectroscopy 142 inductively coupled plasma-mass spectroscopy 153– 4 microwaves 258– 63 solid phase extraction 393 –410 supported liquid membrane extraction 562 –4 thermal desorption interfaces 387 thermally convective wet digestion 199, 200– 1, 204–7, 210 –1 thimersal 1181 time accelerated solvent extraction 349 microwave extraction 275 –6 optimization 377 time-of-ﬂight mass spectroscopy (TOF– MS) 1095–6 tin derivatization 577– 92, 578 –81, 583 –6 environmental samples 991 –1020 1283

European Union 1005–6, 1009–10, 1020 industrial waste 946– 7 isotopes 1019 solid phase extraction 445, 449–50 solid phase microextraction 381–5 speciation 991–1020 Tin speciation project 1005– 6, 1009– 10, 1020 water 892 tissue chromium 1154– 5 contamination/losses 34 mercury 1068 metallobiomolecules 1189 methylmercury extraction 1073–4 organoselenium 1203 selenium 1049– 50 titanium 717, 878, 890 titanium dioxide 287– 8 TMAH 982 TMAH see tetramethylammonium hydroxide TML see trimethyllead TOF-MS see time-of-ﬂight mass spectroscopy tools, losses/contamination 29 torches 143 –4 total... analyte content determination 236–54 fusion 750– 2 reﬂection geometry 178 trace element determinations 106– 8 total-reﬂection X-ray ﬂuorescence spectrometry (TXRF) 891–4 Total suspended particulate (TSP) 923 toxicity arsenic 1028–30 organolead compounds 1083– 5 organotin compounds 991 toxicity characteristic leaching procedure (TCLP) 936, 951–6 TPhT see triphenyltin trace element quantiﬁcation 117– 86 trace metal fractionation 924 –30 traceability 93– 114 1284

trans-D-1, 2-diaminocyclohexaneetetrachloroplatinum(IV) 1175, 1177 trans-L-1, 2-diaminocyclohexaneoxalatoplatinum(II) 1174, 1175, 1177, 1179, 1180 translocation 539 –40 transport 14– 7, 904–6, 919 –21 trapping biotrapping 556 –7 complete 562–3 cryogenic trapping 495–529, 1218, 1220–2, 1223–7 supported liquid membrane extraction 562 –3 water cooled atom trap 130 treatments see also pretreatments chromium 1155, 1157 classiﬁcation 770, 771– 2 organotin compounds 991 –1020 trialkyllead (TAL) 1081–2 tributyltin (TBT) 991–1020 triethyllead (TEL) 1081 trimethylarsine oxide 1037 trimethyllead (TML) 1081–3 triphenyltin (TPhT) 991–1020 trueness, traceability 95 TTA extraction 491– 2 turntables 268–9 TXRF see total-reﬂection X-ray ﬂuorescence spectrometry UE see ultrasonic extraction ultrahigh purity water 873, 966 ultraﬁltration 1194 ultrasonic extraction (UE) 353 –66 ultrasonic nebulization (USN) 858 ultrasound 202, 949, 1251 ultraviolet (UV)... digestion 199, 203, 209 –11, 221– 2 on-line decomposition 211 on-line photo-oxidation 646 ultraviolet visible (UV VIS) spectrophotometry 179 –81, 1130 –3 unattended operation see automation unbroken chains 94– 5 uncertainty 994

external calibration 68–70 general principles 5 internal standardization 80– 1 isotope dilution mass spectrometry 86– 8 linear regression 64– 7 method of standard additions 75–6 recovery data 53–4 underspiking 88–90 United States Environmental Protection Agency (US EPA) 354, 857–8, 939 –46, 949 –50 unreactive buffer analyte recovery 1188–90 uptake 538 –41, 1083 urban environments aerosols 913–14 ambient particulate matter 921 –2 dust 1108– 9 volatile metal(loid)s 520 –1 urine arsenic 1032, 1196–7 chromium 1136, 1139– 41, 1142– 4 contamination/losses 37 drug 1173 –83 selenium 1046– 9 US EPA see United States Environmental Protection Agency USN see ultrasonic nebulization UV see ultraviolet UV-VIS see ultraviolet visible Vacutainer 35 vanadium 718, 887, 891, 893, 1181 vapor generation atomic absorption spectrometry (VG-AAS) 638–41, 644–5 vapor generation (VG) 577– 90 applications 585– 9 chemical 127– 9, 140–1, 150, 163 ﬂow injection 635–41 geological samples 755 vapor-phase acid digestion 199, 213–6, 222 vapor-phase aerosol generation 593 vapor pressure 502 vapor steaming 26, 27

vaporization 150–2, 769, 859 variability in sampling 2 vegetables 1051, 1053 vegetation see plants vents 274 vessels dry ashing 245 –6 event detection 272 –3 materials 29, 31– 2, 195–8 microwave extraction 270 –5 relief 273 VG see vapor generation VG-AAS see vapor generation atomic absorption spectrometry vinylpyrrolidine copolymers 423 viscosity 314 –6 visible spectrophotometry 179 –81, 1130– 3 vitreous silica 24–5 VMCs see volatile metal compounds VOCs see volatile organic compounds volatile metallic compounds (VMCs) 495– 529, 1211 –30 adsorption 1218– 20, 1227–9 cryogenic trapping 1218, 1220–2, 1223–7 gases 1212–4, 1216 methodology 1218–30 sample preparation 1211– 30 solid phase microextraction 380– 7 speciation 1211 –30 volatile metalloid compounds 495– 529, 1211 –30 volatile organic compounds (VOCs) 495– 529, 1217– 8 volatile species 502 –3, 865, 877–9 volatilization 29, 526 –8, 540 volcanic activity 1031 voltammetry 183– 5, 885, 888 –91 volumetric ﬂasks 973 wafers 965, 966, 968, 984–5 washing 433, 630 waste industrial 935– 61 water 874, 876 water 857 –94 arsenic 1031– 3 1285

chromium 1120– 36 cryogenic trapping 512 –7, 518, 1227 direct analysis 876, 893 drinking 868, 1031–3, 1121 lead 1108–9 luminescence 880 mercury 883–5 metal ion extraction 477– 80, 486– 8 methylmercury 1071– 2 organotins 991–1020 preconcentration techniques 860 selenium 1056– 7 solid phase extraction 411 –32 T-XRF 891 vapor generation 879 volatile metals 1215, 1227 voltammetry 885 water cooled atom traps (WCAT) 130 water solubility of polysaccharides 1191–2 waveguides 263, 267 wavelength isolation 145– 6 waxes 294 –7 WCAT see water cooled atom traps wear metals 710–11 websites 960–1 weighted linear regression (WLR) 59–67 weighting factors 65 well water 874 well-mixed particulates 9 –10 wet acid digestion 199– 227 closed systems 204, 214– 6 efﬁciency 216– 9 ﬂow systems 210– 3 instrumentation 220, 223 microwave heating 199, 201, 207 open systems 4, 214, 739, 838 safety 202, 204, 223– 4 UV digestion 203, 209– 11, 221– 2 vapor-phase digestion 214–6 wet ashing 694–5 wet decomposition 40, 41 wet digestion 193 –227

1286

acid digestion 199 –227 contamination/losses 40, 41 decomposition 199 –227 dissolution 199– 227 food analysis 794– 809, 812 –29, 836– 8, 843–6 reagents 195– 8 techniques comparison 219 –20, 221 –3 vessel materials 195–8 wet oxidation 697– 704 wetlands 1058 wheat 1054 white clover 1053– 4 WLR see weighted linear regression workstations 662– 3, 664, 668–76 World Health Organization (WHO) 1118 World Wide Websites, industrial waste 960– 1 sample preparation 961 X-ray ﬂuorescence (XRF) see also T-XRF chromium 1120 fusion 301 geological samples 728–9, 730– 2, 734– 5 instrumentation 177 matrix effects 177– 8 quantitation 176– 8 trace analysis 178 X-ray intensity maximization, fusion and ﬂuxes 308–9 XRF see X-ray ﬂuorescence yeast 541 –3, 1201–3 yields 170 ZD0473 anti-cancer drug 1174, 1179, 1180 Zeeman correction system 123 zinc 888, 891 –3 zircons 745