Capillary Electrochromatography

J O U R N A L OF C H R O M A T O G R A P H Y LIBRARY - - v o l u m e 62 capillary e l e c tro ch ro rn a to g ra p h y...

Author: F. Svec | Z. Deyl

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J O U R N A L OF C H R O M A T O G R A P H Y LIBRARY - - v o l u m e 62

capillary e l e c tro ch ro rn a to g ra p h y

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J O U R N A L OF C H R O M A T O G R A P H Y LIBRARY -- v o l u m e 62

capillary electroch romato graph y edited by

ZdenOk Deyl Institute of Physiology, Academy of Sciences of the Czech Republic, Prague, Czech Republic

and

v

Fran ti#ek Svec Department of Chemistry, University of California, Berkeley, USA

2001

ELSEVIER Amsterdam

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London

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New

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E L S E V I E R S C I E N C E B.V. S a r a B u r g e r h a r t s t r a a t 25 P.O. B o x 211, 1000 A E A m s t e r d a m , T h e N e t h e r l a n d s

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F i r s t e d i t i o n 2001 L i b r a r y o f C o n g r e s s C a t a l o g i n g in P u b l i c a t i o n D a t a C a p i l l a r y e l e c t r o c h r o m a t o g r a p h y / e d i t e d b y Z d e n e k D e y l a n d F r a n t i s e k Svec. p. c m . - - ( J o u r n a l o f c h r o m a t o g r a p h y l i b r a r y ; v. 62) I n c l u d e s b i b l i o g r a p h i c a l r e f e r e n c e s a n d index. ISBN 0-444-50432-X (hardcover) 1. C a p i l l a r y e l e c t r o p h o r e s i s . 2. C a p i l l a r y l i q u i d c h r o m a t o g r a p h y . I. D e y l , Z d e n e k . II. Svec, F r a n t i s e k . III. Series. Q P 5 1 9 . 9 . C 3 6 C 3 4 2001 543'.0894--dc21 2001018989

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V

Preface Capillary electrochromatography (CEC) is a rapidly emerging technique that adds a new dimension to current separation science. The major "news" in this method is that the hydrodynamic flow of the eluting liquid, which is typical of HPLC, is replaced by a flow driven by electro-endoosmosis. This increases considerably the selection of available separation mechanisms. For example, combinations of traditional processes such as reversed-phase- or ion-exchange- separations with electromigration techniques are now possible. Also, CEC is opening new horizons in the separation of non-polar compounds, and thus represents an alternative to the widely used micellar electrokinetic chromatography. As a matter of fact, different separation procedures such as "classical" chromatography, capillary zone electrophoresis, isoelectric focusing, and isotachophoresis, are now merging to create a unified concept of separation science that was envisioned quite some time ago. This wealth of techniques also enables their combination in a variety of operational modes to meet the continuously growing requirements of separating ever-smaller samples (both in mass and volume), removing interferences from complex matrixes, and of making on-line identification of entities separated from mixtures. As with many other instrumental methods, an increase in the complexity of the method often leads to a reduction in reliability unless the technical background has reached a level of maturity (although this is difficult to define). Thus, capillary electrochromatography benefits largely from the use of packed columns. However, this packed column technology, originally developed for HPLC, does not meet completely the needs of CEC, and new approaches are desirable. One of those new techniques is the creation of the column-bed in situ by the preparation of monolithic separation media. Obviously, history repeats itself. Many scientists will remember that the columns current in gas chromatography have evolved from packed columns and that coated open-tubular capillaries only came later. The ingenious introduction of micellar/microemulsion electrokinetic chromatography by Terabe extended the possibilities of electromigration techniques into the area of uncharged compounds. A combination of these approaches is seen to emerge in electrochromatography. As a result, today's open-tubular electrochromatography exploits the interactions of solutes with the modified inner surfaces of the capillaries, a phenomenon that many researchers active

VI in electrophoresis (and, in particular, those in protein and peptide chemistry) attempted to avoid for a number of years. The fact that electrochromatography has overcome its typical "childhood problems" is proved by the steadily increasing number of applications, as confirmed in the last chapter of this book. Clearly, electrochromatography provides the analyst with a new tool which understandably, and as with many other instrumental analytical methods, is not generally applicable to all separation tasks. However, the number of difficult separations that has been achieved elegantly using CEC within recent years is, in our opinion, sufficient to justify publication of a monograph on this subject. Within the following pages, we have attempted to provide the reader with the necessary theoretical background, description of the instrumentation, modes of operation and methods of detection, and an overview of the broad variety of applications. A common problem of all monographs concerned with rapidly developing fields is that they cannot include everything. For example, the number of papers related to CEC published during the preparation of this book increased by several tens. Similarly, it is almost impossible to cover all aspects of the subject. While some facets of the current CEC are reviewed to an adequate depth, some others are only mentioned briefly. Our intention was to publish quickly a book that may be less complete, rather than endlessly improving a work that would never see the printer's press. Since virtually nothing is constant in this world but is continuously developing, it is a fair assumption that CEC will also be developed further in the very near future, and a new monograph might be required. Therefore, we would appreciate greatly any comments and suggestion from readers which will help us to improve future editions.

Frantigek ~;vec Zden6k Deyl

Berkeley and Prague December 2000

VII

List of Contributors Luis A. Col6n

Department of Chemistry, State University of New York at Buffalo, Natural Sciences Complex, Buffalo, NY 14260-3000, USA Anna Dermaux

State University of Gent, Organic Chemistry Department, Krijgslaan 281 (S. 4.), B 9000 GENT, Belgium Zden6k Deyl

Institute of Physiology, Academy of Sciences of the Czech Republic, Videhsk6 1083, CZ-14220 Praha 4, Czech Republic Melvin R. Euerby

AstraZeneca R&D Charnwood, Bakewell Road, Loughborough, Leicestershire, LE11 5RH, United Kingdom Adam M. Fermier

The R.W. Johnson Pharmaceutical Research Institute, Science and New Technology~Analytical Development, OMP Bld. B-236, 1000 Route 202, Raritan, NJ 08869, USA Georg H61zl

Institute of Analytical Chemistry and Radiochemistry, Leopold-FranzensUniversity, Innrain 52 a, A-6020 Innsbruck, Austria Csaba Horwith

Department of Chemical Engineering, Yale University, New Haven, CT, USA Christian G. Huber

Institute of Analytical Chemistry and Radiochemistry, Leopold-FranzensUniversity, Innrain 52 a, A-6020 Innsbruck, Austria Christopher M. Johnson

AstraZeneca R&D Charnwood, Bakewell Road, Loughborough, Leicestershire, LE11 5RH, United Kingdom Hiroshi Kobayashi

Department of Polymer Science and Engineering, Kyoto Institute of Technology, Matsugasaki, Sakyo-ku, Kyoto 606-8585, Japan Todd D. Maloney

Department of Chemistry, State University of New York at Buffalo, Natural Sciences Complex, Buffalo, NY 14260-3000, USA

VIII Maria T. Matyska

Department of Chemistry, San Jose State University, One Washington Square, San Jose, CA 95192, USA Alan P. McKeown

AstraZeneca R&D Charnwood, Bakewell Road, Loughborough, Leicestershire, LE11 5RH, UnitedKingdom Ivan Mik~ik

Institute of Physiology, Academy of Sciences of the Czech Republic, Videhsk6 1083, CZ-14220 Praha 4, Czech Republic Joseph J. Pesek

Department of Chemistry, San Jose State University, One Washington Square, San Jose, CA 95192, USA Anurag S. Rathore

Bioprocess Sciences, Pharmacia Corp., Mail Code GG3K, 700 Chesterfield Parkway North, Chesterfield, MO 63198 Gerard P. Rozing

Agilent Technologies GmbH, Waldbronn Analytical Division, P.O. Box 1280, D 76337 Waldbronn Pat Sandra

State University of Gent, Organic Chemistry Department, Krijgslaan 281 (S. 4.), B 9000 GENT, Belgium Volker Schurig

Institute of Organic Chemistry, University of Tidbingen, Auf der Morgenstelle 18, 72076 Tiibingen, Germany Franti~ek ~;vec

Department of Chemistry, University of California, Berkeley, CA 94720-1460, USA Nobuo Tanaka

Department of Polymer Science and Engineering, Kyoto Institute of Technology, Matsugasaki, Sakyo-ku, Kyoto 606-8585, Japan Dorothee Wistuba

Institute of Organic Chemistry, University of Tiibingen, Auf der Morgenstelle 18, 72076 Tfibingen, Germany

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . List of Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 1

1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8

V VII

Migration of Charged Sample Components and Electroosmotic Flow in Packed Capillary Columns ...................... Anurag S. Rathore and Csaba Horv~th

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F l o w o f ions in o p e n tubes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E l e c t r o o s m o t i c f l o w in o p e n tubes . . . . . . . . . . . . . . . . . . . . . . . . . . F l o w o f ions in p a c k e d c o l u m n s . . . . . . . . . . . . . . . . . . . . . . . . . . . Electroosmotic flow through packed columns . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S y m b o l s and a b b r e v i a t i o n s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 2

Instrumentation for Capillary Electrochromatography

........

1 2 3 5 6 12 35 36 37

39

Gerard P. Rozing, Anna Dermaux and Pat Sandra 2.1 2.2 2.3 2.4 2.5 2.6

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Instrumentation requirements . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gradient-CEC instrumentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S u m m a r y and c o n c l u s i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 3

Modes of CEC Separation . . . . . . . . . . . . . . . . . . . . . . . . .

40 41 58 75 82 83

87

Christopher M. Johnson, Alan P. McKeown and Melvin R. Euerby 3.1 3.2 3.3 3.4 3.5 3.6 3.7 3.8

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Definitions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Unmodified packings . . . . . . . . . . . . . . . . . . . . . . . . Modified packings . . . . . . . . . . . . . . . . . . . . . . . . . Chiral stationary p h a s e s . . . . . . . . . . . . . . . . . . . . . . Gel C E C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monoliths . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Size e x c l u s i o n C E C . . . . . . . . . . . . . . . . . . . . . . . .

3.9 3.10 3.11 3.12

Gradient CEC . . . . . . . . . . . . . . . . Selectivity c o m p a r e d with L C . . . . . . G u i d e l i n e s for the analysis o f acidic basic Conclusions . . . . . . . . . . . . . . . . .

. . . . . . . .

. . . . . . . .

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. . . . . . . .

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. . . . . . . .

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. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . and neutral c o m p o u n d s . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. . . .

88 89 89 94 100 101

101 102 103 103 104 106

3.13 3.14

106 106

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 4

Packed Bed Columns . . . . . . . . . . . . . . . . . . . . . . . . . . .

111

Luis A. Col6n, Todd D. Maloney and Adam M. Fermier 4.1 4.2 4.3 4.4 4.5 4.6 4.7 4.8

Introduction . . . . . . C o l u m n fabrication . . Packing methods . . . Comparison of packing Conclusions . . . . . . Acknowledgement . . Abbreviations . . . . . References . . . . . .

Chapter 5

. . . . . . . . . . . . . . . . . . . . . procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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

Capillary Electrochromatography on Monolithic Silica Columns

112 112 150 156

158 158 159 159

. . 165

Nobuo Tanaka and Hiroshi Kobayashi 5.1 5.2 5.3 5.4 5.5 5.6

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . M o n o l i t h i c silica c o l u m n s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preparation procedure o f silica m o n o l i t h s from silane m o n o m e r s . . . . . . . . . Structural properties o f m o n o l i t h i c silica c o l u m n s prepared in a capillary . . . . . P e r f o r m a n c e o f m o n o l i t h i c silica c o l u m n s in C E C . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 6

165 167

169 170 173

180

Capillary Column Technology: Continuous Polymer Monoliths . . . 183

Franti~ek Svec 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8 6.9 6.10 6.11 6.12

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184 R e p l a c e a b l e p o l y m e r i c stationary phases . . . . . . . . . . . . . . . . . . . . . . 185 P o l y m e r gels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 H i g h l y crosslinked a c r y l a m i d e - b a s e d m o n o l i t h s . . . . . . . . . . . . . . . . . . 189 I m p r i n t e d enantioselective m o n o l i t h s . . . . . . . . . . . . . . . . . . . . . . . 206 Polystyrene-based monoliths . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207 M e t h a c r y l a t e ester-based m o n o l i t h i c c o l u m n s . . . . . . . . . . . . . . . . . . . 212 A s s e s s m e n t o f p o r o u s structure . . . . . . . . . . . . . . . . . . . . . . . . . . . 220 Effects o f properties on the separation ability . . . . . . . . . . . . . . . . . . . 223 Other applications o f p o r o u s p o l y m e r m o n o l i t h s in C E C c o l u m n t e c h n o l o g y . . . 232 Acknowledgment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238

Chapter 7

Open Tubular Approaches to Capillary Electrochromatography

. . 241

Joseph J. Pesek and Maria T. Matyska 7.1 7.2 7.3 7.4 7.5 7.6 7.7

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C h e m i c a l etching process . . . . . . . . . . . . . . . . . . . . . . . . . . . . C h e m i c a l m o d i f i c a t i o n process . . . . . . . . . . . . . . . . . . . . . . . . . . C h a r a c t e r i z a t i o n o f etched, c h e m i c a l l y m o d i f i e d capillaries . . . . . . . . . . . . Applications of O T C E C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

242 . .

. . .

244 245 247 257 266 267

XI

7.8

References

Chapter 8

8.1 8.2 8.3 8.4 8.5 8.6 8.7

Hyphenation of Capillary Electrochromatography and Mass Spectrometry: Instrumental Aspects, Separation Systems, and Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Christian G. Huber and Georg HOlzl

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I n s t r u m e n t a t i o n and t e c h n o l o g y for c o u p l i n g o f C E C and M S .......... Stationary p h a s e - m o b i l e p h a s e systems used for C E C - - M S ........... O p t i m i z a t i o n o f e l e c t r o c h r o m a t o g r a p h i c and m a s s s p e c t r o m e t r i c c o n d i t i o n s Examples of application . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 9

9.1 9.2 9.3 9.4 9.5 9.6 9.7 9.8 9.9 9.10

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

268

271

.

272 273 296 . . . 298 . 305 . 312 . 313

Pressure Supported CEC: a High-Efficiency Technique for Enantiomer Separation . . . . . . . . . . . . . . . . . . . . . . . Dorothee Wistuba and Volker Schurig

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . T e c h n i q u e s o f pressurizing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Theory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Instrumentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A d v a n t a g e s and d i s a d v a n t a g e s o f pressurized C E C . . . . . . . . . . . . . . . . C o n c l u s i o n and future trends . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

317 318 319 321 325 326 335 336 337 337 337

Chapter 10 Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Zden~k Deyl and Ivan Mik~/k

341

10.1 10.2 10.3 10.4 10.5 10.6 10.7 10.8 10.9 10.10 10.11 10.12 10.13 10.14

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preconcentration procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ketones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Carbohydrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fatty acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Triglycerides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Steroids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A m i n o acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . N u c l e i c acids constituents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Drugs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Antibiotics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pesticides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

341 342 343 347 348 350 352 355 358 359 362 363 368 369

10.15 A v a i l a b l e applications ( s u m m a r i z i n g Table) . . . . . . . . . . . . . . . . . . . . 10.16 R e f e r e n c e s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

369 413

Index of Compounds Separated . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

421

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Chapter 1

Migration of Charged Sample Components and Electroosmotic Flow in Packed Capillary Columns Anurag S. RATHORE* and Csaba HORV/kTH

Department of Chemical Engineering, Yale University, New Haven, CT, USA *Present address: Bioprocess Sciences, Pharmacia Corp., Mail Code GG3K, 700 Chesterfield Parkway North, Chesterfield, MO 63198

CONTENTS

1.1 1.2 1.3 1.4

1.5

1.6 1.7 1.8

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Flow of ions in open tubes . . . . . . . . . . . . . . . . . . . . . . . . . . . Electroosmotic flow in open tubes . . . . . . . . . . . . . . . . . . . . . . . Flow of ions in packed columns . . . . . . . . . . . . . . . . . . . . . . . . 1.4.1 Conservation of electric current . . . . . . . . . . . . . . . . . . . . 1.4.2 Evaluation of conductivity of the packed and open segments . . . . . 1.4.3 Evaluation of the potential drop across the packed and open segments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4.4 Evaluation of the equivalent length of the packed segment . . . . . . 1.4.5 Evaluation of the electric field strengths in the two segments . . . . 1.4.6 Conductivity ratio in CEC . . . . . . . . . . . . . . . . . . . . . . Electroosmotic flow through packed columns . . . . . . . . . . . . . . . . 1.5.1 Overbeek's model for EOF in porous media . . . . . . . . . . . . 1.5.2 Effect of charged capillary wall . . . . . . . . . . . . . . . . . . . 1.5.3 Conservation of volumetric flow rate . . . . . . . . . . . . . . . . 1.5.4 Flow equalizing intersegmental pressure . . . . . . . . . . . . . . 1.5.5 Flow velocities in the packed and open segments . . . . . . . . . . 1.5.6 EOF inpacked columns . . . . . . . . . . . . . . . . . . . . . . . 1.5.7 Migration times, velocities and M-factors . . . . . . . . . . . . . . 1.5.8 Higher separation efficiencies in CEC . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Symbols and abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 3 5 6 6 6 7 8 10 11 12 12 14 18 20 23 26 27 32 35 36 37

2

Chapter 1

1.1 INTRODUCTION Capillary electrochromatography (CEC) is a liquid phase analytical separation technique that is carried out with capillary columns by electroosmotically driven mobile phase at high electric field strengths in an apparatus similar to that used in capillary zone electrophoresis (CZE). The history of CEC could be traced to almost 60 years back when Strain [1,2] applied electric field across an adsorption column to demonstrate higher selectivity due to combination of electrophoretic and chromatographic separation forces. Almost thirty years later in 1974, Pretorius et al. suggested the use of EOF as "pumping mechanism" alternative to pressure driven flow in order to expand the scope of the then newfangled technique of HPLC [3]. The viability of CEC in packed capillary columns was demonstrated by Jorgenson and Lukacs in 1981 [4] and examined in more detail by Knox and Grant in 1987 [5,6]. One of the primary reasons of interest in the technique is the mixed separation mechanism of CEC that is borrowed from both HPLC and CZE [7]. In order to exploit the potential of CEC, it is of utmost importance to successfully control and optimize electrochromatographic conditions to generate high EOF velocities. Several experimental and theoretical studies have been performed and have tried to address these issues [8-30]. However, theoretical understanding of the EOF and the associated transport phenomena in porous media such as packed columns in CEC requires an exact description of the flow field in porous media under conditions prevalent in CEC by simultaneously solving the Poisson-Boltzmann and NavierStokes equations, which remains an unsolved and complicated task [9,10,31-54]. Columns employed so far in capillary electrochromatography (CEC) contain both a packed and an open segments with concomitant changes of the electric field strength and the flow velocity at the interface of these two segments in such columns. Figure 1.1 shows the schematics of a CEC column of total length L, which consists of a

Packed Floating segment tretaining rit ~ ,t

Fixed retaining frit

:

Open Lopen se/~l

::1

I~ L ~n L ---packed ~ op~:, ~~-" o~en,2 Detection window

Fig. 1.1. Schematic illustration of a CEC column consisting of a packed and an open segment. The latter is divided into pre-detection and post-detection open segments by the detector window.

Migration and Electroosmotic Flow

3

packed and an open segment of lengths Lpackedand Lopen, respectively. This complicates the exact calculation of the pertinent CEC parameters in the packed and the open segments, the column design and the interpretation of electrochromatographic data [18,20,21,24,26,27]. In view of the great present interest in CEC, we hope that this chapter reveals the complexity of the electrokinetic phenomena underlying EOF in porous media and will stimulate the experimental and theoretical work necessary for a better understanding of the physicochemical basis of CEC and to develop this technique into a powerful analytical tool. The goal of this chapter is to review previous work, to reveal the complexity of the electrokinetic phenomena underlying electroosmotic flow (EOF) in porous media and to compare the case of an open capillary that is used in capillary zone electrophoresis (CZE). It follows from the conservation of the volumetric flow rate that in most cases a "flow-equalizing intersegmental pressure", which is different from the pressures at the two ends of the column, develops at the interface of the packed and open segments and has significant effect on the magnitude as well as the radial distribution of the flow velocity in the open segment. A framework is presented that uses measurements of current and EOF performed on CEC columns for the evaluation of parameters such as the conductivity ratio and the interstitial EOF mobility that are useful tools for characterizing them. The actual EOF mobility, that is obtained after taking into account the porosity and tortuosity of the packing, is a better measure of surface properties of the packing. Further, a modified form of Overbeek's model that was originally developed for porous media of infinite dimensions has been used to account for the wall effect in the packed capillary columns used in CEC. Finally, migration of charged components in a CEC column is simulated and expressions are presented for estimating the retention time if the chromatographic and electrophoretic properties of the sample components are known. It is shown that, by varying the length of the packed segment and maintaining same the total length of the CEC column and the position of the detection window, the balance of the chromatographic and electrophoretic forces can be shifted and the selectivity can be adjusted if the separation involves the interplay of both mechanisms. 1.2 FLOW OF IONS IN OPEN TUBES

When an open tube with fixed charges at the tube wall is filled with an electrolyte solution, the ionic atmosphere forms an electrical double layer [31-33]. Since the double layer has a higher concentration of counterions than the bulk solution, electroneutrality requires that the bulk electrolyte outside the double layer has the same amount of excessive coions. Flow of ions in an open capillary tube occurs via one of the three possible modes. References pp. 37-38

4

Chapter 1

First is ionic conduction through the bulk electrolyte, where ions migrate under the influence of the electric field by virtue of their electrophoretic (ionic) mobilities. Second is convection of the bulk electrolyte, where the mobile phase is voltage and/or pressure driven and carries the excess charge in the bulk with it [34-36]. This contribution to the total conductivity is found to be negligibly small at ionic concentrations higher than 1 mM [34,35]. Third is ionic conduction through the double layer at the tube wall due to migration of the excess counterions under the influence of the electric field [34,36-38]. The contribution of surface conduction to the total conductivity is significant only at ionic concentrations smaller than 10 mM and zeta potentials smaller than 150 mV [34,36-38]. When ionic conduction through the bulk electrolyte is the dominant mechanism of ionic migration, the conductivity of an electrolyte filled cylindrical capillary, CYopen,is expressed as [18]

i Lopen (~open : Vopen Aope n

(1.1)

where, i is the current flowing through a capillary of length, Lopen, and cross-sectional area, Aopen, when a potential drop, Vopen, is applied across it. The conductivity of an electrolyte solution is an intensive property that is independent of the length or diameter of the capillary tube and can be expressed in terms of the concentration charge and mobility of the constituent ions as follows [ 18,39]

Oopen = [ 72 ~

Z2 Vj Cj

(1.2)

J

where, F is the Faraday constant and zj, vj and cj are the valency, mobility and molar concentration of the j th ionic species, respectively. Equation 1.2 can be rewritten as

~open= F2 C Z z2 Vj Xj

(1.3)

J

where, C is the molar concentration of the buffer and xj = cj/C is the number of moles of jth ionic species per mole of buffer. For weak electrolytes, xj depends on the dissociation of the ionogenic species and thus on the pH. It follows from Eq. 1.3 that

Migration and Electroosmotic Flow

5

the conductivity of sufficiently dilute electrolytes increases linearly with the molar concentration of the electrolyte and the slope depending on the charge, size and concentration of the ionic species. 1.3 ELECTROOSMOTIC FLOW IN OPEN TUBES

In open tubes with thin double layers and when there is no polarization, the EOF mobility, kteo, open,

can

be expressed by the following relationship [40-44]

~: eo ~.~ ~eo,open

-"

(1.4)

"

where, e is the dielectric constant of the medium, eo is the permittivity of the vacuum, and rl is the viscosity of the bulk solution. ~w is the zeta potential of the wall and is defined as potential on a hypothetical 'surface of shear' close to the tube wall that can be assumed to exist so that while the bulk electrolyte beyond this surface is moving, there is no motion between this surface and the charged wall. The mobility, as defined by Eq. 1.4, depends on the properties of the charged surface and the bulk electrolyte but is independent of the capillary diameter or the applied voltage. When the surface

(z e qJo~

charge is independent of the electrolyte concentration and exp~ kB T ) >>1 for the simpler case of z:z electrolytes [33]

Wo= (ksT~ logC + constant

(1.5)

~,ze)

where, ~o is the surface potential, kB is the Boltzmann constant, T is the absolute temperature and e is the elementary charge. Since the zeta potential is of the same order as the surface potential for the present case, the above equation can be rewritten to relate the EOF mobility to the buffer concentration as follows

~eo,open

=

eeo~_(e_eoksT)logC+constantq Tlze

Referencespp. 37-38

(1.6)

6

Chapter 1

According to Eq. 1.6, the dependence of the EOF mobility on the logarithmic electrolyte concentration is linear. In practice, the EOF mobility in the bulk electrolyte is estimated from migration data obtained with a suitable neutral and inert tracer in an open tube by using the following expression

LdL

(1.7)

~[eo,open- to,open V

where, Ld is the distance between the inlet and the point of detection of the capillary and to, open is the migration time of the tracer in an open tube of length, L, with an applied voltage, V. 1.4 FLOW OF IONS IN PACKED COLUMNS 1.4.1 Conservation of electric current Besides the three modes of ionic migration discussed above for an electrolyte filled open silica capillary, current in a packed column can also flow via conduction through the stationary phase. However, when ionic conduction through the bulk electrolyte solution is still the primary form of ionic migration, conductivity of a packed column, ~packed, can be expressed along similar lines as Eq. 1.1 in the following manner [18]

i' Lpacked

(1.8)

(~packed = VpackedAopen

where, i' is the current flowing through the intraparticulate and interstitial mobile phase spaces of a CEC column fully packed with a stationary phase, length of the column and Vpackedis the applied voltage.

Lpacked is the

1.4.2 Evaluation of conductivity of the packed and open segments Columns in CEC often have a packed and an open segment with a detection window in between. The conductivities of the open and packed segments,

(Yopenand

Crpacked, are then evaluated from the currents measured with the capillary tubing in the absence, iopen, and presence, ipacked, of the packing by using Eqs. 1.1 and 1.8 as

Migration and Electroosmotic Flow

7

follows

(1.9)

L iopen (Yopen= V A open

and (1.10)

L iopen ipackedLpacked (Yopen-- V [iopenL - ipackedLopen]Aopen

where, Lopen and Lpacked a r e the respective lengths of the open and the packed segments of the CEC column. 1.4.3 Evaluation of the potential drop across the packed and open segments

Once CYopenand Cypackeda r e known, the potential drops across the two segments of either column with packed length, Lpacked, are evaluated by using the following expressions [ 18,20]

I

I

(1.11)

(YopenLpacked+ (YpackedLopen

Vopen--V_Vpacked:V~~--VI

(YpackedL~ I (YopenLpackedd- (YpackedLopen

(1.12)

where, Ropen, Rpacked and R are resistances of the open segment, the packed segment and the total column, respectively. Equations 1.11 and 1.12 express that the total potential drop, V, is distributed across the packed and open segments of the column according to the relative magnitude of their resistances. Figure 1.2 shows that with increasing dimensionless packed length, ~, = Lpacked/L, the total resistance of the column and hence the potential drops across the two segments vary nonlinearly according to Eqs. 1.11 and 1.12. As ~, goes from 0 to 1, the two potential drops,

Vpackedand Vopen, monotonicaly increase and decrease from zero

to V and V to zero, respectively. Further, a higher conductivity ratio, CYopen/Cypacked, means a higher resistivity of the packed segment. Thus, a higher potential drop across References pp. 37-38

8

Chapter 1

20

.

.

.

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Packed segment

>~e

i,i o p e n segmen t . . . . . . ~

15

cs

2 ~ e-, o o

(Yopen /(Ypacked

\, /1.5 _ \X/3.1 10 **~

1.5

-.

5

"

~

% ~

9"'~.. -..

10

~ 9.,..

~. 9-. ......

.

0

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0.25

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0.5

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0.75

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~

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0

0.25

0.5

0.75

1

Dimensionless packed length, ;~ Fig. 1.2. Plots of the potential drop across the packed and the open segment of a CEC column against the dimensionless packed length, ~,, with conductivity ratio, CYopen/CYpacked,as the parameter. The respective currents with columns having packed segments of 0, 10 and 20 cm were reported as 6.6, 3.9 and 2.6 pA, respectively. The conditions were: fused silica capillary, 50 p,m x 30 cm; applied voltage, 18 kV; column packing, 3.5 mm Zorbax ODS, 80 A; mobile phase, 50 % (v/v) ACN in 10 mM sodium borate, pH 8.0.

the packed segment is required to generate current equal to that in the open segment causing an increase in the nonlinearity of V vs ~ plots in Figure 1.2. Since the total potential drop is kept constant, the plots of Vpacked and Vopen against ~, are mirror images of each other. The data plotted in Figure 1.2 was obtained by measuring the current in columns differing only in the length of packed segment, Lpacked, under otherwise identical condition [ 19]. The conductivity ratio, CYopen/CYpacked,was calculated to be 3.1 and the potential drops across the two segments, Vpacked and Vopen, were evaluated using Eqs. 1.11 and 1.12. 1.4.4 Evaluation of the equivalent length of the packed segment

For the sake of simplicity, we may replace the packed segment by a hypothetical open tube of length, Le, that is the same as the distance traveled by the neutral and inert tracer in the packed segment. The lumen of this hypothetical tube, Apacked, is assumed to be the same as the free cross-sectional area of the packed column so that 2 Apacked = gae , where, ae is the radius of the hypothetical tube. The equivalent length,

Le, is determined from the ratio of the conductivities of the packed and the open

Migration and Electroosmotic Flow

9

O'open/~packed -o

4

0 O.

3

,.J ............... .

.

3.]

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

.

.

I

0

.

1.5

,

,

,

,

0.25

I

0.5

,

~

,

,

I

0.75

1

Dimensionless packed length, Fig. 1.3. Plots of the ratio of the equivalent and the actual lengths of the packed segment, Le/Lpacked, against ~ with conductivity ratio as the parameter. Conditions same as in Figure 1.2.

segments of the column, as [20,24,43]

Le L 41 +LPLked ( (Yopen ) _( LopenI Lpacked-- Lpacked ~(~packed-- 1 ~Lpacked)

(1.13)

Altematively, the equivalent length can be evaluated directly from the current measurements in the absence and presence of the packing, topen and Zpacked, respectively.

_ q ioen I Lpacked Lpacked i;acked ~LpackedJ According to Eq. 1.13, Le depends on the length of the packed segment, Lpacked, as well as the conductivities of both the mobile and stationary phases. The ratio of lengths, Le/Lpacked, as a function of ~ is plotted in Figure 1.3 with the conductivity

Referencespp. 37-38

10

Chapter 1

ratio, t~open/CYpacked,a s the parameter. A higher t~open/t~packedratio signifies an increase in the resistivity of the packed segment which is equivalent to an increase in the Le/Lpacked. Further, if the resistivities of the two segments are close, as for C~open/t~packed= 1.5 or 3.1, Le/Lpacked is nearly a constant as assumed in the following discussion. 1.4.5 Evaluation of the electric field strengths in the two segments

The actual electric field (under which the migrants travel) in the packed segment, V p a c k e d , and the equiva-

Epacked, can be evaluated from the pertinent potential drop, lent length, Le, as

Epacked =

Vp,~a Le

(1.15)

whereas, the electric field in the open segment is obtained by the relationship

Vopen

(1.16)

gopen = Zopen

Evidently, both Epacked and Eopen a r e quite different from the "fictitious electric field", E*, that is obtained by dividing the total potential drop across the column, V, by the total column length, L.

E* =__V L

(1.17)

Clear distinction between Epacked, Eopen and E is of particular importance when accurate knowledge of the electric field strength is required for evaluation of the electroosmotic and electrophoretic mobilities from the observed migration times. Similar situation arises in other systems that are subject to similar conservation principles, e.g., displacement electrophoresis. Figure 1.4 illustrates plots of the above three electric field strengths against the dimensionless packed length ~, according to Eqs. 1.15, 1.16 and 1.17 with

CYopen/t~packed=

3.1. It is seen that, the field strength is higher in the packed than in

Migration and Electroosmotic Flow

11

TE*

I Eopoo

LU

-J~

r

100

|

K,

80

"'~,

oLU

!

|

,E packed

~'-~

L

~E

! Eop en

E packed /X

.

"-,~,

60

E

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

~ .............................................................

40

.m

dl=a

o

20

u.I

0

,

,

|

O

I

,

,

,

,

I

,

,

,

,

I

,

.25 0.5 0.75 Dimensionless packed length,;~

,

,

,

1

Fig. 1.4. Plots of the actual, Epackedand Eopen, and the fictitious, E*, electric field strengths in the packed and the open segments against )~ for Case I with (yopen/CYpacked = 3.1. At the top, the discontinuity in the electric field strength at the interface of the two column segments is illustrated. Rest of the conditions as in Figure 1.2. the open segment, i.e., Epacked > Eopen. Both field strengths, Epacked and Eopen, increase with decreasing )~ and concomitantly the mobile phase velocity increases in both segments. Figure 1.4 also illustrates the inequality of the actual electric fields in the two segments that serves as a caveat that instead of the fictitious electric field strength, E*, the actual electric field strengths, Epacked and Eopen, should be evaluated and reported together with other chromatographic conditions in CEC. 1.4.6 Conductivity ratio in CEC The ratio of the conductivities of the packed and the open segments of a CEC column, % is given by the following relationship

Clp~kea

(1.18)

(Y open

The reciprocal of q~ is often called "formation factor" in reservoir engineering and

Referencespp. 37-38

12

Chapter 1

geology [45-47]. It is independent of mobile phase properties except when the ionization of the liquid is affected by the stationary phase and therefore, offers a simple means to characterize properties of the stationary phase. The conductivity ratio has been used to estimate the porosity of rock samples [45,47] and is related to the electrokinetic porosity, eT, by Archie's law as follows

(Ypacked q)----- ~ - -

(1.19)

m ET

(Y open

where, m is an empirical constant. The electrokinetic porosity in Eq. 1.19 depends on the morphology of the packing particles and is not necessarily the same as the porosity employed in chromatography. When the porosity of the media is greater than 0.2, as is the case generally, m = 1.5 provides a very close approximation to the experimental data and so this value will be used here [46,47,49,50]. 1.5 ELECTROOSMOTIC FLOW THROUGH PACKED COLUMNS 1 . 5 . 1 0 v e r b e e k ' s model for EOF in porous media

The following analysis is based on Overbeek's work [43,52,53] and is valid for porous/nonporous packing particles of any arbitrary shape as depicted in Figure 1.5. The assumptions are that the particles be non-conducting, have uniform zeta potential and a double layer thin compared to the radius of the pores in the plug. Overbeek, upon integration over the whole interstitial volume of the bed, obtained the following expression for the average velocity

(1.20) " ~ U P > - - W c cl I v f fc p

dVc=- ~r~l ~

d~

where, ~ is the zeta potential at the surface, Vc and Vcfare the total column Vvolume and the volume of the interstitial space, respectively. The integration is performed over Vcf only since flow is only in the interstices. Up is electroosmotic velocity that is generated locally at the packing surface and is given by an expression similar to Smoluchowski's equation for the EOF, as follows

Migration a n d Electroosmotic Flow

13

Lpacked

/

/

Packing

Flow oath of length, Le and tortuosity, "c = L e/Lpacked

Fig. 1.5. Schematic illustration of the tortuous flow path in the packed segment of a CEC column.

Up

~

~eo~E

(1.21)

lq

For the current, i, the following relationship holds

i = OpackedE = ~(YopenIV? d

(1.22)

where, ~packedand Oopen are the conductivities of a completely packed column and an open tube, both filled with the electrolyte solution, respectively. Combination of Eqs. 1.20 and 1.22 yields the average velocity as

> = e eo ~ E (Opacked~

1]

(1.23)

~ (Yopen) !

with the conductivity ratio,

Opacked/Oopen,that is readily determinable experimentally.

Equation 1.23 was derived by Overbeek and Wijga [52] and Overbeek [53] to describe EOF in a porous medium without boundaries and will be used in the next section to express the effect of the capillary wall on the flow distribution inside the column.

References pp. 3 7-38

14

Chapter 1

1.5.2 Effect of charged capillary wall It is assumed that the EOF is generated only at the charged wall and the packing particles are uncharged. The flow can then be visualized in the form of very thin annuli of liquid in the packed column. Each annulus faces a force in the forward direction (the direction of EOF) from the annulus enveloping it and a force in the backward direction from the annulus inside it. The inertia terms and the compressibility of the fluid are assumed to be negligibly small. The net viscous force, Fv, on such an annulus of unit volume in absence of any particles is given by

Fv = Ar -d-rrd(r d u~rW)d)

(1.24)

where, r is the radial coordinate and Urw is the local velocity in the axial direction. In such a column with charged tube wall and uncharged packing, the flow velocity would be a rapidly fluctuating function of radial position with zero value at the surface of the uncharged particle and maximum in the intraparticular space. Hence, the velocity under consideration, Urw, is more like a volume average velocity for the volume element. Besides the viscous forces, the fluid in the shell also experiences a drag force from the packing particles in the shell. The total drag force, Fd, offered by spherical packing particles of diameter dp that are located far enough from each other to act like isolated spheres, is given by the product of the drag force by an isolated sphere and the number of spherical particles in a shell of unit volume as follows

1-ST) Fa = (6 n 1"1dp Urw) 4 ~ d3p/3

(1.25)

where, ~;T is the total porosity of the column. Since in packed columns the particles are in close contact with each other, the actual drag force is different than that given by Eq. 1.25. This is corrected for by introducing the dimensionless packing parameter, c~, which depends on the structure of packing and shape of the particles and should be easily determinable from experimental data. Performing the balance between the viscous and the drag forces and simplifying the equations we have that

Migration and Electroosmotic Flow 1 d (r du~w~=9 c~(1-~:r) u,.~ 132

5

4

15 (1.26)

where 13 is another dimensionless constant that is readily evaluated if et and ~;T are known from the following relationship

./

(1.27)

13=3-N et (1 - ~;'' 2

The boundary conditions for solving the differential equation as given in Eq. 1.26 are

d u~ _ 0

at r = 0

(1.28)

u~=uw

atr-a

(1.29)

dr

and

where, Uw is the EOF velocity at the wall and a is the tube diameter. Equation 1.29 simply reflects no slip condition at the plane of shear that is very close to the tube wall for thin double layers. Solution of the system of Eqs. 1.26, 1.28 and 1.29 for the potential distribution can be found in the literature [39,42] and written for the local velocity as

io (~ ~/r ] u :uw Io(

(1.30)

a/r

where, Io is the Bessel's function of the zeroth order. For ~r/dp greater than 3.5, which is generally the case, the zeroth order Bessel's function can be approximated by

Referencespp. 37-38

16

Chapter 1

!

l

Packing particle

Uw

Electrosmotic

velocity

Fig. 1.6. Schematic illustration of loci of the wall effect where EOF decays in the case of charged tube wall and uncharged packing particles.

e ~r/dp Io (f3 r/dp)= 42 rt [3r/dp

(1.31

)

which in turn can be substituted back into Eq. 1.30 to give the simplified solution as [11]

I e ~'/4 {2x[3a/dp I u~ = u~ ~12 rt [3r/dp e f3a/dp

= Uw

I~f~)e~("-')/4

(1.32)

Equation 1.32 for the region close to the wall is illustrated in Fig. 1.6. The flow velocity is maximal at the plane of shear and then decays quickly as we move away from the wall. This should not be surprising as in a column packed with uncharged particles, EOF is generated at the tube wall and the interior of the tube contributes only the drag resistance. It should be noted that the flow velocity falls significantly as soon as we move a distance of one particle diameter away from the column wall. Previous experimental studies on packed beds that have been published in the literature support these findings [55].

Migration and Electroosmotic Flow

17

The average velocity in the column for the above case can be easily determined using the velocity profile given in Eq. 1.32 and accounting for the tortuosity in the bed as

- -

~o27~rurwd a2 ~,((Ypackedl=cYopen Uwl(YopenCYpacked II.a21o(_~a/dp))2 ~orlo(~r/dp)dr(1.33)

that can be simlified for the case when [~r/dp is greater than 3.5 as follows [ 11]

I

< blzw> = blw O a'e ll (~open

a2

l

(1.34)

Equations 1.34 suggests that the average velocity in the packed column varies linearly with the dimensionless particle diameter, dp/a. Under conditions employed in CEC, the EOF is generated not only at the capillary wall but also at the surface of the packing. When the zeta potential of the wall is same as the packing (~w = ~p), the velocity profile should be flat according to Overbeek's expression in Eq. 1.23. For the case when the zeta potentials at the tube wall and the particle surface are not equal, the total velocity could be evaluated by adding a term to Overbeek's velocity expression to account for the wall effect. This term would be given for the local and average velocities with the zeta potential of the wall being replaced by the 'excess zeta potential' on the wall (~w - ~p) that is responsible for the wall effect, i.e.,

(1.3:5)

where, Ur is the net local velocity from both contributions. Figure 1.7 illustrates the effect of the magnitude of the excess zeta potential on the radial profile of the EOF velocity. The plots show that the wall effect is limited to a narrow annulus at the wall that increases in width with the magnitude of the excess zeta potential. The effect of particle diameter on the radial flow profile is illustrated in Fig. 1.8 for certain typical cases. It is seen that the wall effect increases with the particle

References pp. 37-38

18

Chapter1 '

'

'

i

,

,

,

i

,

,

,

,

,

,

i

[-?

2.5

r

,

,

'

4

..

[ ~ P ~ = L_~c~ = 2 ~ P ~ ' ~ , . j ~

faster at the wall

=mm

o (D > i3. r ~ A L_

i

m

1.5

"<_,/1 ,,,~.,~ t

~o~ = ~ p

N o wall effect

-i,~..

-.

"~.~.

L

o

=l

0}

0.5

a

co =-~

l o w e r at the wall

E

=l

-0.5

P f

-1

,

0

,

J

I

,

a

0.2

,

I

,

~

0.4

n

I

0.6

a

n

,

I

n

0.8

~

,

1

Dimensionless Radial Position r/a Fig. 1.7. Graphs illustrating the hypothetical radial flow distribution when both the tube wall and the packing are charged. The ratio of zeta potentials of the wall and the particles in the packing is the parameter. The velocity is made dimensionless by using the local electrosmotic velocity at the particle surface as the reference. Conditions; R = 50 ~tm; ~ = 0.4 and

(~w/~p) dp- 5 ~tm.

diameter of the packing for fixed tube diameter. This is because with increasing particle diameter, the area of the one particle diameter thick annulus relative to the total cross-sectional area increases and so the effect becomes more evident. By integrating Eq. 1.35 over the bed volume we obtain the expression for the average velocity in the column as

< blr ~> :

< blrw >

I 1-~~+=I1+l-~ll-~ll~-111

(1.36)

As mentioned above, the wall effect would be palpable only in the case when the two zeta potentials are unequal and then increase with the magnitude of the difference. As the particle diameter increases, the wall effect increases too. 1.5.3 Conservation of volumetric flow rate

The packed and open segments of the CEC column being connected in series, the

Migration and Electroosmotic Flow

'

.

i

'

i

'

'

!

'

'

'

~

d = lO~rn

.-~

1.8

O

l

!

19

dp = 5 lam \ r ] ~

o

>

~=~

1.6

d =3 ~tm

----Iw~a]---;ee"Sing

t! /

Im

e-

~.

.2 ~ c

1.4

E

1.2

o

1 0

0.2

0.4

0.6

Dimensionless

0.8

1

Radial Position r/a

Fig. 1.8. Graphs illustrating the hypothetical radial flow distribution when both the tube wall and the packing are charged with the particle diameter as the parameter. The velocity is made dimensionless by using the local electrosmotic velocity at the particle surface as the reference. Conditions; a = 50 ).tm; ~p = 0.4 and ~w = 2 ~ .

mass conservation law requires that the volumetric flow rate of the mobile phase has to be the same in the two segments. The actual flow velocities in the open and packed segments, denoted by Ueo, a packed and Ueo, a open and taken as the average velocities over the total free cross-sectional area, can be related as follows

a

a

bleo, packed Apacked = Ueo, open A open

(1.37)

However, if the two segments were operated individually under conditions prevalent in the CEC column, the "virtual" flow velocities would be given by the following expressions

(1.38)

and

References pp. 37-38

Chapter 1

20

t Vope~t

(1.39)

l'leo, open : ~leo, open Zopen

where kteo,packed and ~teo,open a r e the respective mobilities of the EOF marker measured independently in the packed and open segments of the column. Even though the flow velocities as given by Eqs. 1.38 and 1.39 are "virtual", they are useful for the estimation of the EOF generating ability of the packed and the open segments. Thus, they can be used to characterize the two segments and distinguish among the three possible cases in CEC according to the relative magnitude of the volumetric flow rates, Ueo,packedApacked and Ueo,openAopen. Case I: Ueo,packedApacked < Ueo,openAopen. This is the case when zeta potential at the packing surface is such that the volumetric flow rate generated in the packed segment is lower than that in the open segment due to the tortuosity. Case II: Ueo,packedApacked = Ueo,openAopen. Such equality may occur in practice when the conductivites and relative lengths of the packed and the open segments are such that the velocities in the two segments as given by Eqs. 1.38 and 1.39 fulfill the conservation of volumetric flow rate as given by Eq. 1.37. Case III: Ueo,packedApacked > Ueo,openAopen. The flow rate generated by the packed segment in this case is higher than that by the open segment.

1.5.4 Flow equalizing intersegmental pressure As discussed above, if each segment was operated individually it would generate the virtual flow velocity as given by Eqs. 38 and 39. However, conservation of volumetric flow rate in the form of Eq. 1.37 puts a restriction on the flow velocities. A close analysis of the various variables in the system and of the different possibilities that may occur leads to the conclusion that pressure at the interface of the packed and open segments, Pi, has to be different from that at the inlet and outlet of the CEC, P0. This "flow equalizing intersegmental pressure", Pi, provides a mechanism to alter the O

t2

velocities, Ueo,packed and Ueo,open, to Ueo,packed and Ueo,open, in order to satisfy the conservation of volumetric flow rate as given by Eq. 1.37. Development of similar local pressure gradients inside the packed bed as a result of abrupt velocity changes in the vicinity of packed particles were mentioned in recent publications [23,24,28]. Figure 1.9 illustrates pressure distribution in a typical CEC column when

Ueo,packedApacked < Ueo,openAopen (Case I) and the intersegmental pressure, Pi, is smaller than the ambient pressure, PO. In the open segment this pressure gradient builds up opposite to the direction of the EOF, slowing the flow down and making the volumetric flow rate equal to that in the packed segment. There is a continuos dissipation of energy in the system in the form of work done by this induced pressure

Migration and Electroosmotic Flow

P

21

o-

P.

I I

>p Po open

.i, P o

/g

t

i packed

I I I [ I I

=3.1

r PACKED

f" P

4~ OPEN

P

o

i

0.5 r

/~packed

o .._.....1.o

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

-0.5

-1 '~

~/

1.5.t" j,,,a"

~= -1.5

..

-2 0

0.25

0.5

0.75

1

Dimensionless packed length, Fig. 1.9. Plot o f the pressure difference, Pi-Po, across the packed segment against ~, with the conductivity ratio as the parameter. At the top is a schematic illustration o f the pressure gradients across the open and the packed segments of the column when ~open/CYpacked = 3.1. Conditions used are Al~acked/Aopen= 0.4; ~;p = 0.4; 11 = 10-3 N m -2 S -1, laeo,packed = ~teo,open =3.33 x 10 -8 m 2 V 1 s " . Rest of the conditions same as in Figure 1.2.

gradient in equalizing the flow rates in the two segments. The electrical energy being the sole input in the system, part of it is dissipated in this manner. The conservation of volumetric flow rate for an incompressible fluid under conditions of steady state can be expressed [20,39,42,54] as follows.

Referencespp. 37-38

Chapter 1

22

B~176176 kLpackedJ

[Vpacked'][ 2I,(Kae) ] = Le ) l--Kaelo(Kae)J (1.40)

--

Lope. ~ Aopen q- ~eo,operg~open Lopen

Kalo(~ca)

where, tc is the Debye screening parameter taken as the reciprocal of the double layer thickness, and B ~ is the specific Darcy's law permeability. The Kozeny-Carman equation is often used to evaluate B ~ for packed beds and is given by

B~ = d2p

3p

(1.41)

180 (1 - ~:p) 2

where dp is the particle diameter and Sp is the interstitial porosity of the packed bed. For the case of thin double layers (Ka and ~cae > 50), Eq. 1.40 can be solved to obtain an expression for the intersegmental pressure Pi as [20]

pi = p 0 _ [ ~eo, openAopen Vopen-- ~.leo,packedApackedVpacked(Lopen,/Le)] (B ~ Lope. mopen/rl Lpacke3 + (a 2 Aopen/grl)

(1.42)

The final expression for the flow equalizing intersegmental pressure Pi as given by Eq. 1.42 can be rewritten in terms of the dimensionless packed length, )~, as follows

Pi : Po - [ ~te~~ A~ Vopen -- ~teo,packedApacked Vpackea(1- X)L/Le)l [B~ (1 - )~) AopeJr 1 ;~] + (a 2 Aope./8rl)

(1.43)

Equation 1.43 shows that the magnitude of the intersegmental pressure depends on the applied voltage and the lengths, free cross-sectional areas and resistivities of the two column segments. The pressure difference, P i - PO, as calculated from Eq. 1.43 is plotted as a function of the dimensionless packed length, ~, with the conductivity ratio as the parameter in Figure 1.9. The pressure drop across any of the segments, P i - PO, is maximum at small ;~ values and becomes zero at the extremity )~ = 1. In the present

Migration and Electroosmotic Flow case

23

Ueo,packedApacked< Ueo,openAopen, and

negative. Further, it is seen that

as

so the pressure drop,

Pi- PO, is

always

CYopen/CYpacked increases, the pressure drop de-

creases. This is so because smaller conductivity of the packed segment leads to a higher potential drop across the packed segment and, consequently, a smaller inequality in the flow rates generated in the two segments. On the basis of above discussion, the role of the flow equalizing intersegmental pressure in Cases I to III can be described as follows. Case I: In this c a s e , Ueo,packedApacked< Ueo,openAopen and so as discussed above, Pi < PO in order to equalize the flow rates in the open and packed segments. Case II: Here, Ueo,packedApacked = Ueo,openAopen and so it follows from Eq. 1.43 that

Pi=Po. Case III: This case is opposite of Case I in that Ueo,packedApacked > Ueo,openAopen and consequently it follows from Eq. 1.43 that Pi > PO. 1.5.5 Flow velocities in the packed and open segments

The flow velocities through the packed and open segments in series can be expressed by contributions from the EOF and the intersegmental pressure driven flow so that for the case when the double layer thickness is smaller than 2% of the mean channel diameter, i.e., ~:a and ~:ae > 50, it follows that [20]

a lPO-PilfB~176 bleo,packed= Le ~rl ApackedJ

packedIV~kedl

(1.44)

and

Hea~176

(P~ tV~ Lopen --~ + ~[eo,open Lopen

(1.45)

When the packed segment is much less permeable than the open segment, the flow velocity in the packed segment is not significantly influenced by the pressure gradient. Then the actual velocity in the packed segment turns out to be identical to the virtual velocity as given by Eq. 1.38, and we obtain that

a

(Voc,eq Le )

ldeo,packed= ldeo,packed= ~eo,packed

Referencespp. 37-38

(1.46)

24

Chapter 1

Ueo, packed

I Ueo, open 7 U eo, packed

l,,.ml

o

3

E E

2

o}

(n

"J

7 U eo, open

Packed segment

-....

~""~". ,~.....,. ,..,~..

o 0

,,..

1

Open ..................... " ' " - , , - ~ g m e n t P" ......~... eqn. (39)

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

o} >

.~.

,,..

=~,, ~.. , , ~ ~ . . , = = = _ _

~ = = ~ _ ,===~_ ====~_==,=_._ =,==_ _ ==.

eqn. (45) 0

'

'

'

'0.

s

. . . .

0.

I5

'

'

'

'0.

5

'

,

,

,

1

Fig. 1.10. Dependence of the flow velocity in the interstices of the packed segment and the velocities in the open segment on ~ for Case I and CYopen/C~packed = 3.1. The inequality of the two velocities in the open segment, Ueo,open and uaeo,open, illustrates the effect of the pressure differential, Pi-Po. The schematics of the CEC column at the top illustrates the discontinuity in flow velocity at the interface of the two column segments. Conditions as in Figure 1.9.

Since in columns most commonly used in CEC at present, the detection window is located right after the packing, the chromatograms will not be palpably affected by the pressure gradient. Therefore, once the potential drop across the packed segment is evaluated, Eq. 1.46 gives a satisfactory estimation of the actual interstitial flow velocity through the packed segment of the column. Figure 1.10 illustrates the dependence of the mean actual mobile phase velocities through the interstices of the packed segment, Ueo, a packed, and in the open segment, O

Ueo, open, when CYopen/CYpacked=

3.1.

In order to illustrate the effect of the intersegmen-

tal pressure on the flow velocity in the open segment, the open segment velocity was plotted against ;~. The magnitude of this effect is greater at low ;~ and appreciable also in the range 0.6 < ;~ < 0.9, which is likely to be of practical importance. As shown by Figure 1.10, the velocities decrease about three fold in both the packed and the open segments when X increases from 0 to 1. The flow equalizing intersegmental pressure also has an effect on the flow profiles in the open segment. The flow field in the open segment may be affected strongly by

25

Migration and Electroosmotic Flow

Case l

e=

1 I~ /

Case II

0.25

To.8

/ t l!

oE

.

2i . . . . . . . . . 0.~5--~. . . . . . . .

Z

[ t

.~J.,050

Case III

.

.

.

.

.

6 [_

0.25_

t

/

/

",,.

o.so ~. /J

/

\~

1.1

t/i

.

9

,

,

0.9

9. . . . . . . . . . . . . . . . . . .

-1

0

1

0.7

-1

0

1

OL~ -1

0

1

Radial position, rla Fig. 1.11. Fully developed flow profiles in the open segment of the CEC column for the three cases with Z as the parameter. The dimensionless radial coordinate is given by fla. Conditions as in Figure 1.9.

the axial pressure gradients, whereas in the packed segment the flow field is expected to remain unaltered. This is illustrated in Figure 1.11 where the flow velocity in the open segment, Ueo,open(r), is plotted against the dimensionless radial coordinate, r/a, with ~, as a parameter for the three cases defined earlier. The results presented in this section strongly support other findings [29] and explain the flow profiles that were obtained via flow visualization in the CEC column right after the intersegmental frit. C a s e I: The conservation of volumetric flow rate yields the following expression for

the radial distribution of the flow velocity in the open segment when ~:a > 50.

2

Ueao,open(r) : --

Zopen

~

1 --

(1.47) + ~eo, open Zopen

Equation 1.47 shows that in this case the velocity profile in the open segment is also the sum of two contributions: one from the EOF and other from the intersegmental pressure. As a result, as shown in Figure 1.11, the flat velocity profile of the References pp. 37-38

26

Chapter 1

electroosmotically driven flow is distorted by the presence of the pressure gradient. The resulting flow profile is reversed parabolic with the lowest velocity in the center line ( r / a - 0). Further, it is seen in Figure 1.11 that with decreasing X, the flow velocity increases and the flow profile becomes more parabolic. Case II: Since Pi = PO, Eq. 1.44 simplifies to

(1.48)

a

Ueo,ope,(r) : ~teo,ope, Lope,

Equation 1.48 predicts a perfectly flat flow profile as shown in Figure 1.11 since the pressure effects are absent. As X increases, the electric field strength across the open segment decreases resulting in a reduction of the flow velocity. Case III: For the special case when the inner wall of the open segment is neutral and no EOF is generated there, Eq. 1.44 becomes

2

blea~176

Lopen - ~

(1.49)

1-

which is the equation of parabolic flow profile obtained with pressure driven laminar flow as in the open segment only viscous flow prevails. As shown in Figure 1.11, the pressure gradient increases with decreasing X and as a result, the flow velocity increases in the open segment. 1.5.6 EOF in packed columns

EOF mobility in the interstices of a packed column, Ueo,packed, c a n be expressed in a way similar to Eq. 1.4 [40,43]

bteo, p a c k e d - -

~o G q

(1.50)

where, ~ is the zeta potential at the surface of the packing. Equation 1.50 holds also for EOF through the pores of the packing particles when the pore diameter is sufficiently large (typically, greater than 300 A for electrolyte concentrations greater than

Migration and Electroosmotic Flow

27

10 mM of a 1:1 electrolyte) compared to the thickness of the double layer. The mobility of an inert, neutral EOF marker through the packed segment of a CEC column has been expressed in the literature [53] by the following relationship

LpackedL

(1.51)

~eo, packed : to,packed V

where, to,packed is the migration time of the inert tracer in the packed segment and

~teo, packed is the "apparent" EOF mobility that is evaluated for the packed segment by using the voltage applied to the whole column. Since ~teo,packed depends both on the zeta potential and the porosity of the packing, a higher ~eo, packed c a n be a result of a higher charge at the packing surface as well as a higher porosity of the packing structure. Further, it follows from Eq. 1.51 that the apparent EOF mobility would also depend on the column architecture (i.e., Lpacked/L) and so will be different for two columns having different Lpacked/L under identical conditions of stationary and mobile phases. In view of the above discussion it follows that the actual interstitial mobility of an inert, neutral EOF marker through the packed segment that would be useful for determining the zeta potential of the packing should be calculated as [ 18]

L~

2 Lpacked 2

D

( 1.52)

~eo, packed: to, packed Vpacked- to, packed Vpacked

where "c = Le/Lpacked is the tortuosity of the packing. It follows from a comparison of Eqs. 1.51 and 1.52 that in the calculation of the actual EOF mobility, the effect of tortuosity is taken into account so that ~teo,packed can be used to calculate the zeta potential of the packing. In the literature of electrokinetic phenomena in porous media [44], constriction factor is employed to describe the effect arising from the variability of cross-section along a given flow path. However, in packed CEC columns this factor is estimated to be in the range 0.93-1.0 and so may be neglected in the following treatment.

1.5.7 Migration times, velocities and M-factors The total migration time, tr, of a sample component across the column is the sum

References pp. 37-38

28

Chapter 1

of the residence times of the sample component in the packed and the open segments of the column and is given by

tr--"

Le

+

1,lpacked

(1 53)

Lope., t Uopen

Upacked and Uopen, are the migration velocities of a charged retained sample component in the packed (interstitial) and the open segments, respectively. They have been expressed [7,21] as

where

a Upacked:~e~

(Vpacked~ Ze fl

(1.54)

and

Uopen = ~ea, open Mce, open Lope.

(1.55)

where Pep is the electrophoretic mobility of the sample component under consideration, k' is its chromatographic retention factor and Mlc and Mce are the respective chromatographic and electrophoretic M-factors (from "mechanism" or "migration") of a given sample component and defined as

M/e-

1 l +k'

(1.56)

Mce, packed= 1 +

Pep

(1.57)

~l,ao,packed

and

Mce, open = 1 +

(1.58) a

~eo, open

Migration and Electroosmotic Flow

29

Evidently, the two M-factors are particular to the chromatographic and electrophoretic properties of a sample component, namely, the retention factor and the electrophoretic mobility and are the weighing factors for the relative importance of the chromatographic and the electrophoretic mechanisms on the migration velocity of the component under investigation. When either the chromatographic or electrophoretic M-factor is unity, the migration of the sample component does not involve that mechanism. Further, an M-factor greater than unity means that the sample component moves with a velocity higher than that of the EOF by virtue of that mechanism represented by that M-factor. On the other hand when a component's M-factor is smaller than unity, it will migrate at a velocity lower than that of the EOF in that particular segment. In view of the above equations, we can define M-factor, Mcec, for the electrochromatographic process in each segment as follows

Mcec, packed = Mlc

Mce,packed

(1.59)

and

(1.60)

Mcec,open -= Mce, open

Further, Eqs. 1.54 and 1.55 can be rewritten in terms of the electrochromatographic M-factors as

blpacked = Mcec, packed bleao,packed

(1.61)

Uopen ~- Mcec, open Uea, open

(1.62)

and

a

a

where Ueo,packed and Ueo,open are the mobile phase velocities in the packed and the open segments of the CEC column, respectively. For a migrant pair X and Y, the ratio of their M-factors that are listed in Table 1.1 for the four conceivable cases. In Cases 1 and 2 the two components differ only in one M-factor, in Cases 3 and 4 both M-factors, Mlc and Mce, are different for X and Y. Figure 1.12 illustrates the effect of changing the length of the packed segment in a CEC column on the separation of five sample components A, B, C, D and E. The

Referencespp. 37-38

Chapter 1

30

TABLE 1.1 FOUR CASES TO ILLUSTRATE THE CEC SEPARATION OF TWO COMPONENTS, X AND Y, IN TERMS OF THE RELATIVE MAGNITUDE OF THEIR CHROMATOGRAPHIC AND ELECTROPHORETIC M-FACTORS

Case No.

Relative magnitude of M-factors of X and Y

Mlc, x Mlc, Y

Mce, packed, X Mce, packed,Y

1

41

=1

2

=1

41

3

>1

>1

4

>1

<1

simulated electrochromatograms correspond to separation performed using four columns having )~ = Lpacked/L values of 0, 0.3, 0.7 and 0.8. The chromatographic and electrophoretic properties of the components are listed in the inset in Fig. 1.12. It follows that when )~ = 0.9, the whole pre-detection segment of the column is packed so that Lopen, 1 = O. Case 1. The electrophoretic mobilities and the corresponding M-factors of the two components are identical. Since their chromatographic retention factors and thus their Mlc are different, their Mcec,packed will also be different so that X and Y are separable by CEC, e.g., components A and B are both neutral, differing only in their chromatographic retention factors and comigrating when X = 0 as illustrated in Fig. 1.12. The separation of A and B improves with increasing ~ and optimum separation is achieved when )~ is 0.9, i.e., the detection window is located immediately after the retaining frit as in conventional CEC columns. Case 2. The chromatographic retention factors and the Mlc of X and Y in this case are identical. Since their electrophoretic mobilities and Mce are different, there Mcec will be different too and the two components can be separated in CEC, e.g., components B and C have the same chromatographic retention factors, but different electrophoretic mobilities. Hence, they are separated by virtue of differential electrophoretic migration as illustrated in Fig. 1.12. Case 3. X has a smaller chromatographic retention factor and a higher electrophoretic mobility in the EOF direction than Y. So Mlc, y > Mlc, Y and Mce,X > Mce, Y in the two segments. Thus, Mcec, X > mcec, Y in both the segments so that chromatographic and the electrophoretic forces act in concert to facilitate the separation of X and Y

Migration and Electroosmotic Flow

Separation parameters

Sample components

kt ~ep [ [Ll'eo,packed

r

31

A

nl

cI

D

E

1

2

2

1

0.5

0

0

0.4

0.5 - 0.35

I +All

0.8

0.7 o t~

"~~I

0.3 e,, o

ffl t-

E

a

0

D 0

A+B

~~E

i 6

12

Time [min]

18

24

Fig. 1.12. Simulated chromatograms illustrating the separation of components A, B, C, D and E by CEC upon changing the dimensionless packed length, ~, from 0 to 0.8.. Values of the chromatographic retention factor, k', and the velocity factor, ~tep,/~teo,packed, are listed in the inset. Conditions used are" V = 20 kV; L = 30 cm; Lopen,2 = 3 cm; Apacked/Aopen = 0.4; ~;p = 0.4; r I = 10-3 N m -2 s"l, ~teo,packed= ~teo,open =3.33 x 10-8 m 2 V -1 s-1

by CEC, e.g., component D is not only less retained chromatographically than component C, but also has a higher electrophoretic mobility in the direction of the EOF. As a result of this synergetic effect, the separation of C and D is facilitated both by chromatographic and electrophoretic mechanisms as seen in Fig. 1.12. Case 4. X has smaller chromatographic retention factor and smaller electrophoretic mobility in the EOF direction than Y so that Mlc, X > Mlc, Y and Mce, X < Mce, Y in the two segments. The final order of migration in this case depends on the relative magnitude of the chromatographic and the electrophoretic forces participating in the separation that can be adjusted by changing the relative lengths of the packed and open segments, e.g., component C has a higher chromatographic retention factor and electrophoretic mobility in the direction of the EOF than component A. As discussed earlier in Case 4, interactions with the stationary phase tend to slow down the migration of C with respect to that of A. On the other hand, due to its codirectional electrophoretic mobility C migrates in the mobile phase with a velo-

References pp. 37-38

Chapter 1

32

city higher than that of the neutral component A, that migrates with the EOF velocity. At low ~, values, where only a small fraction of the capillary is packed, the order of migration is determined by the electrophoretic mobilities of the compounds and C becomes the faster migrating component. However, when ~ values are high, most of the capillary is packed and the migration velocity of the components is dominated by chromatographic retention. It is seen in Fig. 1.12 that A elutes after C when L < 0.8. However, when ~, = 0.8, the two peaks comigrate due to the countervailing effect of the chromatographic and electrophoretic separation forces and then separate again albeit in a reversed migration order at higher ~,'s. The same applies to components B and E. Due to its smaller chromatographic retention factor, E migrates faster than B, while its counterdirectional electrophoretic mobility slows it down. It follows from Fig. 1.12 that at low X values E migrates slower than B as electrophoretic migration dominates the separation. However, at higher ~, the chromatographic retention is the dominant mechanism and so B slows down due to its higher k' value. So the separation of peaks B and E decreases upon increasing of ~, from 0 to 0.3 and further to 0.7. When ~, = 0.7, the countervailing chromatographic and electrophoretic forces make the bands of B and E comigrate. Finally at ~ = 0.8, the two components are separated again though their migration order is reversed. Figure 1.12 illustrates that the design of the CEC column in terms of the dimensionless packed length, ~, has a profound influence on the separation when the sample contains both neutral and charged components. It is seen that satisfactory separation of all five components can be achieved with CEC columns having ~, values in the range 0.2-0.6. Nonetheless, the speed of separation has also to be considered in most cases. The separation is almost two times faster when X is about 0.2 than when ~ = 0.6. Thus in the case illustrated, a CEC column with a packed length of 20 % can provide a rapid and efficient separation of components A, B, C, D and E.

1.5.8 Higher separation efficiencies in CEC CEC has emerged as technique offering very high efficiencies in comparison to HPLC indicating superior mass transfer in the column in CEC [8,15,17,19,30]. The mass transfer processes of convection and diffusion that are the key to separation in CEC, take place in the interstitial space and so it is important to understand flow characteristics in the near vicinity of the packing particles. Figure 1.13 illustrates flow through a bed packed with porous particles and Ul and u2 are the EOF velocities through the small intraparticle pores and the larger interstitial channels, respectively. For pressure driven flow, assuming that the path length and the pressure drops are the same for the two cases the flow velocity varies as square of

Migration and Electroosmotic Flow

33

E

,~

Intraparticle channel

jt

~ Interparticle Y~"- channel u2

u1

Fig. 1.13. Schematic illustration of intra- and interparticle EOF through porous particles in CEC.

a

packed bed of

the channel diameter and thus u2 is much larger than Ul as given by the following expression

(1.63)

where dl and d2 are the channel diameters for the intraparticle pores and the interstitial channels, respectively. Evidently, this difference in velocity caused by varying cross sectional areas of the flow channels causes excessive peak broadening as the sample travels from inlet to outlet of the column. On the contrary, since the EOF in CEC as given by Eq. 1.4 is independent of the channel diameter

U2 ~ Ul

(1.64)

and so the two velocities are nearly equal for similar path lengths. This feature of CEC Referencespp. 37-38

Chapter 1

34

I ~:~,

I~ , ~

~~ ~ m~,~l

Fig. 1.14. Schematic illustration of EOF generated in a cylindrical tube with non-uniform zeta potential at the tube wall [56].

where the EOF velocity is independent of the channel diameter plays an instrumental role in attaining high plate efficiencies. In a recent study the influence of the longitudinally non-uniform zeta potential in CZE was studied [56]. The results showed that when the electrolyte goes through a charged tube with a neutral band in the middle with zeta potential zero, vortices of the kind shown in Fig. 1.14 are formed. Similar results are expected for the case when instead of the zeta potential the field strength varies in an intermediate portion of the tube. This result attains importance in the case of CEC as there the pores in the packing particles go in all directions like those in Fig. 1.13 and so mass transfer in and out of such pores could be enhanced by vortices set up due to change in electric field strengths inside such pores in comparison to the interparticular channels. Presence of these tiny 'mixers' along the path length provides enhanced mass transfer in comparison to pressure driven chromatography where such vortices would be absent. Further, when electric field is applied to a capillary that is closed at one end, EOF as shown in Fig. 1.15 is set up in the tube. This situation is very similar to that of a dead end pore in a packing particle in CEC, where flow is generated at the pore wall and causes the electrolyte to accumulate at the dead end, followed by pressure driven flow at the center in the reverse direction. Unlike in pressure driven flow where mass transfer through such a pore would depend entirely on the diffusive flux, in CEC this flow would greatly enhance the mass transfer.

E

Fig. 1.15. Schematic illustration of EOF generated in a dead end pore in CEC.

Migration and Electroosmotic Flow

35

The above discussion highlights some of the phenomena that contribute to higher separation efficiencies in CEC. The net effect of these is enhances mass transfer and more uniform flow distribution through the column, both of which contribute to obtaining higher number of plates when compared to HPLC. 1.6 CONCLUSIONS This chapter contributes to the understanding of flow through the CEC column and of the generation and control of the EOF. It also provides a set of equations allowing calculation of the various electrochromatographic parameters including electrical conductivities, actual potential drops, electric field strengths and the flow velocities in the different column segments. In handling and reporting experimental results it is important that the parameters be evaluated in the correct manner. In order to facilitate this, we present a scheme in Figure 1.16 that makes it very convenient to carry out such calculations [26,27]. The results of are expected to clarify the meaning and significance of the parameters used for characterization of porous media with regard to some electrokinetic phenomena. Rapid separations in CEC can be obtained with CEC columns having a relatively short packed segment and a subsequent pre-detection open segment. Moreover, when the separation of the sample components comes about by

Applied potential (V), Measured currents ( i*oPen, i* Packed), Segment lengths (L o en, L acked), Capillary radiuPs (a) p

~ Eqs. 9,10 Conductivity ((~open, (~,packed)

~ Eqs. 11,12 Potential drops (Vopen,Vpacked)

f Eqs. 18 Conductivity ratio

((Topenl(~packed)

Capillary radius (a), Viscosity 01), EOF mobilities (l~eoopen, ~eo packed), Free cross-se~t,onal are~ (A_acked), Bed permeability (B~ .~mbient pressure, (Pn)

Effective length (Le)

/

~ Eqs. 15,16 Electric field strenghts (Eopen Epacked)

Intersegmental pressure (Pi)

Eqs. 44,45 Flow velocities (U aeo.open~Uaeo,packed)

Fig. 1.16. Sequential evaluation of various electrochromatographic parameters from experimentally measured quantities in the two upper blocks by using the indicated equations of the present work. References pp. 3 7-38

Chapter 1

36

an interplay of chromatographic and electrophoretic forces, the lengths of the packed and open segments can be adjusted to shift the balance of the two forces, and thus enhance the selectivity of the system particularly when the two forces are acting in opposite directions. This is of particular interest in CEC method development, where tailoring the packed length of the CEC column to the separation problem at hand can afford higher speed and better selectivity than would be obtained by using CEC columns of conventional configuration in the analysis of samples containing both neutral and charged components.

1.7 SYMBOLS AND ABBREVIATIONS

a ae

A B~ c

C d e

E F Fv i Io kB L Ld Le m Mce Mcec Mlc Pi PO r R T to tr u u

a

V

% q/cf

x z

Capillary tube diameter Radius of the hypothetical tube Free cross-sectional area of a column segment Specific Darcy's law permeability Molar concentration of an ionic specie Molar concentration of the buffer Diameter of a particle of a pore Elementary charge on an electron Electric field strength across a column segment Faraday's constant Net viscous force Current through a column segment Bessel's function of zeroth order Boltzmann's constant Length of a column segment or the capillary Distance between the inlet and the point of detection of the capillary Equivalent length of the hypothetical tube substituting the packed segment Empirical constant Electrophoretic migration factor Electrochromatographic migration factor Chromatography migration factor Intersegmental pressure between the packed and open segments Ambient pressure Radial coordinate Resistance of a column segment or the capillary Absolute temperature Migration time of an inert tracer in a column segment Total migration time of a sample component Electrosmotic or electrophoretic velocity Actual electrosmotic or electrophoretic velocity Potential drop across a column segment Total column volume Volume of the interstitial space Moles of an ionic specie per mole of buffer Valency of an ionic specie

Greek Alphabets (x

E

Eo

Dimensionless packing parameter Dimensionless constant Dielectric constant of the medium Permittivity of the vaccum

Migration and Electroosmotic Flow

37

Interstitial porosity of the packed bed Electrokinetic porosity of the medium Ratio of the conductivities of the packed and the open segments Viscosity of the bulk solution Debye screening parameter Dimensionless packed length Mobility of the electrosmotic flow or electrophoretic migration Mobility of an ionic specie Conductivity of a column segment Totuosity of the packed segment Surface potential on a charged surface Zeta potential at the surface of the particle or the capillary wall

~p gT q~ n K

V G"

~o

Subscripts eo

open P packed w

Pertains to electrosmotic flow Pertains to properties of open segment Pertains to packing particle Pertains to properties of packed segment Pertains to the wall of the capillary

Acronyms CEC CZE EOF HPLC

Capillary electrochromatography Capillary zone electrophoresis Electrosmotic flow High performance liquid chromatography

1.8 REFERENCES

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

H. Strain, J. Am. Chem. Soc., 61 (1939) 1292. H. Strain and J. Sullivan, Anal. Chem., 23 (1951) 816. V. Pretorius, B.J. Hopkins and J.D. Schieke, J. Chromatogr., 99 (1974) 23. J.W. Jorgenson and K.D. Lukacs, J. Chromatogr., 218 (1981) 209. J.H. Knox and I.H. Grant, Chromatographia, 24 (1987) 135. J.H. Knox and I.H. Grant, Chromatographia, 32 (1991) 317. A.S. Rathore and Cs. Horwith, J. Chromatogr. A, 743 (1996) 231. K.D. Altria, N.W. Smith and C.H. Turnbull, J. Chromatogr. B, 717 (1998) 341. J. Sffthlberg, Anal. Chem., 69 (1997) 3812. A.I. Liapis and B.A. Grimes, J. Chromatogr. A, 877 (2000) 181-215. A.S. Rathore and Cs. Horvfith, J. Chromatogr. A, 781 (1997) 185. M.G. Cikalo, K.D. Bartle and P. Myers, J. Chromatogr. A, 836 (1999) 25. D. Li and V.T. Remcho, J. Micro. Sep., 9 (1997) 389. L.A. Colon, K.J. Reynolds, R.A. Maldonado and A.M. Fermier, Electrophoresis, 18 (1997)2162. M.M. Dittmann, K. Wienand, F. Bek and G.P. Rozing, LC-GC, 13 (1995) 800. A.L. Crego, A. Gonzalez and M.L. Marina, Crit. Revs. Anal. Chem., 26 (1996) 261. S.E. van de Bosch, S. Heemstra, J.C. Kraak and H. Poppe, J. Chromatogr. A, 755 (1996) 165. A.S. Rathore, E. Wen and Cs. Horwith, Anal. Chem., 71 (1999) 2633. G. Choudhary and Cs. Horv~th, J. Chromatogr. A, 781 (1997) 161. A.S. Rathore and Cs. Horvfith, Anal. Chem., 70 (1998) 3069. A.S. Rathore and Cs. Horvfith, Anal. Chem., 70 (1998) 3271.

38 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

Chapter 1 C. Yang and Z. E1 Rassi, Electrophoresis, 20 (1999) 18. H. Poppe, J. Chromatogr. A, 778 (1997) 3. R. Stol, H. Poppe and W.Th. Kok, J. Chromatogr. A, 887 (2000) 199. M.G. Cikalo, K.D. Bartle and P. Myers, J. Chromatogr. A, 836 (1999) 25. C.K. Ratnayake, C.S. Oh and M.P. Henry, J. High Resol. Chromatogr, 23 (2000) 81. C.K. Ratnayake, C.S. Oh and M.P. Henry, J. Chromatogr. A, 887 (2000) 277. G.E. Eykholt, J. Haz. Mat., 55 (1997) 171. R.N. Zare, D.J. Rakestraw, D.S. Anex, R. Dadoo, H. Zhao, D. Nier, J.-R. Chen, M. Dulay and C. Yan, Lecture No. 1277, presented at the 49th Pittsburgh Conference, New Orleans, LA, March 1-5, 1998. M.M. Dittmann and G.P. Rozing, J. Chromatogr. A, 744 (1996) 63. R.J. Hunter, Foundations of Colloid Science, Clarendon Press, Oxford, 1986, p. 395. R.J. Hunter, Zeta Potential in Colloid Science Principles and Applications, Academic Press, London, 1988, p. 11. P.C. Hiemenz and R. Rajagopalan, Principles of Colloid and Surface Chemistry, Marcel Dekker, New York, 3rd ed., 1997, p. 499. F. Urban, H.L. White and E.A. Strassner, J. Phys. Chem., 39 (1934) 311. Q. Wan, J. Phys. Chem. B, 101 (1997) 4860. J. Lyklema and M. Minor, Colloids and Surfaces A: Physicochemical and Engineering Aspects, 140 (1998) 33. R.W. O'Brien and W.T. Perrins, J. Colloid Interface Sci., 99 (1983) 20. N.I. Zharkikh, X. Pendze and S.S. Dukhin, Colloid Journal, 56 (1994) 573. J.H. Masliyah, Electrokinetic Transport Phenomena, Aostra, Alberta, 1994, p. 99. M. von Smoluchowski, in I. Graetz (ed.), Handbuch der Elektrizit/~t und des Magnetismus, Barth, Liepzig, 1921, p. 366. M. von Smoluchowski, Bull. Intern. Acad. Sci. Cracovie, (1903) 184. C.L. Rice and R. Whitehead, J. Phys. Chem., 69 (1965) 4017. J.T.G. Overbeek, in H.R. Kruyt (ed.), Colloid Science, Elsevier, New York, 1952, p. 194. J.R. Boyack and J.C. Giddings, Arch. Biochem. Biophys., 100 (1963) 16. G.E. Archie, Trans. AIME, 146 (1942) 54. S. Liu and J.H. Masliyah, in L.L. Schramm (ed.), Suspensions: Fundamentals and Applications in the Petroleum Industry, American Chemical Society, Washington, DC, 1996; p. 227. P. Wong, J. Koplik and J.P. Tomanic, Phys. Reb. B, 30 (1984) 6606. R.W. O'Brien, J. Colloid Interface Sci., 110 (1986) 477. R.E.D. Rue and C.W. Tobias, J. Electrochem. Soc., 106 (1959) 827. M.A. Ioannidis, M.J. Kwiecien and I. Chatzis, Transport in Porous Media, 29 (1997) 61. Q. Wan, J. Phys. Chem. B, 101 (1997) 8449. J.T.G. Overbeek and P.W.O. Wijga, Rec. Trav. Chim., 65 (1946) 556. J.T.G. Overbeek, in H.R. Kruyt (ed.), Colloid Science, Elsevier, New York, 1952, p. 115. K. Kobayashi, M. Iwata, Y. Hosoda and H. Yukawa, J. Chem. Eng. Japan, 12 (1979) 466. C.E. Schwartz and J. M. Smith, Ind. Eng. Chem., 45 (1953) 1209. B. Potocek, B. Gas, E. Kelmdler and M. Stedry, J. Chromatogr. A, 709 (1995) 51.

Chapter 2

Instrumentation for Capillary E le c t r o c h r o m a t o g r a p h y Gerard P. R O Z I N G 1,*, Anna D E R M A U X 2 and Pat S A N D R A 2

1Agilent Technologies GmbH, Waldbronn Analytical Div&ion, P.O. Box 1280, D 76337 Waldbronn 2State University of Gent, Organic Chemistry Department, Krijgslaan 281 (S. 4.), B 9000 GENT, Belgium

CONTENTS

2.1 2.2

2.3

2.4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Instrumentation requirements . . . . . . . . . . . . . . . . . . . . . . . . 2.2.1 General . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.2 Temperature control . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.2.1 Thermal effects . . . . . . . . . . . . . . . . . . . . . . 2.2.2.2 Cooling of capillaries . . . . . . . . . . . . . . . . . . . 2.2.2.3 Effect of temperature on mobility . . . . . . . . . . . . 2.2.2.4 Band broadening caused by temperature effects . . . . . 2.2.3 High-voltage power supply . . . . . . . . . . . . . . . . . . . . . 2.2.3.1 Applying voltages above 30 kV . . . . . . . . . . . . . 2.2.4 Injection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.4.1 Mode of injection . . . . . . . . . . . . . . . . . . . . . 2.2.4.2 Quantitative aspects of injection . . . . . . . . . . . . . 2.2.4.3 Influence of the injection plug length on the efficiency Gradient-CEC instrumentation . . . . . . . . . . . . . . . . . . . . . . . . 2.3.1 Voltage programming . . . . . . . . . . . . . . . . . . . . . . . . 2.3.2 Step gradient of mobile phase composition . . . . . . . . . . . . . 2.3.3 Continuous gradient of mobile phase composition . . . . . . . . . 2.3.3.1 Field generated gradient . . . . . . . . . . . . . . . . . 2.3.3.2 EOF driven gradient . . . . . . . . . . . . . . . . . . . 2.3.3.3 Pressure assisted gradient CEC . . . . . . . . . . . . . . Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4.1 UV/VIS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4.1.1 High-sensitivity detection cell . . . . . . . . . . . . . .

40 41 41 43 43 47 48 48 52 52 54 54 54 . 56 58 58 59 60 60 62 71 75 76 76

Chapter 2

40

2.5 2.6

2.4.1.2 Fluorescence detection . . . . . . . . . . . . . . . . . . 2.4.1.3 Nuclear magnetic resonance detection . . . . . . . . . . 2.4.1.4 Photothermal absorbance detection . . . . . . . . . . . . 2.4.1.5 Condensation nucleation light scattering detection . . . . 2.4.2 Alternative methods to enhance detection sensitivity . . . . . . . . 2.4.2.1 Bandbroadening due to detection . . . . . . . . . . . . . Summary and conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

78 79 80 80 81 81 82 83

2.1 INTRODUCTION Capillary Electrochromatography (CEC) has enjoyed a high interest and fast development in the past 5-10 years leading to a quickly growing number of publications in this field. Much progress has been achieved and the technique is on the verge of becoming an alternative to HPLC and CE in practical, routine analyses. CEC is a chromatographic separation method in which the mobile phase is driven through a stationary phase by electroosmotic flow (EOF) using an electrical field. Typically, a polyimide clad fused silica (FS) capillary tube, 50-200 lam I.D., which is packed with reversed phase HPLC type stationary phase particles, is used as a separation column. Precautions are made to retain the particles in the packed bed. Solvents like in HPLC are used to affect separation e.g. acetonitrile-aqueous buffer mixtures. An electrical field up to 1000 V/cm is applied to generate an adequate velocity of the EOF. Partitioning between mobile and stationary phase like in HPLC separates neutral solutes. Charged solutes are separated by the combined action of partitioning between stationary and mobile phase and by their difference in electrophoretic mobility in the electrical field. Normally, UV-VIS spectrophotometric light absorption through a window on the capillary tube just after the packed bed, which is freed from the polyimide layer, is used for detection. With this global description the basic elements of a system for CEC have been identified viz. a column, high voltage power supply and UV-VIS spectrophotometric absorbance detector (see Fig. 2.1). Theory of CEC and methodological aspects of the technique are described exhaustively in the literature [1,2,3,4,5] and in this monograph and will not be dealt with here further. In this chapter, the current status of instrumentation for CEC is reviewed. General requirements, solvent delivery, and detection will be dealt with in some detail. Coupling of CEC separation with electro-spray ionization mass spectrometry (ESI MS) is of particular importance and is treated in a separate chapter of this monograph.

41

Instrumentation

la.p,el v

4

1, 3 " Buffer vials, 2 Packed 9 Fused Silica Capillary 50-200 ~ i.d., 2 0 0 - 5 0 0 mm length 4 Electrodes 9 5 "Power supply 6 'Point of detection 7 Ext. 9 pressure 2-12 bar Fig. 2.1. Schematic view of a CEC system. 1 and 3, buffer vials; 2, packed fused silica (capillary 50-200 ~m i.d., 200-500 mm length); 4, electrodes; 5, power supply; 6, point of detection; 7, ext. pressure 2-12 bar.

2.2 I N S T R U M E N T A T I O N R E Q U I R E M E N T S

2.2.1 General

In its simplest form a CEC instrument includes following basic components: a. A high-voltage power supply. b. A system for delivering solvent and/or sample to the inlet and outlet of the packed capillary.

References pp 83-85

42

Chapter 2

c. A column in which an EOF is generated and electrochromatographic separation processes take place. d. A column compartment for thermostatting the capillary column. e. A detection device able to register the concentration profile in the eluent leaving the separation column. A simple sketch of such a system is depicted in Fig. 2.1. Since the elements of this system are present in instrumentation for capillary electrophoresis, CE equipment, homemade or commercial, has been successfully used for CEC. The high voltage power supply, 5, is required to deliver a high voltage (30 kV) at only a low power (10 Watt). Automated change of solvent and sample vials, 1 and 3, with the added capability to fill or replenish the vials is provided in commercial CE equipment. The capillary column, 2, typically fused silica tube, 0.05-0.2 mm I.D., 5-50 cm long, and packed with 3-5 btm reversed phase HPLC type stationary phases, is hold in a thermostatted compartment. At the point of detection, 6, the polyimide coating on the fused silica is removed, to allow spectrophotometric detection at low UV wavelength. However, one modification of commercial CE equipment, which is regarded obligatory, is the pressurization of the solvent vials. There is a general consensus that pressurization at inlet and/or outlet ends of the CEC column is needed to prevent formation of bubbles. The pressurization techniques can be divided into two different modes. One possibility is the application of a hydraulic pressure gradient, sufficiently high (> 50 bar) to cause an additional pressure driven flow through the column. The resulting mode is called pressure-assisted electrochromatography (PEC) and will be described in section 3 of this chapter. The second approach is an equal pressurization of both inlet and outlet vials with an inert gas at relatively low pressures of 10 to 12 bar as shown in Fig. 2.1. The reasons for formation of bubbles in packed column CEC has not been explained satisfactorily. Bubbles may be formed due to local differences in EOF velocity (e.g. between unpacked and packed sections of the capillary [6,7]), by local differences in field strength (leading to "hot spots"), by release of gas trapped in the pores or electrochemically formed [8]. Whichever mechanism applies, it was suggested by early workers in CEC, to pressurize the inlet and outlet vials, in order to keep the gas dissolved [3,9,10]. Once the bubbles form, the detector base line becomes very noisy and the current unstable. This may lead to break down of the current and the flow stops. Robson et al. illustrated that using pressurization of the solvent vials CEC can be carried out routinely at high fields with high speed and high efficiency [ 11]. Performing reproducible CEC requires stringent control of parameters like temperature, voltage, and pressure. In commercial CE equipment, many of these

Instrumentation

43

parameters are automatically controlled which has led to significant improvements in reproducibility and accuracy of the separations. Currently, most manufacturers of CE instrumentation also provide the option for pressurization of both outlet and inlet vials (see Table 2.1). The Hp3DcE system (now Agilent Technologies Capillary Electrophoresis System, Waldbronn, Germany) has the option to apply a gas pressure of 0.2 to 1.2 MPa to the outlet and/or inlet vial [ 12]. A pictorial view of the system is shown in Fig. 2.2. Isocratic CEC separations are easily performed in this equipment, which accounts in part for the initial success and acceptance of CEC. However, for advanced operation, an extensive modification of the equipment is obligatory and will be described later in this chapter. In a review of the technique, Majors claimed that CEC needs dedicated instrumentation, designed specifically for CEC and not adapted from CE. According to the same author, this was the general consensus in discussions amongst leading researchers in CEC [13, 14]. Shortcomings in current commercial CEC instrumentation include the lack of a reliable instrument allowing (i) gradient elutions and voltages higher than 30 kV, (ii) rapid changes in column temperature, (iii) use of capillaries with varying lengths, and (iv) interfacing to modern sampling formats such as 96 (384) well plates. These shortcomings have led to further development of CEC instrumentation.

2.2.2 Temperature control 2.2.2.1 Thermal effects

The control of temperature is vital in CEC. During the electrochromatographic separation process, electrical energy is released as Joule heat. Part of the thermal energy is dissipated to the environment, while the other part is consumed by the medium resulting in an increase of the temperature in the capillary. The ion mobility depends on the solvent viscosity. Upon the increase in the temperature of the solution, its viscosity decreases causing a further increase in the current, thus releasing more heat (Fig. 2.3) [15]. If the heat dissipation is insufficient, the temperature may raise considerably to a point at which the eluent boils thus forming vapor bubbles. This leads to breakdown of the current. In addition, the increase in temperature results in an unequal temperature across the column diameter, leading to different flow velocities across the capillary cross section, and, consequently to reduction in the column efficiency

[16]. For that reason, as in CE, narrow capillaries with a high

surface-to-volume ratio are used in electrochromatography, so that heat dissipation is more effective. Since the effectiveness of radial heat dissipation decreases with the increase of the capillary diameter, a maximum internal diameter suggested for packed CEC capillaries is 0.2 mm. Obviously, effective temperature control, actually cooling

References pp 83-85

TABLE 2.1 LIST OF COMMERCIAL SYSTEMS USEFUL FOR CEC

Essential specifications

URL

Manufacturer

System

Agilent Technologies, Waldbronn, Germany

Agilent Capillary Forced-air temperature cooled with Peltier element. http://www.chem.agilent.com/cag/ products/hp3dce.html Electrophoresis System Temperature range 10°C below ambient to 60°C (0.1 "C) with a minimum of 4°C. Minimum total capillary length 33.5 cm Minimum effective capillary length 8.5 cm. Vial pressurization to 12 bar. Several modes of operation, CE, CEC, CE-MS, CE + pressure. CEC capillaries 250 x 0.1 mm and 400 x 0.1 rnrn, CIS-, C8- and Phenyl type phases

Beckman Coulter Inc., Fullerton, USA

http://www.beckman.com/ PIACE MDQ Methods Temperature control of capillary column with rapidly beckman/biorsrch/prodinfo Development System recirculating liquid coolant. Selectable temperature control Icapelecl paceseries.asp between and 15°C and 60°C (0.1"C). Solvent delivery provided by both variable pressure (0.1 to 100 p.s.i).

Biomolecular Instruments, Santa Fe, USA

Spectraphoresis Ultra

http://www.biomolecularl .corn/ Peltier controlled 10°C below ambient to 60°C in O.l°C increments. Constant voltage, constant current, voltage gradients, positive or negative polarity, time-programmed polarity switches current gradients, constant power. Pressure at one or both ends of a capillary with the voltage or current can be applied. Minimum capillary length 22.5 cm; maximum length 63.5 cm; shortest detection length ( L d ) 6.3 cm.

5 h

Crystal 300 Series

Constant voltage or constant current, positive or negative pressure, simultaneous pressure and voltage. 10400°C max. (identical with PrinCE System) Pressure: -180 - 250 mbar, 1 mbar increments.

2

$ 9

%

Microtech Scientific &, Inc., Sunnyvale, USA CII

http://www.biomolecularl.corn/

http://www.microlc.com/ Ultra-Plus I1 Capillary Temperature controlled from 5°C above room to 85°C. CE 1 CEC Module Columns 50 pm to 500 pm fused-silica capillary, 1.0 rnm to 2.0 rnm for micro bore. Lenght 1 cm to 100 cm for capillary, 1 cm to 25 cm for micro bore. Flow rate range 0.1-500 pllmin. pressure limit 10,000 p.s.i. Power supply voltage from 0 to 60,000 V.

Unimicro Technologies TriSepTMCEC System Gradient CEC system which includes two PU-980 HPLC Inc. Pleasanton, USA CEC, HPLC and CE pumps, flow rate range 1 pVmin to 10 ml/min, -30 or +30 kv. Electropak CEC columns. ProLab Instuments Evolution 200, GmbH, Kagenstrasse MicroHPLC System 17, CH - 41 53 Reinach, Switzerland

Binary, high-pressure gradient pump. Flow rate range 1-200 pllmin in gradient mode, isocratic 0.1-200 pllmin. Pressure range 0 4 0 0 MPa. Integrated vacuum degasser. Programmable high voltage power supply 30 kV, 200 pA.

http://www.unimicrotech.com/ index.htrn1

Chapter 2

46

"v" t~ ~.,=_

~o t~

"0

~ .Ig e~ o t~

t~

~D r/3 rat3 ~r/3

~E e~

~D

m

el-

O

O

E t_ ,

e.. m

O .,..a ~D

-n O t_ t~

~

o~ t_

r..)

!

(D

"o

(D Cr

.,..~ ~z0

<

(D

O ~~D

.o (D

.X:

C./3

e,i e-i 60

of the packed capillary is as mandatory in CEC as in CE. A quantitative treatment of this subject is given in the next section. Using a system fumished with an effective thermostatting, Euerby et al. [17] claimed that 15~ is an ideal temperature for separations in CEC.

Instrumentation

~'T (w/m~

47

0.025 (air) 0.15 1.4 0.6 0.2-0.6 (liquid)

parabolic logarithmic

z (oc)

polyimide (10-20gm)

y[(25-100 lumen gm) !

fused silica (100-200 gm) Fig. 2.3. Thermal effects in electro-driven separation techniques. Temperature profile across a capillary containg electrolyte and heated by passage of an electric current.

Since increased temperature is known to improve separation in HPLC, Sandra et al. performed CEC separations of triglycerides at temperatures of up to 50~ and did not observe any bubble formation or breakdown of the current [ 18]. However, Knox and McCormack [ 19] demonstrated that high temperature might also affect the sample injection. If the rate of thermal expansion of the liquid within the capillary is faster than the rate of electromigration of the slowest component, some compounds present in the injected sample may not enter the capillary. 2.2.2.2 Cooling of capillaries The applied field E, the equivalent conductivity A of the buffer (m2f2-1mo1-1) and the buffer concentration c (mol/L) determine the heat that is generated in an open capillary, Qv (W/cm3).

Qv= EZA c

Referencespp 83-85

(2.1)

Chapter 2

48

Qv can reach hundreds of watts per cubic centimeter. Because the heat is exchanged with the environment, an unequal temperature profile is generated over the capillary cross-section (Fig. 2.3). Therefore, for separations using high electric field strengths exceeding 200 V/cm, active cooling is essential. It has been shown that a strong stream of air (> 10 m/s) can reduce the temperature in the capillary by a factor of 5 [2]. Therefore, thermostatting systems of some CEC instruments are based on forced air circulation, which can effectively stabilize temperature even in capillaries run at high current levels. As opposed to liquid cooling, the use of forced air flow eliminates the need to disassemble the cooling system when installing or removing capillary columns. Forced air also requires less maintenance than liquid and is cheaper. In contrast, cooling using a liquid appears to be more efficient [20,21,22]. It should be born in mind that even in effectively cooled systems, an increase in temperature can result in a non-linear increase in current [ 15].

2.2.2.3 Effect of temperature on mobility The mobility of the solutes and ions present in the buffer solution and, therefore, the EOF and migration/retention times of the solutes depend on the temperature through the effect of temperature on viscosity and dielectric constant. Mobilities increase approximately 2-3 % per I~ This may affect resolution but enable faster analyses if the separations at hand are relatively simple. However, reproducibility of migration times is strongly affected by changes in viscosity, which in turn is a further argument to well-control the temperature in the packed capillary column. By using the constant current-mode instead of the constant voltage-mode (see part 3 of this chapter), the effect of viscosity changes can be eliminated.

2.2.4 Band broadening caused by temperature effects While electro-driven flow affords some remarkable advantages over pressuredriven flow with regard to zone broadening [1,3,4], the realization of the highest performance is limited by the heating occurring in the former systems. Poppe et al. [23,24] considered the effects of homogeneous heat release in packed columns in detail. Although their equations are general and complex, they lead to a simple result for an infinitely long cylinder within which the heat is generated homogeneously and lost by conduction through the walls. The temperature difference between the center of such a cylinder and its walls is given by equation

Instrumentation

49

AT= Q ~ 16~,v

(2.2)

where Q is the rate of heat generation per unit volume of the cylinder, ~T is the thermal conductivity of the medium, and de is the cylinder diameter. In a pressuredriven system, variation in temperature leads to a corresponding variation in eluent viscosity, and consequently in flow rate. Thus, a velocity variation of 1 or 2 % across the column is found in standard pressure-driven chromatography using a 5 mm bore column [ 1]. The situation is different in electro-driven systems. Here, the generation of heat arises from the current through the electrolyte filling the capillary and is given by

V2

(2.3)

Q-R

where V is the voltage across the capillary and R is its resistance (f2.m). Introducing the molar electrical conductivity of the solution ~, and replacing (l/R) we obtain

Q = E2 ~ c g

(2.4)

where c is the concentration of the solution and ~ is the total porosity of the medium. The value of ~ is 1 for an open capillary, in which case equation 4 reduces to equation 1, and 0.4-0.8 for a packed capillary. The excess of temperature at the center of the capillary is then calculated using equation 2 and is given by

AT=

E2 L c e ~

(2.5)

16 Xv

The heat release per unit volume is about 1,500 times larger for electro-driven systems than for pressure-driven systems under typical operation conditions! Although

self-heating

can

have

deleterious

effects

on

performance

in both

pressure-driven and electro-driven systems, there is an important difference between the overall effects in these two modes. In pressure-driven systems, which use wide

Referencespp 83-85

Chapter 2

50

bore columns, the dispersing effects of self-heating can rather easily be removed by radial diffusion. In a CEC capillary, the parabolic temperature profile across the capillary results in a corresponding parabolic velocity profile, which is superimposed upon the main flow velocity of the eluent (EOF) within the column. The dispersion effect of this velocity profile may be treated, using a slightly modified Taylor equation developed for the calculations of dispersion in an open capillary. The velocity excess at the center of the column is denoted by 2Av and the velocity of the main band by v. To analyze this situation, imagine an observer moving at a velocity v with the main band. He will observe a situation similar to a stationary observer monitoring the flow in an open capillary: he will see a band moving slowly along the capillary, with a velocity Av. During the movement of the band along the whole column length L, the moving observer will see a band which appears to move a distance (Av/v)L. The Taylor equation can then be applied to obtain the dispersion arising from this effect:

o2 (Av/v)L

~ Av 96 Dm

(2.6)

In equation 2.6, Dm is the diffusion constant of the solute in the mobile phase. The height equivalent to theretical plate H is then given as:

H = t~'L--

~v

(~/~

(2.7)

The velocity-spread Av which arises from the self-heating can be expressed as

Av q wall qce,,,re zXn 2-" = = = 0.026 AT V

q wall

(2.8)

q

in which 1"1is the viscosity of the solvent and, therefore,

Av

(2.9) - 0.013

AT

Instrumentation

51

From the Smoluchowski equation,

(2.10)

y= r ~;~~ E 1]

where e0 is the permitivity of a vacuum, er dielectric constant of the eluent and ~ is the zeta potential. Substitution of equation 9 for

(Av/v), equation

2.5 for AT and

equation 2.10 for v leads to the final expression for H:

H~

~

~'~ O'O13E2Xc~'~}I] 16 XT

(2.11)

96 Dm

7" 10-9 ~30~r ~ ~2 2 a~cE s c 2 H~

(2.12)

Dm TI ~2

Using representative values: ~ = 50 mV, ~,= 0.015 m-2mollf2 -1, ~ = 0.75, dc = 100 lam, E = 50,000 V.m-1, c = 0.01 M, Dm = 10-9m2s -1, rl = 10-3 N.s.m -2, and )~Y = 0.4 W.m-IK -1 gives,

H = 0.006 ~tm

(2.13)

Clearly, this value is very small and, in practice, negligible when compared to typical HETP-values of 5 to 10 ~tm. However, one notes that H is proportional to

dc6E5c2. Therefore,

small changes in column diameter, applied field and electrolyte

concentration can have dramatic effects on H arising from the self-heating. For example if the column diameter is increased to 200 ~tm, AT increases 4 times to 1.8 K and H rises 64 times to 0.4 ~tm. If, in addition, E is doubled, H may reach the unacceptable value of 12 ~tm. However, a decrease in ionic strength c to 0.002 mol/L would bring H down again to 0.5 ~tm.

Referencespp 83-85

Chapter 2

52

2.2.3 High-voltage power supply Typical velocities in pressure-driven LC columns packed with 3 to 5 ~tm diameter particles are 0.5-3.0 mm/s. To obtain these flow rates in packed CEC columns using the same stationary and mobile phases, requires a field of 100-1000 V/cm. Appropriate power supplies will be able to apply 30 kV but the current is limited to 300 ~A. Under standard CEC conditions, the currents are in the range of 1-30 ~tA and very stable. With state-of-the-art instrumentation, the current can be monitored which is very useful. Usually, the voltage is applied through electrodes in the solvent vials. The high voltage is on the injection side of the column and the detector side is grounded. This reduces the risk of damage to the detector and the noise. On most commercial systems, the polarity and magnitude of the voltage, the current, and the power can be controlled using the controlling software. The standard polarity setting is positive meaning that the positive electrode is at the inlet vial. Reversing the polarity to the negative mode means that the inlet vial becomes the negative electrode. The outlet electrode is grounded and is connected with the power supply to measure the current flowing through the capillary. Voltage, current, and power are related to each other. The control software of most commercial instruments allows to set values for each of these three parameters independently thus operating in "constant voltage", "constant current" or "constant power" - mode. The "constant voltage"- mode applying 0 to 30 kV is commonly used by CEC-practitioners. An interesting feature of commercial instruments is voltage programming during the analysis (see section 3 of this chapter). This allows applying high voltage in a 10 s ramp at the beginning of the analysis to avoid stress to the column [ 12].

2.2.3.1 Applying voltages above 30 kV Applying voltages higher than 30 kV results in an increase in mobile phase velocity and thus reduces the time of analysis. Hutterer and Jorgenson demonstrated the feasibility of extending the applied potential above 30 kV and the separation power of CE using a home built ultra-high voltage capillary electrophoresis system. A commercial power supply was extensively modified in order to provide electrical potentials of up to 120 kV. A unique electrical shielding system was developed to prevent capillary breakdown and corona or spark discharges [25]. Choudhary and Horv~ith constructed a homemade CEC system, which can be operated at up to 90 kV [26]. The system is presented in Fig. 2.4. The 90 kV voltage power supply was composed of a bipolar 30 kV and a unipolar 60 kV power supply. The system allows the application of 90 kV with a cathodic EOF and a voltage drop

53

Instrumentation

10

11

~

6

"

Ibl

7

I

I

m

m

12"~"" 15 IB

mt

I

n

"-"16

oH 1

3

mmn n

-

17

Fig. 2.4. Schematic view of the modular capillary electrochromatograph with a 90 kV dual power supply and pressurisable chambers for column inlet en outlet (reproduced from Ref. [26] with permission of the publisher). 1, 60 kV power supply; 2, 30 kV power supply; 3, digital electrometer; 4 and 5, electrodes; 6 and 7, reservoir for mobile phase or the sample; 8, pressurisable chambers; 9, packed capillary column; 10, cell for on-column detection; 11, detector; 12, four-port two-way valve; 13, four-port three-way valve; 14, pressure gauges; 15, from nitrogen cylinder; 16, vent; 17, ground.

up to 30 kV with an anodic EOF. A 1.4 cm thick PVC protective spacer block was mounted between the UV detector and the on-column detector cell to prevent arcing. Yan et al. [27] manufactures an instrument, which can be operated in both CEC and micro-HPLC mode. The high voltage power supply can provide up to 60 kV. A stainless-steel six-port rotary valve that also includes an injection port serves as the injection manifold, which is inserted in the electric circuit. The instrument is partially housed within a Faraday cage.

References pp 83-85

Chapter 2

54

2.2.4 Injection 2.2.4.1 Mode of injection Because of the narrow capillary diameter, small sample volumes in the range of nL have to be injected in order to preserve the high efficiency. Samples may be introduced either in hydrodynamic (by applying pressure) and/or electrokinetic mode. In the latter, the sample is introduced in the column by applying a voltage (0-30 kV). Since the sample is introduced by combining electroosmotic flow and electrophoretic mobility, the amount of sample loaded depends on the charge of individual analytes and the nature of the sample matrix. While this is likely to have little impact in qualitative analysis, it requires that both standards and samples are run in a homogeneous, closely defined matrix for quantitation. Rigorous control of the sample matrix is less important for hydrodynamic injection, since sample loading is only a function of applied pressure. The use of hydrodynamic injection does have some limitations because of the very high backpressure required to drive the mobile phase through packed capillary columns. Therefore, the injection pressure using packed columns must be in the 0.2-1.2 MPa range.

2.2.4.2 Quantitative aspects of injection For both electrokinetic and hydrodynamic injection the injected quantity Qi in the plug with length Li, can be calculated from the injection time ti, the linear velocity of the injection plug vi, and the sample concentration ci (Fig. 2.5):

~t~

rt~

Q~= Li c~ ~4 = vi ti c i - - ~

(2.14)

Where de is the capillary diameter. Note that the following equations are only valid for open capillaries. In the case of electromigration, vi is related to the mobility of the sample components, but also to the EOF and the applied voltage during injection Vi [28]:

~t Vi v~= L =

(~eo -b ~ep) Vi L

(2.15)

Instrumentation

55

Li -4

t.-

driving force

injectiontime

Fig. 2.5. Plug-like injection by applying voltage or pressure difference.

or

~:~

(2.16)

Qi = V, t~ (~eo + ~tep) Ci 4 L

Because laep is different for each of the sample components, faster moving ones are proportionally oversampled, although this effect can be reduced when electroosmosis is much faster than the electrophoretic mobility of the sample constituents [29]. Pyell and Rebscher [30] proposed that the injection length in CEC for neutral compounds is

L~ =

t, ~l,eo V/ L

(2.17)

where laeo is the electroosmotic mobility, Vi is the injection voltage, and L is the total column length. Inserting equation 2.17 in equation 2.14 gives equation 2.16 for ~tep=0. The value of ~teo is determined using a non-retained neutral compound.

References pp 83-85

Chapter 2

56

•m = gep -t- ~teo-

LaL tmV

(2.18)

Combining equations 216 and 2.18 results in

V~t~ r i f L e Q,-- V eo

(2.19)

-

For hydrodynamic injection, the Poiseuille relation determines the linear velocity.

vi-

AP, ~ 8Lr I 4

(2.20)

Combining equations 2.14 and 2.20 results in

1 rt ~

Q; = APi ti c , 1"1 128L

(2.21)

From equation 2.21 it is clear that discrimination among the solutes does not occur.

Until now, electrokinetic mode of sample injection was employed in the majority of CEC analyses. Ross et al. [ 16] compared the reproducibility of both electrokinetic (20 kV/5 s) and hydrodynamic injection (5 bar/15 s). The relative standard deviation (r.s.d.) for retention times (n=5) varied between 0.26-0.37% and 0.28-0.40% for electrokinetic and hydrodynamic injection, respectively. The r.s.d. (n=5) for the peak heights varied between 0.81-1.64% and 2.81-3.80% for electrokinetic and hydrodynamic injection, respectively. Sandra et al. confirmed this accuracy for the analysis of triglycerides [ 18].

2.2.4.3 Influence of the injection plug length on the efficiency The injected plug length is an important parameter that affects extra-column band broadening [30]. Assuming that the contribution due to detection and data processing can be neglected, the tolerable injection plug length for a column of a given size and

57

Instrumentation

efficiency can be derived. The variance for the injection plug can be calculated from the injection plug length Li [31 ].

2

L2

(2.22)

o~- 12

The variance for the separation process is given by

2 L2 or N

(2.23)

where Ld is the length of the column to the detection window and N is the plate number. The total variance can be obtained by addition of equations 2.22 and 2.23

4L~ +

L~

(2.24)

-i5+

If reduction in the calculated plate number NR is considered as criterion for the tolerable injection plug length and if NR is given by

NR = N (1 - 6)

(2.25)

where 6 is the reduction parameter, the maximum plug length which meets this requirement is

12 8 L L~= 4 1 - 8 4-N

(2.26)

A reduction parameter 6 of 0.091 corresponds to a 10 % increase in peak variance. The dependence of the peak width on the injected plug length for alkyl and aryl benzoates in packed CEC was investigated experimentally [30]. At low injection plug Referencespp 83-85

Chapter 2

58 length

(Li

_< 1 mm) CYT is independent of

Li, while at high volume overload

~T is

linearly dependent on Li. 2.3 GRADIENT-CEC INSTRUMENTATION The vast majority of CEC analyses reported until now were performed in the isocratic mode with equipment similar to that shown in Fig. 2.1, with the option of pressurizing one or both ends of the capillary column. However, the analysis of a complex mixture containing compounds with a wide range of molecular weights and polarities may require acceleration of the elution of late-eluting components to shorten analysis time. In LC, this is done by solvent gradient elution. In contrast, gradient elution is more difficult to achieve in CEC because of constraints related to high voltage, sample introduction, and pressure. So far, several approaches to gradient CEC have been published. Moreover, three gradient CEC systems are currently commercially available (see Table 2.1). To broaden the applicability of CEC however, reliable gradient instrumentation must be generally and broadly available. Several types of gradients can be applied in CEC such as temperature gradient, voltage gradient, solvent step gradient, and continuous solvent gradient. Voltage programming, stepwise gradient, and several types of continuous gradients have already been reported in the literature and will be discussed in the following part.

2.3.1 Voltage programming Xin and Lee [32] described the technique of voltage programming to accelerate the elution of late-eluting components. The advantages of voltage programming can be summarized as follows: 9Voltage programming is simple to realize and does not require expensive equipment like LC gradient pumps or additional high voltage power supplies. 9 Voltage programming is efficient since the voltage can be controlled precisely. It is also highly reproducible. 9 Problems related to sample introduction, high voltage isolation, or other typical of other gradient methods are avoided. Samples can be injected in both electrokinetic and hydrodynamic mode without affecting other aspects of the instrument operation or causing sample waste. 9 The selectivity of separation does not change in the course of the voltage programming, making method development easy and straightforward. 9 The baseline is very stable because the composition of the mobile phase does not change, thus making quantitation more accurate.

59

Instrumentation

power

supply

!1 l/V l:~ete~r -

-II

I

I I

~_

Fig. 2.6. Systematic diagram of the home-built CEC system with voltage programming capability. (Reproduced from Ref. [32] with permission of the publisher).

9 Both column ends can be pressurized to prevent bubble formation without inducing pressure driven flow during the analysis. This mode appeared to be an effective alternative to mobile phase gradient programming for the analysis of polyaromatic hydrocarbons (PAHs). The equipment for the voltage programming experiments consisted of a computer controlled high voltage power supply, two pressurized electrolyte reservoirs, and a fiber optic assembly for UV detection (Fig. 2.6). A constant voltage of 15 kV for 11 min was applied and then linearly increased to 40 kV at a rate of 10 kV/min. The flow velocity changed from 0.4 to 2.8 mm/s, using the acetonitrile-50 mM TRIS pH 8.1 (80:20 v/v) mobile phase.

2.3.2 Step gradient of mobile phase composition An alternative approach to a continuous gradient, which can be performed using standard CEC instrumentation, is a stepwise gradient. This is achieved by changing the inlet vial during the chromatographic run for a new buffer vial containing a different mobile phase. Ding and Vouros demonstrated that stopping the flow during the CEC run does not affect the efficiency of the analysis even with a stop-time as long as several minutes at room temperature [33]. The same group demonstrated the use of the step gradient to improve speed of the CEC analysis of DNA adducts derived from syn benzo[g]chrysene-11,12-dihydrodiol13,14-epoxide. They used a three-step gradient, which started with a quaternary mobile phase followed by two ternary mobile phases. The analysis was completed in References pp 83-85

60

Chapter 2

78 min, which is about 25 % of the total analysis time required to separate the same 8 adducts by LC. While applying a step gradient, it is important to stop the flow at the appropriate time to prevent overlap of the EOF peak with any of the analyte peaks [34]. Euerby et al. [35] applied this technique to the analysis of a mixture of six diuretics with widely differing lipophilicity. The step gradient resulted in a remarkable reduction in analysis time and improved peak shape for the later eluting analytes. However, unstable baseline with numerous disturbances were observed which may limit the applicability of this approach in screening of impurities in pharmaceuticals. Zhang et al. investigated two modes of step gradients for the separation of mixtures of ketones, aldehydes, and aromatic hydrocarbons. The first approach involved changing the inlet and outlet vials as described above. The second method involved dropping another mobile phase directly into the stirred inlet vial using a pipette. Both methods shortened the analysis time and improved the detection limits compared to isocratic elution. The separation of a mixture of 13 aromatic hydrocarbons was achieved in 14 min by changing the mobile phase from 80% methanol to 80% and 90% acetonitrile, respectively [36].

2.3.3 Continuous gradient of mobile phase composition The interest in continuous gradient CEC is increasing. At present, three approaches have been used to create this type of gradient in CEC. In the first, two high-voltage power supplies are used to generate an EOF from two different mobile phases in two separate capillaries, which are connected via a T-piece where they mix, and enter the column. No mobile phase is wasted in this system. However, the exact composition of the mobile phase entering the column is unknown. In the second approach, a conventional gradient HPLC-pump supplies the mobile phase with a varying composition to the inlet vial where the EOF transports the solvent through the column. Although the use of a LC pump allows accurate control of the mobile phase composition, major part of the mobile phase is wasted and a small pressure driven flow may be created, which may reduce column efficiency. In a third approach, a high-pressure gradient pump is connected directly to the inlet of a packed capillary in which electro- and pressure driven flow operate simultaneously. This approach is called pressurized capillary electrochromatography (PEC) or electrically assisted micro HPLC. 2.3.3.1 Field generated gradient

Zare's group used the electrical field applied to two separate fused silica capillaries to generate a gradient of mobile phase composition [37,38]. The system is shown in Fig. 2.7. Merging two flows, which voltages were controlled by a computer, generated

Instrumentation

61

Computer

power =.p,.+l

i

i

Higl~voltaoe I '

.

~]

power j -I

supply 2

= L--_ ....J

Captllaz7 2

Capillary 1

Mobile-phase reservoir 1

Microscope objective

Separation column

Filters

Lens

.~.",-'"

....... , .....

,,

Moblle-,phasa reservoir 2

....

Lens

?

Photomu[1Jplier tube

.......

o~

[O"

acquisition

Laser

Outlet reservoir

=

Fig. 2.7. Schematic diagram of the solvent gradient-elution CEC apparatus. (Reproduced from Ref. [29] with permission of the publisher).

a mobile phase composition gradient with submicroliter flow rates. The two fused silica capillary arms were attached to a T-connector serving as a mixer. The inlet of one capillary arm was placed in the solution reservoir containing first mobile phase, while the inlet of the other was placed in the second reservoir containing a different mobile phase. The resulting mobile phase flowed from the mixer in the packed capillary column. Two independent computer-controlled, programmable, high-voltage power supplies (0-50 kV) were used to apply ramping high-voltage potentials to both mobile phase reservoirs to regulate the EOF in each arm. The ratio of the EOF rates in

both arms was changing as a function of time to deliver the required gradient to the separation column. An open tubular capillary was used for the evaluation of the performance of the solvent gradient system. Fluoranthene, added to the mobile phase in the second reservoir served as a fluorescent tracer to indicate the percentage of this solvent

constituting the final mobile phase. The time between the beginning of the linear portion of the voltage program and the onset of the increase in measured fluorescence References pp 83-85

62

Chapter 2

was used as an indicator for the response time of the gradient system. The relative standard deviation for three consecutive runs of the time interval was found to be less than <1%. Samples were introduced electrokinetically into the separation column by disconnecting it from the T-connector and placing its inlet into the sample vial. After injection, the column was reconnected to the T-connector and the gradient-elution initiated. Similar to gradient-LC, baseline drift represents a problem in gradient CEC. The authors observed severe baseline drift with the increasing acetonitrile content in the mobile phase. 2.3.3.2 EOF driven gradient

Recently Lister et al. [39] demonstrated a flow-injection analysis interface for gradient CEC. In this system, the solvent is driven through the column only by EOF (Fig. 2.8). Solvent gradients were generated in a micro-LC system and delivered to the interface. Injections were carried out on-line using a rotary LC valve controlled by an electric actuator. Using a gradient of acetonitrile-water from 60:40 to 90:10 allowed baseline separation of nine solutes in less than 18 min. This interface includes an auto-injector and dramatically decreases the need for manipulation with the packed column thus improving its lifetime. Taylor et al. [40,41] constructed a gradient LC-CEC interface consisting of a stainless steel T-connector and a restriction capillary (Fig. 2.9). Samples were introduced using a LC auto-sampler into the mobile phase stream, which flows perpendicularly via a fused silica capillary towards and past the CEC column inlet. Voltages of up to 30 kV were applied to the T-piece to generate EOF in the CEC column. On the outlet side, a similar grounded T-piece was used. The split ratio and LC flow rate were adjusted in such a way that both column ends were pressurized to suppress bubble formation without generating a significant hydrodynamic flow. The reproducibility of injection was demonstrated by repetitive injections (n=6) of a standard solution of corticosteroids at two different days. R.S.D. for retention times on day 1 and day 2 were 0.51 and 0.93, and for peak areas 2.49 and 1.78, respectively. The injection linearity in terms of analyte concentration and injection volume was also good (r 2 > 0.998) [40]. Retention time and peak area reproducibility for corticosteroids extracted from urine was characterized by R.S.D. 2 and 7 % respectively [41 ]. Due to differences in UV transmission of the mobile phase, the baseline drifted with increasing acetonitrile concentration. A prototype gradient-CEC instrument has also been developed by Hewlett-Packard (now Agilent Technologies, Waldbronn, Germany). This instrument offered the ability to perform all isocratic as well as gradient micro-LC, CEC, and pressure assisted or pseudo-CEC (PEC) without having to remove the column upon switching between

Capillary

n

T

Pt electrode

Fig. 2.8. Right panel; schematic diagram of instrument setup for EOF driven gradient CEC according to Lister et al. [39]. Left panel; diagram of FIA CEC interface (Reproduced from Ref. [39] with permission of the publisher).

PEEK N l p m . n l Tub.

CEC Columm

-WMr C .pU1.v

Fig. 2.9. Left panel; schematic of a gradient CEC system according to Taylor et al.; 1 and 2, waste capillaries; 3, CEC column; 4, high voltage sampling interface; 5, loading capillary; 6 , gradient HPLC pump; 7, HPLC autosampler; 8, high voltage power supply; 9, groundedwaste interface; 10, UV detector; 1 1, waste reservoir; 12, HV power cable; Right panel; cross section of the CEC sampling interface (Reproduced from Refs. [40,41] with permission of the publisher).

Instrumentation

65

the modes. The instrument consisted of two separate pieces of equipment, namely Hp3DcE and an HP 1100 LC binary pump, controlled by special versions of firmware and software (see Fig. 2.10). A unique cassette provided the coupling between these two instruments. A double tube electrode replaced the inlet electrode of the CE instrument. The solvent was delivered from the pump to the inlet vial via a restriction capillary connected to the double tube electrode (Fig. 2.11). Excess solvent was removed from the vial via the outlet restriction capillary. This vial could be placed in two positions by the inlet lift. In the lower position of the lift, EOF driven isocratic and gradient CEC was possible. The solvent was delivered by the pump to the vial through the inner channel and removed via the outer channel using a pressure of nitrogen (right panel of Fig. 2.11). The resistance of the outlet restrictor was low. The pressure in the vial equaled the external gas pressure and was the same in the outlet vial. In this way, no hydraulic flow was generated and bubble formation was suppressed. The column continuously accepted the delivered solvent from the vial by electroosmotic flow. In this position, the flow delivered by the external pump must be low to avoid solvent overflow in the vial. The upper position allowed isocratic and gradient micro-LC with the possibility to assist the separation with an electric field. The capillary was introduced into the electrode and the solvent was delivered to the capillary via the inner channel. The solvent overflow was liberated through the outlet capillary, which had a higher flow resistance in this case. In this position, the special vial functions as a flow splitter. This versatile system can be used in different modes. For example, a high speed separations are possible upon unidirectional application of both electroosmotic and hydraulic flow (Fig. 2.12). However, the primary use of this device is gradient elution CEC illustrated on the examples in Fig. 2.13. Euerby et al. [42] evaluated this continuous gradient CEC prototype system in pharmaceutical applications. The reproducibility was assessed by repeated separations of a standard test mixture consisting of thiourea, benzamide, anisole and biphenyl. The R.S.D. for the retention times of the analytes was better than 1 % for simple linear, concave, convex and linear gradients with isocratic steps. In the case of a simple linear gradient the R.S.D. found for retention times varied from 0.09 to 0.42%. Using the continuous gradient system, the separation of a mixture of various diuretics was achieved in a shorter time compared to isocratic elution and sharper peaks for the late eluting peaks were observed thus improving the sensitivity. Huber et al. [43] used a commercial capillary electrophoresis system connected to a LC pump to achieve the gradient elution. His approach is shown in Fig. 2.14. Two LC pumps were connected to one end of a capillary restrictor through a static mixing T-piece. The other end of the restriction capillary was connected to an injection valve. This valve was connected to a poly (ether ether keton) (PEEK) cross (17.2 ~tL) which

References pp 83-85

CEC-capillary

I

Restriction capillaries

Binary pump

m-fl\

f i

Normal outlet vial

electrode k i a l

Grounded u n i w

WilXlC

Spceial 2 in I inlet vial

Fig. 2.10. Schematic diagram of the Agilent Technologies prototype gradient CEC instrument.

67

Instrumentation

Capillary

Capillary

Solventremoval ,-Solventdelivery

Solvent removal Solventdelivery Nitrogen delivery

Inlet vial

Inlet vial

Fig. 2.11. Schematic drawing of the special inlet vial of the Agilent Technologies prototype gradient CEC instrument. Left graphic, pHPLC gradient mode; right graphic, EOF driven gradient CEC mode.

mAU 16C 14(

Pressure Voltage

12C

115 bar 25 kV

10C 80' 60'

5

40"

1 2 Pressure

20"

Voltage

'''

2'''

4.'''

6'''

8'''

10'''

12'''

115 bar

OkV 14'''

16'

min

Fig. 2.12. Cmparison of pHPLC with field assisted pHPLC separation of aromatic hydrocarbons using the Agilent Technologies prototype gradient system. Column, CEC Hypersil Phenyl 250 (335) x 0.1 ram. Mobile phase A, acetonitrile-50 mM trihydroxymethylamine, pH 8 (95:5 v/v); B, 2.5 mM trihydroxymethylamine, pH 8; A/B 85:15, dtection 250 nm; sample, 1, thiourea; 2, dimethylphthalate; 3, diethylphthalate; 4, biphenyl; 5, o-terphenyl References pp 83-85

2.5

5

7.5

10

12.5

15

17.5

20

22.5 min

5

10

15

20

25

rnin

Fig. 2.13. Example of EOF driven gradient CEC (right) and field assisted pHPLC (left) separation of a rabbit urine sample with , (335) cyclic peptide drug metabolites on Agilent Technologies prototype gradient system. Column CEC Hypersil C18, 3 ~ r n 250 x 0.1 mm. Solvent A, methanol-12.5 mM ammonium acetate (1:9); B, methanol-ammonium acetate-acetonitrile (10:12.5:77.5); gradient 30-90% B in 20 min. Voltage 25 kV, detection 209 nm, injecton 150s, 5 kV. Sample concentration is few pM.

69

Instrumentation

1

i r

r Fig. 2.14. Capillary electrochromatograph with gradient elution capability according to Huber et al. (reproduced from Ref. [43] with permission of the publisher). 1, high-voltage power supply" 2, inlet reservoir with electrode; 3, outlet,.r~servolr with electrode; 4, packed capillary column; 5, on-line sensing unit of UV detector; 6, detector output (0-1 kV); 7, sample injection valve; 8, purge valve; 9, restrictor; 10, syringe for introduction of sample or buffer; 11, capillary restrictor; 12, static mixing tee; 13, grounding; 14, pumps; 15, pump control panels and readouts; 16, manometer; 17, eluent reservoirs; 18, switching valve; 19, syringe for buffer introduction; 20, waste reservoir at the inlet; 21, waste reservoir at the outlet; 22, thermostated inlet compartment; 23, detector compartment; 24, outlet compartment; 25, CEC instrument control panel; 26, gas pressure control; 27, gas inlet (1.4 MPa nitrogen); 28, temperature control; 29, data acquisition. gill

',

.

was used as an inlet reservoir. The two horizontal ports of the cross were used to connect the capillary column and the platinum electrode. The vertical openings of the cross were connected to the valve. The 3 mL outlet reservoir also made from PEEK was filled with the mobile phase using a plastic syringe. The reproducibility of the gradient profile was determined from the change in the UV absorbance at the column effluent using 5% acetone solution in the inlet reservoir. R.S.D. (n=5) of 3% were found for retention times for phenyl thiohydantoin-glycine (PTH-glcyine) and PTHtyrosine. Recently, Alexander et al. [44] reported the use of an automated separation system developed for micro-LC and CEC using both isocratic and gradient elutions. The complete system is shown in Fig. 2.15. An enlarged view of the coupling of the column to the injection valve presents Fig. 2.16. The mobile phase was delivered by two micro-LC pumps at a flow rate of 30 ~tL/min to a post injection splitter that houses the column inlet. In the CEC mode, pressure was not applied (no restriction on

References pp 83-85

70

Chapter 2

I1

m

m 9--~ 5

I---1

Fig. 2.15. Schematic automated isocratic and gradient elution nano-liquid chromatograph/ capillary electrochromatograph according Alexander et al. (reproduced from Ref. [44] with permission of the publisher). 1, high-voltage power supply (negative polarity); 2, platinum electrode; 3, outlet reservoir vial; 4, UV detector with on-column flow cell; 5, nanocolumn; 6, two-position switching valve; 7, jack stand; 8, fused-silica make-up adapter (split device); 9, ground cable; 10, internal loop micro-injection valve; 11, plexiglas compartment; 12, autosampler; 13, dynamic mixer; 14, micro-LC pumps.

6 ----~

t,

t 2

t 3

t 4

t 5

Fig. 2.16. Schematic of the coupling of the nanocolumn to the injection valve [44]. 1, nano-column (75 ~tm I.D. x 360 ~tm O.D.); 2, knurled nut and ferrules; 3, fused-silica make-up adapter (split device 504 ~tm I.D.); 4, micro-injection valve (column port 250 pm i.d.); 5, rotor; 6, outlet port of the make-up adapter.

splitter) to the column inlet or outlet and the voltage was continuously applied during both sample injection and mobile phase delivery. In the LC mode, a restrictor was attached to the splitter.

Instrumentation

71

Most recently, Que and co-workers proposed a very simple device to generate a solvent gradient for CEC [45]. In this case, a small capillary shaped reservoir 0.4 mm I.D., filled with the weak solvent, was connected to the front of a CEC column. The reservoir was connected to the inlet vial via a very narrow fused silica capillary (0.05 mm I.D.). After sample injection into the CEC column, the reservoir was filled with the strong solvent. Due to the high velocity of the solvent in the narrow inlet capillary, the strong solvent in the reservoir was mixed with the starting buffer solution and the mobile phase with increasing solvent strength driven out from the reservoir. In all of these implementations, the required solvent transport is achieved by electroosmotic flow although some of the systems have the built in pressure driven flow ability. 2.3.3.3 Pressure assisted gradient CEC

Experiments in CEC has shown though that the magnitude of EOF strongly depends on solvent composition and the type of stationary phase. Since these parameters are typically varied to optimize the separation, it may happen that EOF is significantly reduced under conditions, which are optimal for the separation. Coupling of both solvent transport by EOF and separation in a single column is certainly a weakness of CEC. Therefore, many attempts have been described in which the hydraulic flow is assisting the electrochromatographic separation. As mentioned earlier, that it is possible to simultaneously apply a high hydraulic pressure at the inlet vial to drive the solvent through the column and an electrical field. This hybrid method combining both pressure and electrodriven liquid chromatography was first introduced by Tsuda [46] and is referred to as pressure assisted CEC or field assisted micro HPLC. Verheij et al. [47] used this the technique to couple CEC with MS. These authors also proposed the name pseudo-CEC for this technique. Tsuda's group constructed a PEC system [48] consisting of a HPLC pump, an injector, a home made capillary column, a UV detector, and a high voltage power supply (0-50 kV). A fused silica restriction capillary was used to control flow splitting. The high voltage was applied at the stainless steel T-piece while the electrolyte reservoir was grounded. Hugener et al. [49] and Eimer and Unger [50] developed similar home-made systems. The latter applied high voltage to the outlet reservoir and the splitting Tpiece was grounded. The system was operated in both micro-LC and PEC mode. Three organic acids, which could not be separated in the micro-LC mode, were baseline resolved applying an additional electric field across the capillary. Adjustments in pressure and voltage controlled the selectivity.

References pp 83-85

Chapter 2

72

l GradientMixer] [ HPLCPump ]IZ~ Injection i DataAquisitionI Waste ~

Sp ~'~rl--~_ Earth

CapillaryColumn

ElectrolyteReservoir

ElectrolyteRese~'oir

High-Voltage PowerSupply

Fig. 2.17. PEC System according to Behnke and Bayer (reproduced from Ref. [51] with permission of the publisher).

Behnke and Bayer published a similar approach for gradient elution PEC [51 ]. The schematic view is shown in Fig. 2.17. A gradient mixer and a HPLC pump were combined with a modular CE system. A post-injection splitter was used and sample introduced by a conventional HPLC six-port injector. A grounded stainless-steel Tpiece was used to split both eluent and sample. The electrolyte reservoir on the inlet side of the separation capillary was connected to the splitter by a homemade interface. Behnke and Metzger [54] used a LC system with an auto sampler and solvent delivery system. Splitting of the eluent was achieved by a stainless steel T-piece connected to a 50 pm I.D. fused silica capillary that served as flow resistor. In the gradient-pseudo-CEC mode, the LC system was combined with a high voltage power supply. The separation column was connected to the splitting T-piece. The CEC separation of peptides of a tryptic digest of cytochrome c on a 60 mm x 100 ~tm I.D. column packed with 1.5 ~tm ODS afforded resolution comparable to that of conventional LC using a 250 mm long column packed with 5 ~tm particles, however, the CEC analysis time of 10-15 min was considerably shorter than that required in LC

Instrumentation

73

(45-90 min). The elution time window for the peptide mixture did not shift significantly by applying the voltage as could be expected from the small contribution of EOF at low pH to the high flow velocity generated by a pressure of 25 MPa. Gradient-PEC has been coupled to an ion trap storage/reflectron time-of-flight mass spectrometer for the analysis of peptide and protein digests [52]. Analysis of bovine cytochrome c digest was achieved in less than 12 min using a gradient of 0-50% acetonitrile, 10 kV/m, and 70 MPa supplementary pressure. Gfr6rer et al. [53 ] reported coupling of gradient CEC with NMR. The set-up described by Behnke and Metzger [54] was used in this study although only a pressure of 14 MPa was applied. In another approach, Dittmann et aL used a gradient CEC system based on standard commercial HPLC and CE modules [55]. A schematic of this system is shown in Fig. 2.18. A standard HPLC pump and an injector were connected to a T-piece in a CE--MS capillary cassette through a fused silica restriction capillary. A hole was drilled through this T-piece, shown in detail in the panel of Fig. 2.18, to fit a 0.5 mm PEEK capillary, 1.6" O.D. A small hole was also drilled in the PEEK capillary and placed opposite to the third leg of the tee. Both FS connection and CEC capillaries were placed in the PEEK tube leaving a small space between them. The outlet of the T-piece was connected to a restriction capillary through a selection valve. The separation capillary led through the detection interface to the inlet vial where the voltage was applied. The T-piece was grounded. Using this approach, the voltage direction had to be reversed to obtain correct direction EOF. Therefore, the non-metal detector alignment interface had to be used. Besides gradient elution, the hydraulic flow can be also used to assist solvent transport under conditions of very low EOF. This was illustrated by an experiment carried out using the prototype equipment shown in Fig. 2.8. The lower trace of Fig. 2.19 shows the micro-LC separation of 7 non-steroidal anti-inflammatory drugs [56]. The mobile phase consisted of acetonitrile and 25 mM KH2PO4 pH 2.5 (4:1 v/v). Hexylamine, (0.5%) was added to reduce tailing of these components. Because of the low pH, the EOF is very slow reaching only about 0.5 mm/s. The LC pump pressure was set at 120 MPa to afford a flow rate of 1.0 mm/s in the capillary column. In the micro-LC baseline, the separation of 6 analytes was achieved according to their polarity and hydrophobicity. However, timolol did not elute. Applying a voltage of 25 kV during this micro-LC separation resulted in baseline separation of all of the 7 compounds and reduced the analysis time by a factor of 5. The column efficiency calculated for antipyrine was increased by 18.8% in this voltage assisted micro-LC.

References pp 83-85

Fig. 2.18. Schematic of a pressure driven gradient CEC system according to Dittmn et al. (reproduced from Ref. [55] with permission of the publisher).

75

Instrumentation

mAU 5

6

LC (210 bar) + 25 kV

80

60

40

LC (210 bar) 20 7

~ 0

5

1'0

, 1+2

3

1'5

2'0

~5 min

Fig. 2.19. Illustration of selectivity gain in CEC compared with micro HPLC (taken from Ref. [56] with permission of the publisher). Column Waters Spherisorb ODS I, 3 ~tm, 250 (335) x 0.1 mm. Mobile phase, 80% acetonitrile-20% 25 mM phosphate, 0.2% hexylamine, pH 3.8. Voltage 25 kV, temperature 20~ Sample: 1, procaine; 2, timolol; 3, ambroxol; 4, metoclopramide; 5, thiourea; 6, naproxene; 7, antipyrine.

2.4 D E T E C T I O N

The quality of an analytical separation technique is not only characterized by its separation efficiency but also by its detection sensitivity. UV-VIS spectrophotometry and fluorescence are commonly used simple detection methods for CEC. However, not all compounds absorb UV light and even fewer compounds fluoresce. Although high mass sensitivity can be achieved, the concentration sensitivity in CEC is generally poor since short optical path lengths that equal the internal diameter of the capillary are used. Dittmann et al. [55] discussed thoroughly the limits of sensitivity in CEC detection. A number of approaches enhancing the sensitivity have been developed. For example, mass spectrometry and laser-induced fluorescence (LIF) are new detectors, representing one of these approaches. The coupling of CEC to mass spectrometric detector is discussed in detail in a separate chapter in this monograph. A lot of work References pp 83-85

76

Chapter 2

has also been devoted to improvements in the sensitivity of the most common UVVIS detector. 2.4.1 UV/VIS The typical detector used in CEC so far is the UV/VIS absorption spectrophotometer, with light transmitted through the part of the capillary from which the polyimide coating has been removed. This has a number of disadvantages including (i) small path length of the "cell", (ii) most of the light transmits through the capillary wall, (iii) reduced durability of the column as a result of removal of the polyimide coating, and (iv) detection takes place beyond the frit. With packed columns, detection is mostly performed in the unpacked section behind the frit retaining the stationary phase (on-column photometric detection or OCPD). Rebscher and Pyell [57] have reported photometric detection in the packed section of the column (in-column photometric detection or ICPD). Robson et al. reported very low reduced plate heights (h=l) when employing ICPD with columns packed with ODS [11]. Banholczer and Pyell [58] compared the sensitivity for ICPD and OCPD. Baseline noise in ICPD was about twice that of OCPD. The calibration linearity for butyl benzoate did not differ significantly. The precision in the determination and detection limits of 6 benzoates, however, were lower with ICPD compared to that with OCPD. ICPD can be considered the standard detection mode in CEC using continuous bed columns. Hilder et al. recently reported the separation of seventeen inorganic and small organic anions using a column packed with silica based anion-exchange beads and a nitrate eluent with indirect UV detection at 214 nm. The separation selectivity was shown to be a combination of ion-exchange chromatography (IC) and CE [59]. Sensitivity and linear detection range can be improved by increasing the inner diameter of the capillary. Special capillary designs can be used to extend the optical pathway without increasing the overall capillary radius. One of such designs is the "bubble cell" [60] that is made by forming an expanded region, a bubble, directly within the capillary. This bubble extends the pathway for light by a factor of 3 with almost no deterioration in separation efficiency and resolution. 2.4.1.1 High-sensitivity detection cell

A high-sensitivity cell for the Agilent Capillary Electrophoresis System has recently been developed. The cell increases detection sensitivity by an order of magnitude compared to standard detection [61 ]. This new cell is expected to increase substantially the utility of CE/CEC for the detection of enantiomeric purity of chiral drugs and trace analysis in biological and environmental samples.

77

Instrumentation

•

Removable pillary

l~ ~] ~

Fused silica

body

7

!

BillaCk~u~ed

/L

Flanking windows"-~..

.................. ht path

7"7 // // Removable capillary

Fig. 2.20. High-sensitivity cell incorporated in the optical interface.

The cell comprises a fused-silica body and removable capillaries (Fig. 2.20). The improved performance results from the extension of the detection path length from 100 [am, typical of standard CEC capillaries, to 1.2 mm. The light path through the cell is made in black fused silica, which significantly minimizes stray light. In addition, the reflective interior wall functions as a "light pipe", ensuring 100% transmission of the light that enters the cell. These properties result in enhanced linearity beyond 1.4 AU and unsurpassed spectral fidelity with the diode array detector. The cell is incorporated in an optical interface similar to that used for standard and bubble cell capillaries. This feature does not require any changes in installation of both capillary and cassette. Because poor capillary coupling can cause peak broadening and distortion, the capillaries of the high-sensitivity cell have a special geometry. The ends of these capillaries are flared on the inside, and beveled on the outside to ensure that peak shape is not distorted. An example of the increased sensitivity by using the high sensitivity cell is shown in Fig. 2.21.

References pp 83-85

78

Chapter 2

mAU 30

25

20

CEC column with high sensitivity cell

S / N - 315

Standard CEC column

S / N - 47 '

'

'

I

2

'

'

'

I

'

l

4

l

|

6

i

i

i

i

8

'

l

'

I

10

i

i

i

i

12

i

,

'

i

min

Fig. 2.21. Illustration of signal improvement by the usage of a high sensitivity cell in CEC (reproduced from Ref. [55] with permission of the publisher). Column, CEC Hypersil C18, 3 ~tm, 250 (335) x 0.1 mm; mobile phase acetonitrile-50 mM Tris-HC1, pH 8 (90:10 v/v); voltage, 25 kV, injection electrokinetic, pressure, 10 bar both sides, temperature 20~

2.4.1.2 Fluorescence detection

Rebscher and Pyell [62, 63] were the first to report the use of fluorescence detection. The baseline noise with ICFD was about twice that of OCFD. This could be attributed to the minute motions of the packed bed in the applied electric field and the diffuse scattering in the bed that increased the level of background fluorescence. R.S.D.'s found for the retention times of polyaromatic hydrocarbons were less than 1.1 and < 0.4 % for ICFD and OCFD, respectively. Variations in the peak areas were about twice as high in ICFD (2.6-5.1%) compared to OCFD (1.4-2.3%). According to the authors, this could be due to variations in the temperature within the capillary. The power of fluorescence detection was illustrated on the separation of PAH's by Yan et al. [64]. The 16 U.S. EPA priority PAHs were separated in isocratic mode in less then 10 min using 100 lam I.D. columns packed with 1.5 lam nonporous octadecyl silica particles. Separation efficiencies of 750,000 plates/m were obtained when the PAHs were detected by ICFD while 300,000-400,000 plates/m were found for OCFD.

79

Instrumentation

2.4.1.3 Nuclear magnetic resonance detection

Nuclear magnetic resonance spectroscopy (NMR) is one of the most powerful analytical methods for identification and structure elucidation of organic compounds. Since NMR spectra are recorded in solution, no phase transfer like in MS is necessary when coupled with LC techniques. Additionally, NMR is a non-destructive detection technique, allowing the analyte to be transferred for characterization using additional methods. As of today, LCmNMR coupling was used in a wide range of applications [65,66,67,68,69,70,71 ]. On-line CEC--NMR coupling is promising [72,73,74,75]. Of special interest is the coupling of gradient CEC with NMR. A Bruker AMX 600 NMR spectrometer was used to perform isocratic and gradient micro-LC--NMR and CEC--NMR. This hyphenation is presented in Fig. 2.22. Alkyl benzoates were separated on a column packed with 5 lam Gromsil ODS-2 and 10% of 3 ~m Gromsil Si using a isocratic elution with mixture of 2 mM borate D20-CD3CN (20:80 v/v). No baseline separation was achieved in LC due to the high

zoom of detection cell;

~

"1"

!

,,/, rf

9

coil

V = 240 nL fused sitica capillary i.d. 60 pm

NMR probe

, outlet v i a l - - -

I I~ ]

packed or ~ unpacked f.s. capillary ........... --=== .... -

~.._.~~

l~urestrictiOn inlet vial// ."/ T,.,~.~.~,I injection device I capillary wastel

! I

'~

i ......~~:~:~"~............ i ./" voRage

/

Capillary Electrophoresis chromatographyCapi Electroilary

I" +::'' : !,,/ ,pressure[, Io, . . . . . . . . . | ",,,

Capillary HPLC

Fig. 2.22. Coupling of capillary separation techniques with NMR (taken from Ref. [72] with permission of the publisher). References pp 83-85

80

Chapter 2

sample loading required for the NMR detection. In CEC, however, the elution time was reduced by a factor of 2 and a considerable band sharpening with improved resolution was observed [72]. The same experimental setup was used in the separation of metabolites of paracetamol from a human urine extract [73] and Thomapyrin| containing acetaminophen, caffeine, and acetylsalicylic acid [74]. Compared to isocratic CEC--NMR, gradient CECmNMR offers increased sample loading capacity because of preconcentration at the front of the column and higher separation efficiency together with a reduction in analysis time [75]. 2.4.1.4 Photothermal absorbance

An ultraviolet-laser based thermo-optical absorbance detector for micrometer capillaries was used by Qi et al. [76] to monitor the separation of a mixture of 13 phenylthiohydantoin-amino acids. A modulated pump laser beam periodically illuminated the capillary at a point near its end. Complex deflection and diffraction effects occur at the capillary-solution interface. Perturbation of the refractive index at this interface changes the intensity of the probe beam that is measured using a small photodiode. Detection limits for 13 resolved PTH-amino acids ranged from 1.6 to 4.8.10 -7 M, which was by a factor of ten superior to the detection limits reported for micellar electrokinetic chromatography [77]. Most of this improvement in detection limit is due to the higher thermo-optical enhancement produced by the high acetonitrile content in the buffer compared to the aqueous mobile phase used in the micellar electrokinetic separation. 2.4.1.5 Condensation nucleation light scattering

Guo et al. [78] demonstrated the use of condensation nucleation light scattering detection (CNLSD) coupled to a pressurized CEC system using an electrospray interface. Compared to evaporative light scattering detection (ELSD), CNLSD employs an additional step where the desolvated analyte particles are exposed to a supersaturated vapor that condenses onto the particles. This increases their size from a few nanometers to the micrometer range. As a result, the intensity of scattered light is significantly increased and, therefore, improvement in detection limit can also be achieved provided, the level of non-volatile components and contaminants in the mobile phase is minimized. Guo adapted a commercial CEC to reach a reasonable flow rate for electrospray operation and retain the sensitivity of CNLSD. Good reproducibility, comparable sensitivities for a wide range of compounds, including carbohydrates, and

Instrumentation

81

limits of detection down to the 50 ng/ml level, corresponding to 1-2 pg, were determined without the need for derivatization. 2.4.2 Alternative methods to improve detection

Other approaches to enhance the sensitivity can be sub-divided to (i) concentration outside the capillary (solid-phase extraction, etc.) and (ii) concentration within the capillary using hyphenated techniques such as isoelectric focusing, isotachophoresis, and on-column stacking. An elegant study was presented by Thomas et al. who employed selective solidphase extraction by immunoaffinity CEC (IACEC) to enhance detection limits. A model compound, fluorescein isothiocyanate biotin, was electrokinetically applied to a capillary column packed with an immunoaffinity stationary phase. The analyte was first selectively bound to the stationary phase, then eluted, migrated by zone electrophoresis, and detected by LIF [79]. Stead et al. described an on-line concentration of progesterone and its major metabolites. The sensitivity of CEC was improved by injecting analytes from a non-eluting solution [80]. The ability to focus dilute samples at the top of the column and their consecutive elution using a mobile phase with an increasing concentration of organic modifier (17-45% acetonitrile) in gradient-CEC was reported by Taylor and Teale. They injected a large volume (250 1) of a corticosteroid (100 ng/ml) and observed a 25-fold increase in sensitivity [40]. However, these approaches are sample and buffer dependent and, therefore, difficult to automate while increasing the sample preparation time. Finally, it is also possible to improve sensitivity of detection by reducing the noise level and increasing the stability of the background signal from the detectors. This signal originates from a number of potential sources, including impurities in the buffer and electrical pick-up in the detector wiring circuitry. One of the routes to background reduction and stability is digitations of data and use of mathematical algorithms to smooth the background noise and amplify the signal. Saeed et al. describes the use of an ID/10 processor to improve the sensitivity of a commercial CEC instrument. Signal enhancement from this processor for phenols in tobacco smoke was used to show improvements in the linear dynamic range and data collection for peak area, peak height, and retention time [81 ]. 2.4.2. I Band broadening due to detection

The volume restrictions that apply to sample injection are equally valid for detection. Detectors must be miniaturized relative to the column dimensions and a detection volume in the nanoliter or even subnanoliter level is often required. For References pp 83-85

82

Chapter 2

example, Yang [82] and Jorgenson et al. [83] demonstrated that detachable detection cells, a typical part of stand-alone detectors, were no longer useful. Instead, a small section of the column itself has to be used as the detection volume, which results in an on-column detection. Turbulence due to the cell geometry or originating from connectors is in this case avoided and the cell volume obviously depends on the column diameter. Consequently, the remaining variable parameter of such detection cell is the length used for illumination, from which the contribution to zone dispersion can be estimated [84,85,86]:

cY2 : L 2 12

(2.27)

L

(2.28)

Ld<

Although the tolerance is dependent on column length an other zone dispersion factors, calculations by Otsuka and Terabe [85] suggest that the detection cell length should be shorter than 1 mm. Using a different approach Wang et al. [87] came to similar conclusions, claiming that peak distortion becomes significant when Ld is longer than 1 peak standard deviation. This yields a limiting value of 1.3 mm for L = 400 mm and N = 100,000 and 1.1 mm for L=250 mm and N=50,000.

2.5 SUMMARY AND CONCLUSION For isocratic mode of CEC separations, standard CE instrumentation is sufficient. This applies particularly for equipment that has the provision of column pressurization. In practice this is achieved by applying a gas under a pressure of 2-12 bar to both inlet and outlet vials. Column thermostating in CEC is regarded mandatory to avoid excessive radial temperature gradients within the capillary. In such instruments, sample is typically injected electrokinetically and alternatively by applying the external gas pressure to the sample vial. Detection occurs 'on-column' i.e. directly through a non-packed section of the capillary following immediately the end of the bed. However, there are several shortcomings and functions missing if commercial CE systems are used for CEC. For example, the operational temperature range for chromatographic type separations is limited in current CE equipment. The cassettes used to hold the packed capillaries and the controlled temperature environment do not

Instrumentation

83

allow variations in column configurations and do not allow temperature control of the part of the capillary column resting in the vials. Concentration sensitivity of on-column UV-VIS absorption detection is poor compared to HPLC and coupling of CEC with other types of detectors such as mass spectrometry is often troublesome. However, the major deficits of commercial CE equipment used for CEC are the absence of solvent gradient delivery systems and/or assistance of the CEC separation by hydraulic flow. The latter might provide a sample and solvent flow in cases when the selected chromatographic conditions are unfavorable to generate EOF. It has been shown that hydraulic flow does not contribute significantly to zone broadening. On the other hand, pressurized flow is an easy way to optimization of separation in both chromatographic and electrophoretic methods [56]. Finally, the ability to apply field strengths higher than the currently common 30 kV typical of commercial equipment is highly desirable. Adding these functions to a CE instrument will represent a significant change in hardware, firmware, and software and, therefore, a great effort for instrumentation companies. Three innovative small companies listed in Table 2.1 went that way and lead the field in this area. On the other hand, these limitation have provided and will continue to offer a challenge for both academic and industrial research groups in the years to come. 2.6 R E F E R E N C E S

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

J.H. Knox and I.H. Grant, Chromatographia, 24 (1987) 135. J.H. Knox, Chromatographia, 26 (1988) 329. J.H. Knox and I.H. Grant, Chromatographia, 32 (1991) 317. M.M. Dittmann, F. Bek K. Wienand and G.P. Rozing, LC-GC Intemat., 13 (1995) 800. Electric Field Applications in Chromatography, Industrial and Chemical Processes, Ed. T. Tsuda, VCH, Weinheim, Germany. A. S. Rathore and Cs. Horvath, Anal. Chem., 70 (1998) 3271. U. Tallarek, E. Rapp, T. Scheenen, E. Bayer, H. van As, accepted for publication in Anal. Chem. R.A. Carney, M.M. Robson, K.D. Bartle, P. Myers, J. High Resolut. Chromatogr., 22(1) (1999) 29. N.W. Smith and M.B. Evans, Chromatographia, 38 (1994) 649. R.J. Boughtflower, T. Underwood, C.J. Paterson, Chromatographia, 40 (1995) 329. M.M. Robson, S. Roulin, S.M. Shariff, M.W. Raynor, K.D. Bartle, A.A. Clifford, P. Myers, M.R. Euerby, C.M. Johnson, Chromatographia, 43 (1996) 313. M.M. Dittmann, G.P. Rozing, J. Microcol. Sep., 9 (1997) 399. R.E. Majors, LC-GC, 16 (1998) 12. R.E. Majors, LC-GC, 16 (1998) 96. J. Vindevogel, Ph.D. Dissertation. University of Ghent, Ghent, Belgium 1991. G. Ross, M.M. Dittmann, G.P. Rozing, Internat. Lab. (1996) 10A.

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53 P. Gfr6rer, J. Schewitz, K. Pusecker, L.H. Tseng, K. Albert, E. Bayer, Electrophoresis, 20 (1999) 3. 54 B. Behnke, J.W. Metzger, Electrophoresis, 20 (1999) 80. 55 M.M. Dittmann, G.P. Rozing, G. Ross, T. Adam, K.K. Unger, J. Cap. Electrophoresis, 4 (1997) 201. 56 M.M. Dittmann, K. Masuch and G. P. Rozing, J. Chromatogr. A, 887 (2000) 209. 57 H. Rebscher, U. Pyell, Chromatographia, 38 (1994) 737. 58 A. Banholczer, U. Pyell, J. Microcol. Sep., 10 (1998) 28. 59 E.F. Hilder, M. Macka, P.R. Haddad, Anal. Comm., 36 (1999) 299. 60 G.B. Gordon, R.P. Tella and H.A.S. Martins, HP Journal, June 1995, 62. 61 See http://www.chem.agilent.com/cag/products/ce.html and check linked pages. 62 H. Rebscher, U. Pyell, Chromatographia, 42 (1996) 171. 63 H. Rebscher, U. Pyell, J. Chromatogr. A, 737 (1996) 171. 64 C. Yan, R. Dadoo, H. Zhao, R.N. Zare, D.J. Rakestraw, Anal. Chem., 67 (1995) 2026. 65 K. Albert, J. Chromatogr. A, 703 (1995) 123. 66 K. Albert, A. Bayer, Anal. Methods Instrum., 2 (1995) 302. 67 A. H/51tzer, G. Schlotterbeck, K. Albert, E. Bayer, Chromatographia, 42 (1996) 499. 68 S. Strohschein, M. Pursch, H. Handel, K. Albert, Fresenius' J. Anal. Chem., 357 (1997) 498. 69 U.G. Siedelmann, U. Braumann, M. Hofinann, M. Spraul, J.C. Lindon, J.K. Nicholson, S.H. Hansen, Anal. Chem., 69 (1997) 607. 70 J.C. Lindon, J.K. Nicholson, U.G. Siedelmann, I.D. Wilson, Drug Metab. Rev., 29 (1997) 705. 71 J.C. Lindon, J.K. Nicholson, I.D. Wilson, Adv. Chromatogr., 36 (1996) 315. 72 K. Pusecker, J. Schewitz, P. Gfr6rer, L.H. Tseng, K. Albert, E. Bayer, Anal. Chem., 70 (1998) 3280. 73 K. Pusecker, J. Schewitz, P. Gfr6rer, L.H. Tseng, K. Albert, E. Bayer, I.D. Wilson, N.J. Bailey, G.B. Scarfe, J.K. Nicholson, J.C., Lindon, Anal. Comm., 35 (1998) 213. 74 P. Gfr6rer, J. Schewitz, K. Pusecker, L.H. Tseng, K. Albert, E. Bayer, Electrophoresis, 20(1999) 3. 75 P. Gfr6rer, J. Schewitz, K. Pusecker, E. Bayer, Anal. Chem., 71 (1999) 315A. 76 M. Qi, X.F. Li, C. Stathakis, N.J. Dovichi, J. Chromatogr. A, 853 (1999) 131. 77 K.C. Waldron, N.J. Dovichi, Anal. Chem., 64 (1992) 1396. 78 W. Guo, J.A. Koropchak, C. Yan, J. Chromatogr. A, 849 (1999) 587. 79 D.H. Thomas, D.J. Rakestraw, J.S. Schoeniger, V. Lopez-Avila, J. Van Emon, Electrophoresis, 20 (1999) 57. 80 D.A. Stead, R.G. Reid, R.B. Taylor, J. Chromatogr. A, 798 (1998) 259. 81 M. Saeed, D.H. Craston, M. Depala, J. Reilly, D., Jackson, S.J. Walton, J. Chromatogr. A, 836 (1999) 15. 82 F.J. Yang, J. High Resol. Chromatogr. Chromatogr. Commun., 4 (1981) 83. 83 J.W. Jorgenson, K.D. Lukacs, J. High Resol. Chromatogr. Chromatogr. Commun., 4 (1981)230. 84 X. Huang, W.F. Coleman, R.N. Zare, J. Chromatogr., 480 (1989) 95. 85 K. Otsuka, S. Terabe, J. Chromatogr., 480 (1989) 91. 86 H.K. Jones, N.T. Nguyen, R.D. Smith, J. Chromatogr., 504 (1990) 1. 87 T. Wang, R.A. Hartwick, P.B. Champlin, J. Chromatogr., 462 (1989) 147.

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Chapter 3

Modes of CEC Separation Christopher M. J O H N S O N , A l a n P. M c K E O W N and M e l v i n R. E U E R B Y

AstraZeneca R&D Charnwood, Bakewell Road, Loughborough, Leicestershire, LE11 5RH, UnitedKingdom

CONTENTS

3.1 3.2 3.3 3.4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Definitions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Unmodified packings . . . . . . . . . . . . . . . . . . . . . . . . . . . . . M o d i f i e d packings . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4.1 R e v e r s e d - p h a s e C8, C 18, p h e n y l . . . . . . . . . . . . . . . . . . 3.4.2 The use o f m o b i l e phase additives . . . . . . . . . . . . . . . . . . 3.4.3 Ion exchange . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4.3.1 SCX . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4.3.2 SAX . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4.3.3 Mixed mode . . . . . . . . . . . . . . . . . . . . . . . Chiral stationary phases . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Gel C E C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 Monoliths . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.7 Size exclusion C E C . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.8 Gradient C E C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.9 3.10 Selectivity c o m p a r e d with L C . . . . . . . . . . . . . . . . . . . . . . . 3.11 Guidelines for the analysis o f acidic basic and neutral c o m p o u n d s . . . . 3.11.1 Neutral analytes . . . . . . . . . . . . . . . . . . . . . . . . . . 3.11.2 Acidic analytes . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.11.3 Basic analytes . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.12 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.13 A b b r e v i a t i o n s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.14 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

88 89 89 94 95 98 98 98 99

100 100 101

101 102 103

103 104 104 105 105 106 106 106

Chapter 3

88 3.1 INTRODUCTION

The separation scientist with experience gained from a LC background may tend to limit the modes of electrochromatography to reversed phase (RP), normal phase, ion-exchange and, maybe, size-exclusion. Analysts from an electrophoretic background typically use the term "CE" in a much broader sense to include the main modes of capillary zone electrophoresis, micellar electrokinetic chromatography, capillary gel electrophoresis, isoelectric focusing and isotachophoresis. As capillary electrochromatography (CEC) is a hybrid technique between CE and LC, there are actually many modes of operation ranging from those commonly used in CE to those described in LC. This is shown diagrammatically in Table 3.1. The section of the table topped by the heavy bar indicates the "packed CEC" region and it is these areas which are to be covered in this chapter. There is considerable scope for overlap between the described cells, and many further divisions of these artificial boundaries are possible, for example pressure-assisted electrochromatography (PEC) describes a continuum from pure CEC to pure gLC. Many of the modes of CEC illustrated in Table 3.1 are applicable to both gradient and isocratic elution, aqueous and non-aqueous conditions, as well as to chiral and achiral separations and these will be discussed within the appropriate sections. The complex mechanisms responsible for selectivity will not be discussed, rather this chapter will be limited to describing the scope for application of the different CEC modes. Packed CEC columns are prepared from polyimide coated fused-silica capillaries typically with 20 lam to 180 lam internal diameter (i.d.). The packing material is retained as a column in the fused silica between frits that are generated in-situ by controlled sintering of the siliceous packing by heating, although there are examples of fritless CEC columns [ 1]. The columns typically have a packed segment between the inlet frit and a frit prior to the detection window (from 7 cm to 70 cm), with the unpacked region continuing to the outlet vial. However, attempts have been made to carry out UV detection through the pack segment of the capillary [2-4]. Despite the potential advantages of increased efficiency with in-column detection, the decreased sensitivity due to the particle-induced scattering and increased background noise has made this approach less popular.

TABLE 3.1

89

Modes of CEC

3.2 DEFINITIONS

Rather than explain the detailed theory for different modes of CEC which will be dealt with in a later chapter, it was considered appropriate to collate the information as a table where mode descriptions could be compared and contrasted (Table 3.2).

TABLE 3.2 DEFINITIONS OF CEC MODES

Mode

Packing

Basis of separation

CEC gel

Gel e.g. polyacrylamide or agarose Size and charge

CEC monolith

Mouldedrigid porous polymer

Size, charge and partition

CEC packed SEC

Controlledpore size silica

Size*, charge and exclusion

CEC packed unmodified

Silica gel

Size, charge, adsorption & "reverse phase", ion-exchange

CEC modified

Silica modified with C 18, C8, phenyl, SCX, SAX etc.

Size, charge, partition, ion-exchange

PEC

As above

Size, charge, partition, ion-exchange

*In electrodriven size exclusion chromatography elution time is influenced by the exclusion mechanism and for charged molecules by the electromobility.

3.3 UNMODIFIED PACKINGS

Many reviews have been written on the preparation, physico-chemical properties and application of silica in modern separation science [5-7]. RP LC with silica based bonded stationary phases is utilised for the majority of LC separations in laboratories world-wide. Their ubiquity derives from their versatility, in that generally a wide range of both ionic and non-ionic analyte species can be separated with these columns by careful selection of the stationary phase and mobile phase properties. The use of unbonded silica has tended to be limited to techniques such as adsorption chromatography and supercritical fluid chromatography (SFC) [8,9] which are important techniques, but by no means as widespread as their dominant reversedphase LC cousin. A number of reasons can be suggested for this including their biological incompatibility, the use of specialist equipment (in SFC) and the use of

Referencespp. 106-110

Chapter 3

90

toxic non-aqueous solvents (which is actively discouraged in many laboratories from an environmental and disposal cost perspective). The use of unbonded silica with aqueous-organic eluents was reported many years ago in LC, where it was discovered that good peak shape for strong bases could be achieved [ 10]. A number of publications then followed looking at other analytes and included examinations of the separation mechanism [ 10,11 ]. Interest in this mode of LC was short-lived however, as workers realised that using inorganic buffer salts such as phosphates with high aqueous conditions led to severely reduced column life due to the silica dissolution. Recent work at AstraZeneca R&D Chamwood has circumvented this issue by using amine base buffers in conjunction with a high proportion of organic solvent in the mobile phase [12]. As CEC is a relatively new technique there are few reports using unbonded silica as a stationary phase. Reports to date describe the use of unbonded silica with various conditions and additives including non-aqueous solvents [ 13], dynamic coatings using cationic surfactants [14], blending with ODS phases [15] and cyclodextrin selectors [ 16]. Perhaps more interesting from a pharmaceutical perspective are the few preliminary publications detailing separations using various unbonded silicas for strongly basic [ 17-20] and weakly basic analytes [21 ]. It is of interest to note that certain types of silica exhibited the anomalous peak focusing in a similar fashion to the strong cation exchange materials described by Smith & Evans [22]. In recent work within our laboratories, we have evaluated a number of unbonded silicas by CEC specifically for the separation of a range of pharmaceutically relevant basic analytes and mixtures. We purposefully chose strong basic analytes that comprised a wide range of lipophilicities, molecular weights and log P values to robustly test the separation systems. The basic analyte test mixture contained two AstraZeneca R&D compounds, benzylamine, nortriptyline, diphenhydramine and procainamide. The following section describes initial work from an on-going research program to fully appreciate the role that the silica surface, its components and bonding chemistries play in contributing to CEC separations for various analytes. Packing methodology was based on a slurry procedure similar to that described previously [23] and found to be very repeatable once optimised for all phases studied. Initial testing of the capillaries was based on direct comparison of chromatographic parameters such as the magnitude of the electrosmotic flow (EOF) and the selectivity between two components. Figure 3.1 shows four unbonded silicas tested using a test mixture comprising biphenyl, benzamide, benzyl alcohol and thiourea. It was quickly established that thiourea was retained on all unbonded phases, and could not be used as the EOF marker. A number of other analytes was tried, and biphenyl was selected. It can be seen from Figure 3.2 that the purer silicas (HyPURITY and Kromasil) give a lower EOF than the more acidic traditional Hypersil

Modes of CEC

91

mAU 15 10

12 8 4 0

(A)

2

4

6

min

(B) 12 2

4

6

4

8

0 min

(C)

3 12 .

4

2

4

6

.

.

.

.

.

.

.

.

.

10

8

(D)

1 2~

2

4

6

8

12 min

~3

10

min

Fig. 3.1. Separation of the unbonded silica test mixture using in-house packed (A) Hypersil silica, (B) Hypersil BDS silica, (C) Hypersil HyPURITY and (D) Kromasil silica capillaries (100 l.tm i.d., 33.5 cm total length and 25 cm effective). Conditions: 8:2 v/v ACN-50 mM MES, pH 6.1, 20 kV, 20~ 5 kV for 3 sec. injections, 254 nm. Peak identities: 1 - biphenyl, 2 = benzamide, 3 - benzyl alcohol and 4 = thiourea. Adaptation of [20]. Reproduced with the permission of Chromatographia.

silica. These results are explained by the presence of fewer metal cations in the silica phases which will reduce the proportion of acidic silanols, leading to a reduced EOF generating ability. The better peak resolution of the neutral test mixture on the Kromasil material was attributed to its known greater surface area properties and its slightly larger particle diameter (all Hypersil phases dp= 3 lam and surface area of 170 m2/g whereas Kromasil dp= 3.5 lain and surface area of 340 m2/g). Efficiency values could not be compared for these phases, as the sample loading was different due to the use of electrokinetic injection. The traditional Hypersil unbonded silica at pH 7.8 gave baseline separation within 14 minutes of the basic analyte mixture (Fig. 3.2). The poorer chromatographic performance of the procainamide analyte was attributed to its suspected metal chelating properties. The purer HyPURITY unbonded silica gave broader peak shapes with reduced analysis times than the traditional acidic silica, but complete separation of the analytes was not observed. The separation of the bases on Hypersil BDS was observed

References pp. 106-110

Chapter 3

92 mAU 60-

(A)

II

40

IV I ll

0

2

4

VI

6

30

8

10

12

min

II (B)

20 I

IV V

10 VI 2 40

4

S ......

i0

:mi.

II

(C) 30

l/1V

20

1

10

0

2

4

6

8

min

Fig. 3.2. CEC separation of the basic test mixture using (A) Hypersil silica, (B) Hypersil BDS silica and (C) HyPURITY silica capillaries (100 ~tm i.d., 25 cm effective length, 33.5 cm total). Conditions: 6:2:2 v/v/v ACN-H20-50 mM TRIS, pH 7.8, 20 kV, 20~ 5 kV/3 s injections, 210 rim. Injection mixtures in (A) and (C) were equivolume compositions of each base at 1 mg/ml and in (B), equivolume compositions of each base at 0.1 mg/ml. The EOF marked by biphenyl under these conditions and was similar on all phases at approximately 4 minutes. Peak identities: I = AZ compound A, II - Benzylamine, l l l - Nortriptyline, IV Diphenhydramine, V - AZ compound B and VI = Procainamide. Adaptation of [20]. Reproduced with the permission of Chromatographia.

to fall between these two extremes. It is interesting to note that the EOF is similar on all the unbonded phases at pH 7.8, which is not what was observed at pH 6.1. This observation provides that the non-acidic silanol groups are intrinsically involved in EOF generation, which has not been widely appreciated previously.

93

Modes o f C E C

mAU 20 III 10

1 4 ........

S. . . . . . . .

12 . . . .

min

Fig. 3.3. Separation of neutral, acidic and basic components in their ionised form with Hypersil unbonded BDS silica. Peak identities: I- Benzylamine, II- Caffeine and III = p-hydroxybenzoic acid. The arrow denotes the EOF. Conditions = 6:2:2 v/v/v ACN:H20:50 mM MES, pH 6.1, 20 kV, 20~ 214 nm, 8 bar for 15 sec. inj. Adaptation of [20]. Reproduced with the permission of Chromatographia.

The separation mechanism with unbonded silica is extremely complex and attempts to discuss the details have been reported [19,20]. It is postulated that the separation mechanism with unbonded silica is a balance between the analyte's electromobility and an interaction component (comprising of predominantly ion-exchange, adsorption and, surprisingly, a RP character) with the unbonded silica. The ion-exchange properties of the silica are critical as phase purity is thought to dictate which of these factors dominates over the others. For example, with HyPURITY silica, there are reduced interactions with the silica surface due to a lower ion-exchange capacity, so the analyte's electromobility dominates the separation leading to shorter retention times. With Hypersil silica, the ion-exchange interactions dominate, leading to increased retention and different selectivity. A purpose designed CEC stationary phase that gives excellent EOF character yet allows rapid simultaneous acidic, basic and neutral separations under isocratic conditions without tailing is yet to be discovered. Nevertheless, the Hypersil BDS unbonded silica used in this work was taken forward to explore such complex mixture separations as it combines features of both pure and traditional media. As a phase, it possesses a reasonable EOF (as it is based on traditional silica) with a lower number of activated silanol groups on the silica surface (due to the pre-treatment procedure used to remove surface metal contamination). Figure 3.3 shows the separation of benzylamine, caffeine and p-hydroxybenzoic acid in a single chromatographic analysis.

References pp. 106-110

Chapter 3

94

It is important to note that the acidic and basic species are analysed in their ionic form under these conditions and complete separation of the mixture components is observed. Peak shape is seen to be good for all components, with the basic analyte eluting quickly due to its positive electromobility influence. The neutral species elute just after the EOF, whilst the acid is seen to elute slightly later due to its negative electromobility. It is clear that the unbonded Hypersil BDS may be additionally used for the separation of complex mixtures. 3.4 MODIFIED PACKINGS

The packing materials used in CEC are generally derived from LC technology and over 95% of CEC separations and publications are completed using silica based C18 bonded phases [24]. It is thought that the dominance of the C18 bonded phase for separations in CEC is derived from the majority of workers requiring determinations involving non-polar and weakly ionic analytes only. Nevertheless, work using other bonded phases such as C8 and phenyl [25,26], chiral phases [27,28] anion and cationexchange phases [22,29,30] and mixed mode [31 ] media [32] have been published. The hydrophobic selectivity of the stationary phase can be increased in an analogous way to RPLC, by covalently bonding hydrophobic moieties to a silica support material. In CEC this bonding has an influence on the EOF [33]. The total number of free silanol groups is reduced hence reducing the zeta potential and reducing the EOF. The same effects are found when end-capping is performed. The greater the coverage the more the EOF is reduced. Practically, this could make migration times too long for non-ionic species whose elution depends on the EOF alone. The pH of the mobile phase also has a profound influence on the EOF as the pH dictates the number of silanol groups that are ionised. This has been addressed in a number of ways, which are discussed later. A further practical consideration for CEC is bubble formation. This can result from Joule heating in the column or decompression on passing from the packed to open capillary. In addition to bubbles causing spikes as they pass through the detector, there is the danger of a break in the electrical circuit. Once formed, such bubbles are difficult to remove without applying a high extemal pressure to flush the column. Additionally, as the flits that retain the packing material are often made in-column by sintering the packing material, there is a danger of removing the bonded ligand during the hydrothermal process of preparing the frit which may generate undesirable surface active sites. Joule heating is minimised by using low ionic strength buffers, low voltages, organic buffers, low column temperatures and narrow intemal diameters. Decompression is avoided by low voltages or applying an extemal pressure to the capillary. It is

Modes of CEC

95

the latter approach, which is adopted by purpose made instruments as this, allows the use of high voltages with the concomitant rapid analysis times. However CEC may be performed using standard CE equipment by the use of low temperature and limiting voltages to <15kV [34,35]. A further approach to minimise bubble formation and hence allow unpressurised CEC to be performed, has been to use narrow and resilyated outlet frits [36] or incorporate sub-micellar concentrations of sodium dodecyl sulphate into the mobile phase [37,38]. Typically, packing materials designed for LC have been used for CEC [39]. However, there are significant differences in the requirements for packing materials for CEC and LC. Firstly, the EOF does not suffer from the significant increase in back pressures associated with using smaller particles in LC, smaller particles (0.2-3 lam) [40,41 ] and longer columns may be used in CEC and therefore highly efficient separations are possible. Secondly, the requirement for mechanical strength is lessened with electrodriven separations compared with pressure-driven systems. Finally, only extremely small quantities of packing material are required for a CEC column hence packings that would not be economically viable for LC become realistic options for CEC.

3.4.1 Reversed-phase (i.e. C8, C18, phenyl) In the RP CEC of neutral species selectivity is provided primarily by differences in the partition of the analytes between the hydrophobic stationary phase and the more polar mobile phase. There are also contributions from interactions with the silica support, the major one being polar interactions with ionised silanol groups. This is identical to the process in LC, albeit with the advantages of higher efficiencies in CEC resulting from the plug-flow profile. Additional selectivity is introduced in the case of charged species in CEC due to differences in the analytes' electromobilities. Standard C18 LC stationary phases packed into fused silica capillaries were the first phases to be extensively used for CEC [42-47]. Both porous and non-porous particles have been employed in CEC [38,41 ]. These stationary phases have been used in conjunction with familiar buffered aqueous RP mobile phases with organic modifiers such as methanol, acetonitrile and tetrahydrofuran [48]. The relationship between log retention factor (k) and percentage organic for uncharged species has been shown to be linear with eluents containing methanol and acetonitrile [25,26,33,46]. This allows rapid method development in an analogous way to that used in LC. It is important to note that, in addition to the influence on selectivity, changing the percentage or type of modifier has a profound impact on the EOF as a result of changes in the dielectric constant : viscosity ratio [49]. Hence, whilst mobile phases may be isoelutropic the actual retention times, even for neutral species, will differ significantly due to changes in flow rate induced by changes in viscosity (see Fig. 3.4). References pp. 106-110

Chapter 3

96

23

mAU ~ 40

1

,o

4

6

20

_

mAU ]

2

5

4

10

1

15

_

8

6 3

20

10

12

14

min

2

25

30

35

min

Fig. 3.4. Effect of mobile phase selectivity on the CEC separation of barbiturates (1-6). Electrochromatography was performed at 15~ with an applied voltage of 30 kV on a 25 cm, 100 ~tm i.d., 3 ~tm Hypersil Phenyl packed capillary. Sample concentration was 170 ~tg ml 1 of each component with a 15 kV/5s injection. Detection was at 210 nm. a) ACN-50 mM phosphate buffer, pH 4.5-water (4:2:4 v/v/v), b) MeOH-50 mM phosphate buffer, pH 4.5-water (5:2:3 v/v/v). From Euerby et al [26], 9 of Microcolumn Separations, 1999. Reproduced with permission of John Wiley & Sons, Inc.

Typically, acetonitrile yields higher efficiencies and more rapid elution than methanol due to acetonitrile's higher dielectric constant and lower viscosity. The pH of the mobile phase will have a profound effect on acidic and basic analytes and upon the number of ionised silanol groups on the silica surface and hence the EOF. Buffers must be used to ensure reproducible pH conditions. The type and concentration of buffer in the mobile phase is more limited in CEC than in LC as high inorganic buffer concentrations lead to excessive current and hence Joule heating as in CE. Therefore inorganic buffers such as phosphates and borates are used at low concentration (<50 mM) or organic buffers commonly used in CE such as tris(hydroxymethyl)aminomethane (Tris), and 2-(N-Morpholino)ethane-sulfonic acid (MES) are employed. The ionic strength of the mobile phase is also an important factor as increasing the ionic strength of the mobile phase reduces the zeta potential, which results in a decreased EOF and hence increased retention times. The classical stationary phases used for LC possess relatively high levels of metal ions in the silica support material. These give rise to a large number of highly acidic silanol groups. After bonding of the stationary phase the remaining isolated silanols prove troublesome for the analysis of bases. In recent years, a new generation of pure silicas have been manufactured via highly controlled, improved processes. These phases possess lower metal content and hence the acidity of any residual silanol

97

Modes of CEC

mAU

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3

2

.

,.

20 10 0

201

1~

3~

2

4 6

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r

1

4

5

6

7

8

9

10

11

min

Fig. 3.5. Chromatograms of the optimised separation of the barbiturates (1-6) on three different packing materials. Electrochromatography was performed with an applied voltage of 30 kV on a 25 cm, 100 ktm i.d., 3 ~tm. Sample concentration was 170 ~g ml"1 of each component with a 15 kV/5s injection. Detection was at 210 nm. (a) Hypersil C8, ACN-50 mM MES, pH 6.1-water (5:2:3 v/v/v), 15~ (b) CEC Hypersil, ACN-50 mM MES, pH 6.1-water (4:2:4 v/v/v), 15~ (c) Hypersil Phenyl, MeOH-50 mM phosphate buffer, pH 4.5-water (5:2:3 v/v/v), 60~ * corresponds to an artifact in the mobile phase. From Euerby et al [26], 9 of Microcolumn Separations, 1999. Reproduced with permission of John Wiley & Sons, Inc.

groups is greatly reduced, in turn, improving basic analyte separations in LC [50]. Unfortunately, it is the presence of ionised silanols which generate the EOF essential for the elution of neutral species in CEC, therefore the practical working pH range in CEC is reduced with the new generation silica based packing materials. In addition to generating excessively long retention times with these pure phases at low pH, there is the practical problem of the phase drying out resulting in breakdown of the current. It is for this reason that stationary phases are being developed specifically for CEC. Robust and reproducible methods have been developed with traditional RP materials for neutral and ion suppressed acidic analytes [51,52], in application to pharmaceutical analysis [34,53,54], aromatic compounds [55], phenols in tobacco smoke [56], preservatives in creams [40,41] nucleosides [57,58] and cannabinoids [59]. Typical efficiencies were >100,000 plates/m. The analysis of bases, however, remains a challenge (see later section). The use of other column chemistries such as C8 and phenyl phases allows the selectivity of the stationary phase to be optimised in a similar fashion to that used in LC [60-62], (see Fig. 3.5).

Referencespp. 106-110

Chapter 3

98 3.4.2 The use of mobile phase additives

As the generation of a significant EOF requires the presence of ionised groups on the surface of the stationary phase, classical silicas with high metal contents have been widely used in CEC. Before "new generation" silica RP column chemistries became available, undesirable secondary interactions of basic analytes in LC were observed. The traditional approach to the analysis of bases by LC using stationary phases containing acidic silanols was to use small competing bases added to the mobile phase, such as triethy!amine or, in CE, triethanolamine has been used. Recent work in this area has demonstrated that strong and weak acids and bases, and neutrals can all be eluted in one CEC run by the use of triethylamine phosphate or triethanolamine phosphate [63] (see Fig. 3.6) or hexylamine at pH 2.5 [64,65]. By the use of this approach acids are chromatographed in their ion-suppressed mode whilst strong and weak bases are positively charged. The uncharged species are separated according to differences in lipophilicity by interactions with the RP stationary phase giving rise to significantly different selectivity to that in CE. The selectivity also differs from LC for the charged bases due to additional differences in electrophoretic mobility. An added benefit is the ability to distinguish bases in mixtures of acids, bases and neutrals as the positively charged species elute before the EOF at pH 2.5. The orthogonal separation mechanism provided by this approach is extremely attractive to the pharmaceutical industry. The inclusion of basic additives in the run buffer leads to a reduction in the EOF. This is due to the reduction in the number of free silanol sites on the silica surface. However, above 50 mM the continued reduction in the EOF is less pronounced [63]. In practice, sufficient EOF is generated, even in the presence of mobile phase additives, to elute neutral species in acceptable times. The upper limit on the additive concentration is most frequently due to excessive baseline noise arising from high background absorbance. The inclusion of mobile phase additives leads to a further level of complexity in method development and prohibits coupling to mass spectrometry. However, this approach is a practical solution until better stationary phases are developed. 3.4.3 Ion exchange - SAX, SCX and Mixed Mode

3.4.3.1 SCX In order to extend the practical working pH range in CEC the use of stationary phases containing charged functional groups has been utilised. A strong cation-exchange phase (SCX, which contains a sulphonic acid group) will be negatively charged from pH 2 to pH 9. Hence, a good EOF is maintained throughout this region

99

Modes o f CEC

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Fig. 3.6. Electrochromatogram of benzylamine (I), caffeine (II) and benzoic acid (III). Efficiency values of 4642, 76331, and 6399 plates per column were obtained respectively for I, II, and nI. Electrochromatography was performed at 15~ with an applied voltage of 25 kV on a 25 cm, 100 ~tm i.d., 3 ~tm Hypersil Phenyl packed capillary. Mobile phase: ACN-50 mM triethanolamine phosphate, pH 2.5-H20 (6:2:2 v/v/v). Sample concentration was 100 ~tg ml 1 of each component, 5 kV/5s injection. Detection at 214 nm. From [63]. Reproduced with permission of The Royal Society of Chemistry

[22]. Considerable excitement was generated when Smith and Evans reported efficiencies in excess of 8 x 106 plates m 1 for the analysis of the basic tricyclic antidepressants. These high efficiencies have also been obtained by other workers for a range of structurally diverse basic compounds. Unfortunately these workers, including Smith and Evans, have experienced severe non-reproducibility of the phase, in that severe tailing and fronting have been unexpectedly observed in the middle of successful runs [46,66,67]. Explorations into the focussing mechanism have been reported [68], but it is yet to be fully characterised and understood. However, the separation of weakly basic aromatics on this phase has been demonstrated [21]. An additional disadvantage of the SCX phase is that it possesses little hydrophobic character and hence is not very selective for the separation of neutral analytes. Harnessing the phenomenon of high peak efficiencies in a repeatable fashion could lead to a new surge in the interest for CEC. 3.4.3.2 SAX

In the case of a strong anion-exchange phase (SAX, which typically contains a quaternary amine), EOF reversal is observed, and an ion-exchange mechanism with negatively charged species will also result [69]. This approach has been successfully applied to the analysis of iodide and iodate [70], and has recently been reported for the References pp. 106-110

1O0

Chapter 3

separation of a range of large proteins using a custom synthesised tentacular anion exchanger [71]. The resolving power of CEC with this phase for protein work was demonstrated with separations of chicken egg-white conalbumin variants in a single four-minute run with isocratic eluents. Other workers have reported the use of anion exchanger columns for lanthanide and inorganic ion analysis [29] and aqueous and non-aqueous chiral derivatised amino acid analysis [72,73]. 3.4. 3.3 Mixed mode

There are two types of mixed mode phases 1) physically distinct particles possessing separate lipophilic and ion exchange chemistries [74] and 2) the lipophilic and ion exchange chemistries on the same silica particle [26,75]. The use of mixed mode phases (which contain charged functional groups and lipophilic spacers or separate alkyl ligands) has yielded rapid, efficient methods for the analysis of neutral and ion suppressed acids. Custom synthesised multilayer mixed-mode phases containing ligands with a sulphonic acid sub-layer and a C18 top-layer have also been reported [32,58]. These phases have been successfully used to separate nucleobases and even strongly acidic analytes such as nucleic acids of various sizes from dinucleotides to t-RNAs. Excellent peak area, and migration time precision can be obtained for neutral and acidic compounds for a given mixed-mode column. A major advantage of using mixed mode phases is that they allow the rapid analysis of ion suppressed acidic and neutral analytes at low pH [26]. With the exception of one recent report by Rozing [76], using a new Zorbax SCX/C18, no success has been reported in analysing basic compounds with SCX mixed mode phases. 3.5 CHIRAL STATIONARY PHASES Chiral CEC will be discussed in detail later in the book but is included here to exemplify the application of the high efficiencies obtained with electro-driven techniques which makes them attractive for chiral analysis where selectivity factors are sometimes small. CE has made use of chiral additives in the electrolyte whilst LC tends to utilise chiral stationary phases. Both options have been explored for chiral CEC [27,28,77]. The small amount of packing material necessary for capillaries allows the use of chiral stationary phases that would be prohibitively expensive for standard LC. Cyclodextrins, proteins, antibiotics and molecular imprinting have all been used to form chiral stationary phases [78-80]. After some less than encouraging peak efficiencies obtained using the chiral CEC approach, much improved chiral resolutions have been achieved using CEC compared to LC or CE [81-83].

Modes o f CEC

101

3.6 GEL CEC

Columns packed (filled) with natural and synthetic polymers such as acrylamides, dextrans, glycols, poly(ethylene oxides), poly(ethylene glycols), methacrylates, agaroses and various cellulose derivatives have been widely applied and reviewed for a number of applications in capillary gel electrophoresis [84,85]. The rationale for such a diverse range of polymers is associated with their differences in physicochemical properties which allows separation of a variety of molecules and biopolymers over a wide range of polymer concentrations [86]. Gel CEC is included here simply because many publications describe covalently immobilised polymers or mimetic gels within a fused silica capillary as "electrochromatography" and a distinction between Gel CEC and CGE is therefore difficult to define. Nevertheless, Gel CEC will be covered in greater depth in a later chapter, and it is included here simply because, due to the lack of a clear definition, it could be argued to be the most widely used mode of CEC. 3.7 MONOLITHS The use of hydrothermally formed retaining frits in capillary columns packed with stationary phase particles is an accepted limitation in CEC. The introduction of the frit to hold the packed bed is vital, yet introduces problems such as EOF and flow non-uniformities, compromised frit permeability [87], capillary fragility, increased likelihood of bubble formation [88] and a thermally induced modified frit surface chemistry which can detrimentally alter the chromatography [23]. Practical aspects to be considered include the appreciable effort and skill of the analyst who is required to repeatably manufacture capillaries of a particular phase and redevelop the fritting and packing methodology for each different stationary phase type. The use of continuous bed columns (i.e. monoliths) can address many of these points. Early monolith research focussed on the polymerisation of acrylamides analogous to the approach utilised in CGE [89]. As the area developed, workers tried different chemistries but experienced problems that ranged from shrinkage of the monolith from the capillary wall and / or cracking of the monolith structure - leading to poor chromatography. It is these issues that have generally hampered the development of monolith columns for LC as well as CEC. A monolithic column was recently defined as "A continuous unitary porous structure prepared by in situ polymerisation or consolidation inside the column tubing and, if necessary, the surface is functionalised to convert it into a sorbent with the desired chromatographic binding properties" [90]. Whilst quite general, this definition actually covers a range of methods that can be used to produce continuous bed columns. References pp. 106-110

102

Chapter 3

The many reported methodologies for producing monolithic columns published to date differ in their manufacturing process and the end-product monolith. Porous moulded organic polymers are perhaps the most reported in CEC and involve in-situ polymerisation of monomer solutions in the presence of a porogen using various initiator techniques such as free radical polymerisation [91] or photopolymerisation [92]. By carefully controlling the polymerisation components and conditions for this methodology, monoliths of defined physicochemical properties may be produced which are robust and exhibit excellent chromatographic properties [93]. Particle-fixed continuous beds are another example involving the immobilisation of stationary phase particles of known characteristics using whole column sintering [94]. Finally, sol-gel technology involving the hydrolysis and polycondensation of precursors in a defined solvent in-situ to produce a hydrogel, which may then be converted to a xerogel upon drying before functionalising has been reported [95]. A derivation on this theme is the use of particle-loaded sol-gels where polycondensation of a mixture of polyalkoxysiloxanes and the stationary phase to produce end frits to effectively retain the phase [96] or to completely immobilise the column have been demonstrated [97,98]. Thus, the monolith approach is highly customisable for a particular need or application. It is an important area in column technology development with the number of publications in CEC, ~tLC and even monolith based standard LC increasing dramatically. Details and specific application examples of the use of these various monolith technologies in CEC will be discussed in a later chapter. 3.8 SIZE EXCLUSION CEC In size exclusion chromatography, selectivity for neutral molecules is based on molecular size and shape. The stationary phase consists of either a polymeric gel or a silica gel with controlled pore size. Larger molecules are excluded from the pores whilst smaller molecules, can enter the pores and are hence eluted later. In CEC of charged molecules additional selectivity is introduced based on electromobility. The mobile phase is used to change the molecular shape and/or charge and to optimise secondary interactions. Capillaries packed with unmodified silica gel with pore sizes between 100 and 10 nm have been evaluated for CEC [99]. In addition to the practical application to polymer analysis, the technique is of theoretical interest as a demonstration of the appreciable intra-particle flow in CEC with associated efficiency gains, which is absent in LC. Li and Remcho [ 100] studied packing media pore sizes from 6 to 400 nm and deduced that only the larger pore diameters (>200 nm) supported pore flow, which led to higher peak efficiencies. For SEC a low pore flow is required to obtain selectivity based on exclusion/inclusion in the pore. Hence low ionic strengths must be used to induce double layer overlap in the pores. The optimisation of the

Modes of CEC

103

pore-to-interstitial flow for size-exclusion has been investigated by Stol et al [101 ]. The implications to RP CEC have also been explored [99,102] and the findings indicate that higher efficiencies and linear flow rates are obtained with larger pore sizes (e.g. 400 nm) and higher buffer strengths which favour high intra-particle flow. In further work, Stol et al [103] verified this work and demonstrated substantial intraparticulate flow with much smaller unbonded packing media (30 nm) in addition to the 400 nm material. The high efficiencies observed for the neutral test probes were attributed to better electrokinetic flow homogeneity due to the large pore size and enhancement of the mass transfer kinetics. A rigorous theoretical treatment of flow characteristics in packed beds and determination of an ideal packed capillary structure for electrokinetic flow was offered by Luo and Andrade [104]. 3.9 GRADIENT CEC AND VOLTAGE ASSISTED CAPILLARY LC Preliminary work has been published by a number of workers using research instrumentation [ 105-108]. There are, at time of publication, few commercially available continuous gradient CEC instruments. It is possible to perform simple step gradients using standard CEC instruments by changing the mobile phase during the run [109,110]. This will be a critical area for the development and acceptance of the technique. Preliminary reports using a prototype gradient CEC system capable of performing capillary LC with voltage assisted flow have been very encouraging [25]. It has been found that a large number of compounds of widely different lipophilicity may be eluted and resolved using isocratic CEC that would have required gradients for LC. This presumably is due to the high peak capacity of CEC and hence higher efficiencies compared to LC. The combination of pressure-driven and electro-driven flow offers a great potential for optimisation of separations. For example, CEC systems using columns with low EOF may not elute all species by electromobility alone, whereas the application of pressure will ensure a sufficient flow to elute all species [76,111 ]. The selectivity and structural information afforded by being able to elute analytes with and without applied voltage is also of practical benefit [47]. Once again the availability of commercial instrumentation will be the key to the success of this technique. 3.10 SELECTIVITY COMPARED WITH LC The potential influence of the nature of the driving force on the chromatographic selectivity for neutral molecules has been investigated using CEC, gLC and pressurised flow electrochromatography. The comparison was performed on the same prototype instrument and the same column to eliminate all other possible influences References pp. 106-110

Chapter 3

104

on the selectivity. The relationship between k and the percentage of organic modifier present in the mobile phase was used to compare the consequence of the different mechanisms on the separation of non-ionised molecules. The slopes and intercepts of the plots of log k against percentage organic in the mobile phase, obtained by using the different flow generation mechanisms, were statistically evaluated. As expected, a linear relationship was found between log k and the percentage of organic modifier for a series of weak acids and bases analysed in their ion-suppressed mode and for neutral compounds. Under the conditions investigated, the chromatographic selectivity in electro, pressure driven flow and a combination of thereof was shown to be equivalent for the non-ionised molecules studied [25]. Similar findings were reported by another group in that no significant differences could be demonstrated between CEC, PEC and ~tLC modes on the same capillary for separating a range of 27 neutral analytes [ 112]. In contrast other groups have reported the observation that the chromatographic characteristics of porous reverse phase materials depends on the mode of flow generation [113]. Other authors have additionally reported the different elution profiles between PEC and capillary LC for analytes that include basic amines [47]. On a practical basis, numerous workers such as Ross et al [114] have shown that method transfer between LC and CEC for neutral or ion suppressed analytes is straightforward and simple. The obvious benefits of using CEC compared to LC include increased efficiencies and hence enhanced peak capacity and the orthogonal nature of CEC compared to LC when ionic analytes are present. Retention modelling of neutral [ 115] and basic analytes [12,65] taking into account retention factors and electromobilities of the analytes have shown good agreement between predicted and experiment retention times. These results highlight the predictive nature of CEC and the possibility of performing computer optimisation routines. The possibility of using voltage to "fine tune" pressure separations is an attractive technique which will require more attention when commercially available instrumentation becomes available. 3.11

GUIDELINES FOR THE ANALYSIS OF ACIDIC BASIC AND NEUTRAL COMPOUNDS

3.11.1 Neutral analytes

Traditional LC stationary phase, such as Hypersil or Spherisorb ODS1 materials are ideally suited to the analysis of neutral analytes as the phases possess a high content of acidic silanols, hence at pH 7-9 high EOF generation is achieved which facilitates rapid analysis. In marked contrast, many pharmaceutical compounds pos-

Modes of CEC

105

sess an ionizable functionality and hence different approaches must be used for their analysis as discussed in the sections below.

3.11.2 Acidic analytes Ionised acids tend to migrate towards the anode counter to the EOF therefore they are either not loaded onto the column during electrokinetic injection or are not swept towards the detector and hence are not detected. In order to perform CEC of acidic analytes they must be run in their non ionised form i.e. in acidic mobile phase. As a consequence of the low mobile phase pH there is a significantly reduced EOF hence long retention times are observed. It is highly recommended that the use of mixed mode phases such as the SAX/C18 and SCX/C6 are employed when using low pH mobile phases in order to enhance the EOF [ 116].

3.11.3 Basic analytes The analysis of basic analytes on stationary phases with silica support is still problematic in LC due to the possibility of the basic analyte undergoing mixed mode interactions with the stationary phase i.e. hydrophobic and ionic interactions. Residual isolated silanols are responsible for the deleterious ionic interactions, the result of which is excessive peak tailing. The CEC analysis of basic analytes is thought to be problematic because in order to generate a good EOF, an acidic silica is essential. It is these silanols groups which generate the EOF and are responsible for unwanted peak tailing. A number of approaches to improving the peak shape of basic analytes on silica supports for CEC with stationary phase design have been taken from LC. Many of the problems encountered in the analysis of bases by LC are also manifest in CEC. The use of low pHs or reducing the number of acidic silanols leads to extremely low EOFs which may cause excessive retention of concomitantly chromatographed neutral species in addition to the practical consideration of maintaining a "wetted" capillary. These factors have led to a perception that the analysis of bases by CEC is difficult, if not practically impossible. A number of "new" generation silica support materials have also been evaluated. The reduction in EOF is significant leading to excessive retention time for neutral and acidic species. However, pharmaceutical bases can generally be analysed using such phases, as the intrinsic electromobilities of the molecules are sufficient to ensure elution. Unfortunately, there has been little success to date. The best approach to date for the analysis of basic compounds by CEC has been to incorporate a small basic compound such as triethylamine, triethanolamine [63] or hexylamine [64] into the low pH mobile phase. The small bases act in a competitive manner to restrict the access of the basic analytes to the silanol groups on the surface References pp. 106-110

Chapter 3

106

of the silica. This approach has been shown to work for a number of basic analytes [63-65]. The same approach also allows the simultaneous analysis of acids, bases and neutral, the only drawback being the relative slow EOF. 3.12 CONCLUSIONS The enhanced sample loadability together with the high efficiencies obtainable in CEC have stimulated much interest in this technique. The entire ranges of LC and CE modes are potentially available within CEC. Many of these are only in the initial investigation stages. With the advent of stationary phases specifically designed for CEC and a growing theoretical understanding of the mechanisms involved in CEC, the continued development of the technique is assured. Nevertheless, support from stationary phase manufacturers for custom designed CEC phases and a robust column format are critical for its continued development and acceptance as a mainstream technique. 3.13 ABBREVIATIONS

CEC CGE EOF MES PEC RP SAX SCX SFC TRIS

capillary electrochromatography capillary gel electrophoresis electrosmotic flow 2-(N-morpholino)ethane-sulfonic acid pressure assisted electrochromatography reversed-phase strong anion exchange strong cation exchange supercritical fluid chromatography tris(hydroxymethyl)aminomethane

3.14 REFERENCES

1 2 3 4 5 6

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10 B.A. Bidlingmeyer, J.K. Del-Rios and J. Korpi, Anal. Chem., 54 (1982) 442. 11 G.B. Cox and R.W. Stout, J. Chromatogr., 384 (1987) 315. 12 A.P McKeown presented at the Chromatographic Society Spring Meeting, Loughborough, June 2000. 13 A. Maru~ka and U. Pyell, J. Chromatogr. A, 782 (1997) 167. 14 M. Ye, H. Zou, Z. Liu, J. Ni and Y. Zhang, J. Chromatogr. A, 855 (1999) 137. 15 N.M. Djordjevic, F. Fitzpatrick, F. Houdiere and G. Lerch, J. High Resol. Chromatogr., 22 (1999) 599. 16 W. Wei, G.A. Luo and R. Xiang, J. Microcol. Sep., 11 (1999) 263. 17 M.M. Dittmann presented at the 20th International Symposium on Capillary Chromatography, Riva del Garda, Italy, May 1998. 18 P.D.A. Angus, J.F. Stobaugh, C.W. Demarest K.M. Payne K R Sedo L.Y. Kwok and th ' ' " " ' T. Catalano presented, at the 19 International Symposium on Capillary Chromatography and Electrophoesis, Wintergreen, VA, May 1997. 19 W. Wei, G.A. Luo, G.Y. Hua and C. Yan, J. Chromatogr. A, 817 (1998) 65. 20 A.P McKeown, M.R. Euerby, H. Lomax, C.M. Johnson, P. Ross and H. Richie, Chromatographia, submitted, 2000. 21 C.W. Klampfl and P.R. Haddad, J. Chromatogr. A, 884 (2000) 277. 22 N.W. Smith and M.B. Evans, Chromatographia, 41 (1995) 197. 23 R.J. Boughtflower, T. Underwood and J. Maddin, Chromatographia, 41 (1995) 398. 24 K.D. Altria, J. Chromatogr. A, 856 (1999) 443. 25 M.R. Euerby, C. Chevet and C.M Johnson presented at 23 rd International Conference on Chromatography, Granada (1999). 26 M.R. Euerby, C.M. Johnson, S.F. Smyth, N. Gillott, D.A. Barrett and P.N. Shaw, J. Microcol. Sep., 11 (1999) 305. 27 S. Li and D.K. Lloyd, Anal. Chem., 65 (1993) 3684. 28 S. Nilsson, L. Schweitz and M. Petersson, Electrophoresis, 18 (1997) 884. 29 S. Kitagawa, A. Tsuji, H. Watanabe, M. Nakashima and T. Tsuda, J. Microcol. Sep., 9 (1997) 347. 30 M.G. Cikalo, K.D. Bartle and P. Myers, Anal. Chem., 71 (1999) 1820. 31 M.R. Euerby, C.M. Johnson and K.D. Bartle, LC-GC Int., 11 (1998) 39. 32 M.Q. Zhang and Z. E1Rassi, Electrophoresis, 20 (1999) 31. 33 M.M. Dittman and G.P. Rozing, J. Chromatogr. A, 744 (1996) 63. 34 M.R. Euerby, C.M. Johnson, P. Myers, K.D. Bartle and S.C.P. Roulin, Anal. Comm., 33 (1996) 403. 35 M.M. Robson, S. Roulin, S.M. Shariff, M.W. Raynor, K.D. Bartle, A.A. Clifford, P. Myers, M.R. Euerby and C.M. Johnson, Chromatographia, 43 (1996) 313. 36 R.A. Carney, M.M. Robson, K.D. Bartle and P. Myers, J. High Resol. Chromatogr., 22 (1999) 29. 37 R.M. Seifar, W.T. Kok, J.C. Kraak and H. Poppe, Chromatographia, 46 (1997) 131. 38 R.M. Seifar, J.C. Kraak, W.T. Kok and H. Poppe, J. Chromatogr. A, 808 (1998) 71. 39 C. Fujimoto, Trends in Analytical Chemistry, 18 (1999) 291. 40 T. Adam and M. Kramer, Chromatographia, 49(Suppl. I) (1999) $35. 41 T. Adam, S. L~idtke and K.K. Unger, Chromatographia, 49 (1999) $49. 42 T. Tsuda, Anal Chem., 59 (1987) 521. 43 T. Tsuda, Anal Chem., 60 (1988) 1677. 44 J.H. Knox and I.H. Grant, Chromatographia, 32 (1991) 317.

108

Chapter 3

45 N.W. Smith and M.B. Evans, Chromatographia, 38 (1994) 649. 46 M.M. Dittmann presented at the 18th International Symposium on Capillary Chromatography, Riva del Garda, Italy, May 1996. 47 T. Eimer, K.K. Unger and J. van der Greef, TRAC, 15 (1996) 463. 48 M.M. Dittman and G.P. Rozing, Biomed. Chromatogr., 12 (1998) 136. 49 P.B. Wright, A.S. Lister and J.G. Dorsey, Anal. Chem., 69 (1997) 3251. 50 J.J. Kirkland and R.M. McCormick, Chromatographia, 24 (1987) 58. 51 P. Coufal, H.A. Claesens and C.A. Cramers, J. Liq. Chromatogr., 17 (1993)3623. 52 M.T. Dulay, C. Yan, D.J. Rakestraw and R.N. Zare, J. Chromatogr. A, 725 (1996) 361. 53 P.D.A. Angus, C.W. Demarest, T. Catalano and J.F. Stobaugh, Electrophoresis, 20 (1999) 2349. 54 J. Reilly and M. Saeed, J. Chromatogr. A, 829 (1998) 175. 55 L.H. Zhang, W. Shi, H. Zou, J. Y. Ni and Y.K. Zhang, J. Liq. Chromatogr. Relat. Technol., 22 (1999) 2715. 56 M. Saeed, M. Depala, D.H. Craston and I.G.M. Anderson, Chromatographia, 49 (1999) 391. 57 T. Helboe and S.H. Hansen, J Chromatogr. A, 836 (1999) 315. 58 M.Q. Zhang, C. Yang and Z. E1Rassi, Anal. Chem., 71 (1999) 3277. 59 I.S. Lurie, R.P. Meyers and T.S. Conver, Anal. Chem., 70 (1998) 3255. 60 X. Cahours, P. Morin and M. Dreux, J. Chromatogr. A, 845 (1999) 203. 61 N.W. Smith, CAST, 8 (1999) 10. 62 P.D.A. Angus, E. Victorino, K.M. Payne, C.W. Demarest, T. Catalano and J.F. Stobaugh, Electrophoresis, 19 (1998) 2073. 63 N.C. Gillott, M.R. Euerby, C.M. Johnson, D.A. Barrett and P.N. Shaw, Anal. Commun., 35 (1998) 217. 64 I.S. Lurie, T.S. Conver and V.L. Ford, Anal. Chem., 70 (1998) 4563. 65 M.M. Dittman K. Masuch and G.P. Rozing, J. Chromatogr. A, 887 (2000) 209. 66 M.R. Euerby, D. Gilligan, C.M. Johnson, S.C.P. Roulin, P. Myers and K.D. Bartle, J. Microcol. Sep., 9 (1997) 373. 67 N.W. Smith presented at the 1st International Symposium on Capillary Electrochromatography, San Francisco, CA, August 1997. 68 P.D. Ferguson, N.W. Smith, F. Moffatt, S.A.C. Wren and K.P. Evans, Poster presentation (1999) HPLC 99, Granada, Spain. 69 R. Grtiner, B. Scherer, F. Steiner and H. Engelhardt, presented at the 20th International Symposium on Capillary Chromatography, Riva del Garda, Itlay, May 1998. 70 D.M. Li, H.H. Knobel and V.T. Remcho, J. Chromatogr. B, 695 (1997) 169. 71 J. Zhang, X. Huang, S. Zhang and C. Horwith, Anal. Chem., 72 (2000) 3022. 72 M. L~immerhoferand W.J. Lindner, J. Chromatogr. A, 829 (1998) 115. 73 E. Tobler, M. Lammerhofer and W. Lindner, J. Chromatogr. A, 875 (2000) 341. 74 L. Zhang, Y. Zhang, W. Shi and H. Zou, J. High Res. Chromatogr., 22 (1999) 666. 75 N.W. Smith and M.B. Evans, J. Chromatogr. A, 832 (1999) 41. 76 G.P. Rozing presented at the 13th International Symposium on High Performance Capillary Electrophoresis and Related Microscale Techniques, Saarbrfiken, Germany, February 2000. 77 F. Lelievre, C. Yan, R.N. Zare and P. Gareil, J. Chromatogr. A, 723 (1996) 145. 78 S. Li and D.K. Lloyd, J. Chromatogr. A, 666 (1994) 321. 79 D.K. Lloyd, S. Li and P. Ryan, J. Chromatogr. A, 694 (1995) 285.

Modes of CEC

109

80 J.M. Lin, T. Nakagama, K. Uchiyama and T. Hobo, J. Pharm. Biomed. Anal., 15 (1997) 1351. 81 A. Dermaux, F. Lynen and P. Sandra, J. High Resol. Chromatogr., 21 (1998) 375. 82 C. Wolf, P.L. Spence, W.H. Pirkle, E.M. Derrico, D.M. Cavender and G.P. Rozing, J. Chromatogr. A, 782 (1997) 175. 83 A.S. CarterFinch and N.W. Smith, J. Chromatogr. A, 848 (1999) 375. 84 B.L. Karger, F. Foret and J. Berka, Chapter 13: Capillary electrophoresis with polymer matrices in Methods in Enzymology, Vol 271: High resolution separation and analysis of biological macromolecules Part B Applications., B.L. Karger and W. S. Hancock (Editors) 271 (1996) 293. 85 P.G. Righetti and C. Gelfi, Forensic Science Intl., 92 (1998) 239. 86 A.E. Barron, D.D. Soane and H.W. Blanch, J. Chromatogr., 652 (1993) 3. 87 E.F. Hilder, C.W. Klampf, M. Macka, P.R. Haddad and P. Myers, Analyst, 125 (2000) 1. 88 M.G. Cikalo, K.D. Bartle, M.M. Robson, P. Myers and M.R. Euerby, Analyst, 123 (1998) 87R. 89 C. Fujimoto, J. Kino and H. Sawada, J. Chromatogr. A, 715 (1995) 107. 90 I. Gusev, X. Huang and C. Horvath, J. Chromatogr. A, 855 (1999) 273. 91 J. Liao, N. Chen, C. Ericson and S. Hjerten, Anal. Chem., 68 (1996) 3468. 92 J.R. Chen, M.T. Dulay, R.N. Zare, F. Svec and E. Peters, Anal. Chem., 72 (2000) 1224. 93 N. Ishizuka, H. Minakuchi, K. Nakanishi, N. Soga, H. Nagayama, K. Hosoya and N. Tanaka, Anal. Chem., 72 (2000) 1275. 94 R. Asiaie, X. Huang, D. Faman and C. Horvath, J. Chromatogr. A, 806 (1998) 251. 95 Q.L. Tang and M.L. Lee, J. High Resol. Chromatogr., 23 (2000) 73. 96 M. Schmid, F. Bauml, A.P. Kohne and T. Welsch, J. High Res. Chromatogr., 22 (1999) 438. 97 Q.L. Tang, B.M. Xin and M.L. Lee, J. Chromatogr. A, 837 (1999) 35. 98 C.K. Ratnayake, C.S. Oh and M.P. Henry, J. High Res. Chromatogr., 23 (2000) 81. 99 E. Venema, J.C. Kraak, H. Poppe and R. Tijssen, J. Chromatogr. A, 837 (1999) 3. 100 D.M. Li and V.T. Remcho, J. Microcol. Sep., 9 (1997) 389. 101 R. Stol, W.T. Kok and H. Poppe, presented at the 24th International Symposium on High Performance Liquid Phase Separations and Related Techniques, Seattle, Washington, USA, June 2000. 102 E. Venema, J.C. Kraak, H. Poppe and R. Tijssen presented at the 20th International symposium on capillary chromatography, Riva del Garda, Italy, May 1998. 103 R. Stol, W.T. Kok and H. Poppe, J. Chromatogr. A, 853 (1999) 45. 104 Q.L. Luo and J.D. Andrade, J. Microcol. Sep., 11 (1999) 682. 105 M.R. Taylor and P. Teal, J. Chromatogr. A, 768 (1997) 89. 106 R. Daddoo, C. Yan, D.S. Anex, D.J. Rakestraw and G.A. Hux, LC-GC Int., 10 (1997) 146. 107 C.G. Huber, C Choudhary and C Horvfith, Anal. Chem., 69 (1997) 4429. 108 A.H. Que, V. Kahle and M.V. Novotny, J. Microcol. Sep., 12 (2000) 1. 109 M.R. Euerby, D. Gilligan, C.M. Johnson and K.D. Battle, Analyst, 122 (1997) 1087. 110 J. Ding, J. Szelign, A. Dipple and P. Vouros, J. Chromatogr. A, 781 (1997) 327. 111 T. Eimer, K.K. Unger and T. Tsuda, Fresenius J. Anal. Chem., 352 (1995) 649. 112 Y.K. Zhang, W. Shi, L.H. Zhang and H.F. Zou, J. Chromatogr. A, 802 (1998) 59.

110

Chapter 3

113 J. Jiskra, H. A. Claessens, M. Byelik and C.A. Cramers, J. Chromatogr. A, 862 (1999) 121. 114 G. Ross, M.M. Dittmann, and G.P. Rozing, Publication No. 5965-9031E (1997) Agilent Walbronn, Germany. 115 J.P.C. Visser, H.A. Classens and P. Coufal, J. High Resol. Chromatogr., 18 (1995) 540. 116 M.R. Euerby presented at the 20th Intemational Symposium on Capillary Chromatography, Riva del Garda, Itlay, May 1998.

Chapter 4

Packed Bed Columns Luis A. C O L O N * , Todd D. M A L O N E Y and A d a m M. F E R M I E R t

Department of Chemistry, State University of New York at Buffalo, Natural Sciences Complex, Buffalo, NY 14260-3000, USA ?Present address." The R. W. Johnson Pharmaceutical Research Institute, Science and New Technology~Analytical Development, OMP Bld. B-236, 1000 Route 202, Raritan, NJ 08869, USA

CONTENTS

4.1 4.2

4.3

4.4 4.5 4.6 4.7 4.8

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Column fabrication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1 The column . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.2 Chromatographic material . . . . . . . . . . . . . . . . . . . . . 4.2.2.1 Ion-exchangers and mixed-mode phases . . . . . . . . 4.2.2.2 Submicron particulate materials . . . . . . . . . . . . 4.2.2.3 Highly porous particles . . . . . . . . . . . . . . . . . 4.2.3 Retaining frits . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.3.1 Silica-base frits . . . . . . . . . . . . . . . . . . . . . 4.2.3.2 Fritless packed beds . . . . . . . . . . . . . . . . . . . 4.2.4 Fabricating columns . . . . . . . . . . . . . . . . . . . . . . . . Packing methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.1 Pressure packing . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.2 Packing with supercritical CO2 . . . . . . . . . . . . . . . . . . 4.3.3 Electrokinetic and pseudo-electrokinetic packing . . . . . . . . . 4.3.4 Packing by centripetal forces . . . . . . . . . . . . . . . . . . . 4.3.5 Packing by gravity . . . . . . . . . . . . . . . . . . . . . . . . . Comparison o f packing procedures . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement ............................. Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. .

112 112 113

116 129 132 137

139

. . . . . .

140 144 145 150 150 152 152 154 155 156 158 158 159 159

112

Chapter 4

4.1 INTRODUCTION Capillary electrochromatography (CEC) can be performed in open tubes or packed structures. In the open tubular format, the stationary phase is fixed at the inner surface of a capillary column; columns with inner diameter of less than 20 gm are recommended for the best performance [ 1]. Packed structures, on the other hand, consist of a capillary tube filled with chromatographic media. These packed structures can be classified into three different groups: 1) columns packed with particles [2-30], 2) columns containing separation material that has been polymerized in situ, creating a "rod-like" monolithic structure also known as continuous beds [31-39], and 3) columns with entrapped particulate material, which are a combination of the first two groups [40-46]. Columns of the first group are the ones used the most in CEC and will be discussed in this chapter; the other two groups are discussed in Chapters 5, 6 and 7. The chromatographic packing material most commonly used in CEC is HPLC reversed-phase type on spherical particles (1.5-10 gm diameters); although new alternatives are being explored to fabricate materials with more applicability to CEC. As the CEC column technology develops, however, the preference of using columns packed with particles may change, particularly with the emerging approach of monolithic columns. Several protocols can be used to fabricate packed bed structures for use in CEC. In this chapter, we will discuss the packing techniques and column fabrication protocols that have been used for packing particulate material. We concentrate, therefore, on the different approaches used to deliver chromatographic particles into the capillary column. We present an overview of the different packing protocols available to the practitioner, as well as of the CEC column fabrication method, as performed in our laboratory. Our own experiences, practices, and views regarding packing procedures are also provided, when appropriate. 4.2 Column fabrication

Despite the several detailed procedures reported for the fabrication of packed columns for CEC [ 14,17,20,27,30,47-50], column fabrication may still be regarded as an art. A reliable and reproducible performance of a column depends on the column fabrication. Poorly packed columns can lead to low efficiency, poor resolution, and asymmetric peak shapes. The capillary tubes typically used to fabricate CEC columns are fused silica tubes with inner diameters of 100 gm or less, with 50 and 75 lam I.D. being the most popular. The small inner diameter allows for heat dissipation, which is generated by the applied electric field. Packing such columns is an elaborated process and a skill that requires experience.

Packed Bed Columns

113

4.2.1 The column

A packed column in CEC consists of two segments- a packed and an unpacked (or open) section, in most cases. A typical CEC column with a packed and an open segment is illustrated in Fig. 4.1. Most frequently, capillary tubes with less than 100 ~tm I.D. are packed with reverse phase HPLC materials of 1.5-10 ~tm diameter. The chromatographic material is kept in place by means of retaining frits (vide infra). The electroosmotic flow (EOF) velocity in each segment of the CEC column is different [51 ]. The overall EOF velocity depends on the fraction of the packed segment [51,52]. The resulting net EOF is thus a combination from both, the packed and the open segments. To facilitate detection through the column by spectroscopic means, the polyimide coating on the open segment, close to the outlet retaining frit, is removed; this provides the optical window for detection. This can be achieved by any of the methods already in use for capillary electrophoresis (CE) [53]; burning off the polyimide coating is the most common approach. Because of light scattering by particles, optical detection through the packed bed has been reported to decrease detectability [18,24,27]. The length of the column can alternatively be packed completely with the desired separation material (no open segment); however, if detection through the packed bed is not performed, connecting tubing to a detection system is required. Fig. 4.2 shows an example of a 75 ~tm I.D. fused silica column completely packed connected to a piece of a 50 ~tm I.D. capillary used for detection. The butt connector is made by inserting the two capillaries into a piece of PTFE shrinking tubing, which upon application of heat secures the two capillaries in place [54]. It has been reported that butt connection of capillaries has an insignificant contribution to band spreading [55]; however, care must be exercised since connecting of two different pieces of tubing

Packed Section

Open Section

..J

I ~176

""

"-'-'

.. . . . .

i

I Detection Window

Retaining Frits Fig. 4.1. Schematic of a typical packed-capillary column for CEC, illustrating the open and packed segments. References pp. 159-164

114

Chapter 4

Fig. 4.2. Photograph of a butt connection between a 75 ~tm I.D. packed fused silica capillary and a piece of a 50 ~m I.D. capillary tube. Reprinted from ref. [54] with permission. Copyright Wiley-VCH 1999.

2.0

i y

.

1.115

! §

L193x

R=

|

|

0.9986

1.8

-

E

1.6 /

/ 1.4

/

1.0 0.00

/

/ ,

i

0.10

,

i 0.20

,

I

0.30

,

I

0.40

,

| 0.50

,

/

0.60

Fractional length of the capillary segment packed with bare silica

Fig. 4.3. EOF mobility as a function of the fractional length of the bare silica packed segment for a 100 p,m I.D. capillary containing a 20 cm ODS segment. Reprinted from ref. [56] with permission. Copyright Wiley-VCH 1999.

always has the potential of introducing band broadening to the system. Nonetheless, this option is often used to connect detection schemes that offer higher detectabilities, such as Z-cell for UV detection, mass spectrometry (MS) and NMR detection schemes (see Chapters 2 and 8). In such instances, the gain in detectability and/or structural information is far more important than the loss in efficiency. Totally packed capillary columns, having one segment packed with the stationary phase and a second segment with bare silica, have been fabricated to control the EOF [56]. In this case, the segment that is open in a typical CEC column is packed with bare silica to accelerate and provide a steadier EOF. Such a configuration has allowed an increased EOF that translates into shorter analysis times. Fig. 4.3 and 4 show the EOF mobility as a function of the fractional length of a column packed with bare silica and the effect on analysis time, respectively. As the porosity of the bare silica particles is increased, the EOF is also increased [56]. Columns have also been packed

Packed Bed Columns

115

'i 5

a, u = 0.8 mm/s Nay = I01,000 plates/m E tt)

b.

~g

u = 1.0 mm/s Nay = 108,000 platefdm

2

"~'5

C, u = 1.1

mm/s Nay = 130,000 plates/m

r

2'0

Min

Fig. 4.4. Electropherograms illustrating the effect of the length of the bare-silica segment on the separation of probe compounds, a) 0 cm, b) 6 cm, and c) 28 cm. Solutes: 1, benzene; 2, toluene; 3, ethylbenzene; 4, propylbenzene; 5, butylbenzene and 6, pentylbenzene. Reprinted from ref. [56] with permission. Copyright Wiley-VCH 1999.

with a blend of bare silica and reverse-phase silica supports [18,57]. This can provide enhanced EOF due to the amount of silanols groups introduced by the bare silica, decreasing analysis time. This approach also reduces retention because of the decrease amount of stationary phase, as the bare silica replaces the bonded one; hence, retention depends on the blend ratio. Initial work on CEC was performed on drawn-packed capillaries [ 11 ], a procedure originally introduced by Tsuda et al. [58]. In this approach, large bore columns (thick walled Pyrex tubing) were packed with underivatized packing material; then the columns were pulled at high temperatures to a desired diameter using a glass drawing machine. The stationary phase was attached to the underivatized support packing material after the columns were drawn. This column preparation procedure is not currently used because of the low success rate in fabricating the columns.

References pp. 159-164

116

Chapter 4

4.2.2 Chromatographic material Because of the column's dimensions in CEC, it is important to consider a narrow size distribution of the particles. The effect of particle size distribution on separation efficiency in CEC is expected to be similar of that in HPLC. Although of the same nominal particle size, different packing materials can yield different efficiencies in CEC. It has been pointed out that the different efficiencies reported for the separation of polycyclic aromatic hydrocarbons (PAH), for example, using different packing materials of the same sizes, can be attributed in part to the size distribution of each material [59]. The structure of the packed bed can be influenced by the size distribution. A homogeneous packing size leads to well-packed beds, approaching a closed packed structure. This can be seen in Fig. 4.5, where panels A and B show SEM of the packed bed for columns that were packed with silica particles of about 3 and 0.5 ~tm in diameter, respectively. It is apparent that the particle size in panel A is not as homogeneous as that of the particles in panel B. Notice how the particles with the tighter size distribution form a better-packed bed. One of the most important properties of a column packing material for CEC is the ability to support EOF. This is not only necessary for the separation of neutral compounds but also to separate charged species as the EOF is responsible for the bulk transport of the mobile phase and analytes [7,16]. In the absence of EOF, only species with the appropriate charge will reach the detector. Therefore, packing materials with very favorable characteristics for EOF generation are desired in CEC. Many silicabase HPLC packings from 1.5 to 40 ~tm in diameter have been utilized to pack columns for CEC; those with the C18 reverse- phase being used the most. Other materials include C8 [60-62], phenyl [61,62], and C30 [63,64]. The surface silanol groups of the silica impart a negative charge to the packing material, leading to the generation of the EOF upon application of the electric field. The high surface area of the silica-based packed beds provides for the EOF to be generated mostly at the particle surface, with negligible contributions from the fused silica capillary walls [7,16,65]. However, it is apparent that not all C 18 reversed phase HPLC materials are suitable for CEC. For example, Table 4.1 summarizes electrophoretic mobilities observed on typical reverse-phase HPLC chromatographic materials, and Fig. 4.6 illustrates the CEC separation properties of several C 18 packing materials under identical separation conditions. Faster analysis times are achieved with those materials capable of generating a strong EOF. CEC Hypersil and ODS-1 type are popular among the reverse-phase materials since they seem to support the fastest EOF. These materials are HPLC supports that have not been end-capped, and therefore, a relatively large amount of silanol groups are left on the surface, which can generate EOF. The EOF decreases as the alkyl substitution at the packing surface is increased be-

Fig. 4.5. SEM of the packed bed for columns packed with silica particles of about (A) 3 and (A) 0.5 prn diameter, respectively.

118

Chapter 4

TABLE 4.1 ELECTROOSMOTIC MOBILITIES OF VARIOUS CHROMATOGRAPHIC MATERIALS UTILIZED FOR CEC

Stationary phase material

Electroosmotic mobility (x 10-4 cm2/Vs)

BDS-ODS Hypersil a

0.99

CEC Hypersil C 18a

2.26

Hypersil ODS b

0.14

LiChrospher RP-18 b

1.45

Nucleosil 5 C18 b

1.56

ODS Hypersil a

1.47

Partisil 50DS3 b Prontosil polymeric C30 c Purospher RP- 18b

<0.01 1.54 <0.01

Rainin polymeric C30 c

1.54

Spherisorb Dial b

0.80

Spherisorb ODS Ia

2.26

Spherisorb ODS II a

1.79

Spherisorb $50DS2 b

0.50

Zorbax BP-ODS b

0.68

aAdapted from [7]; separation conditions: (80:20) acetonitrile-Tris-HC1 50 mM, pH 8, 20~ EOF marker used: Thiourea bAdapted from [16]; separation conditions: (70:30) acetonitrile-3-cyclohexylamino-2-hydroxy-l-propanesulfonic acid 25 mM, pH 9.53, EOF marker used: Thiourea CAdapted from [63]; separation conditions: (95:5) acetone-1 mM borate buffer, EOF marker used: Acetone. cause of the concomitant decrease in silanols groups responsible for the EOF [65,66]. Packed beds formed with 3 lam particles containing C 18 phases have generated separation efficiencies above 300,000 plates/m [ 17,30]. Separation efficiencies larger than 500,000 theoretical plates/m have been achieved using 1.5 lam non-porous reversephase silica materials in several applications [67-71]. Fig. 4.7 shows an example of high separation efficiency in the separation of 14 explosive compounds using a packed bed containing 1.5 pm non-porous packing; over 500,000 plates/m were observed. SDS has been used in the mobile phase to prevent bubble formation (vide

Packed Bed Columns

119

2 mAU 80

3

BDS-ODS-Hyp~__~~~~,~

risorbODS2 ._.J

SpherisorbODS1 .....

0 ....

._ 1 1 ~ 2~ ....

~_

+ ....

7~5 . . . .

CECHypersllC18 1'0 . . . .

12.5

1"5 . . . . . . . 1~.5

2'o

mi,

Fig. 4.6. Separation of test compounds on five different reverse-phase C18 stationary phases under identical conditions. The samples were not identical but contained 1, thiourea; 2, naphthalene and 3, fluoranthene. Reprinted from ref. [65] with permission. Copyright Wiley & Sons 1997.

TNB

2,4DNT

TNT NB RDX

0

~

2,6DNT

2-Am DNT

4-Am

~

RetentionTime (minutes)

Fig. 4.7. Separation of 14 explosives in a 75 ~tm I.D. capillary 21 cm (total length of 34 cm) column packed with 1.5 ~tm non-porous reverse-phase silica particles. The mobile phase consisted of 20% methanol, 80% 10 mM MES, and 5 mM SDS; separation voltage of 12 kV. Reprinted from ref. [69] with permission. Copyright American Chemical Society 1998.

References pp. 159-164

Chapter 4

120

,= ~ ~

40, ,.

9

4: [:] V 30 I O O

Pressure ElectroPressure ElectroElectro-

D r i v e n 8pro D r i v e n 5#In D r i v e n 3/~m D r i v e n 3pro D r i v e n 1.5#m

+

z0

f

101

o.o

+ D

"

§

. 12o

Linear

. . . .

z:o

'

Velocity/ram

3:o

s "~

Fig. 4.8. Plate height as a function of mobile phase linear velocity for different particle diameters using pressure and electrically driven flows. Reprinted from ref. [11] with permission. Copyright Friedr. Vieweg & Sohn 1991.

infra) and to stabilize the EOF through dynamic modification of the alkylated surface, which also has an effect on selectivity [69]. The effect of particle diameter on efficiency in CEC is better appreciated in a plot of plate height versus linear velocity of the mobile phase, known as a vanDeemter plot, as depicted in Fig. 4.8. It is clear that as the particle diameter is decreased, the separation efficiency is improved. Further, and contrary to pressure driven LC, the use of EOF in CEC also allows for the use of relatively high linear velocities of the mobile phase without a detrimental effect on efficiency. Silica-base stationary phases have also been employed for enantiomeric separations in CEC [6,72-81]. In the initial work on chiral CEC, commercially available HPLC materials were utilized, including cyclodextrins [6,74,81] and protein-type selectors [73,75,80] such as human serum albumin [75] and Otl-acid glycoprotein [73]. Fig. 4.9, for example, depicts the structure of a cyclodextrin-base stationary phase used in CEC and the separation of mephobarbital enantiomers by capillary LC and CEC in a capillary column packed with such a phase. The column operated in the CEC mode affords higher separation efficiency than in the capillary LC mode. Other enantiomeric selectors are also use in CEC, including the silica-linked or silica-coated macrocyclic antibiotics vancomycin [82,83] and teicoplanin [84], cyclodextrin-base polymer coated silicas [72,78], and weak anion-exchage type chiral phases [85]. Relatively high separation efficiency and excellent resolution for a variety of compounds have also been achieved using columns packed with naproxen-derived and Whelk-O chiral stationary phases linked to 3 ILtm silica particles [79]. Fig. 4.10 shows the

121

Packed Bed Columns

,,co,_./~ %._

A

H~CO~ ~" 01

.,c

oc%.

~

co-[....o

O

OCH3

.,co-

~ , ~ ~

?

(I H2la (CH2)3

•

o~ '~

B

CH3 I N ...O

'~j

CH2CH,

HINO ~ ~ p-LC

p-CEC N, = 5606 N2= 5453

T"

!

I

o

1o

I

20

N, = 16 870 N2 = 19 415

~

[mini

9

0

I

10

I

20 [mlnl

Fig. 4.9. (A) Structure of permethyl-13-cyclodextrin linked to silica support by means of a sulfide spacer and (B) a CEC separation of hexobarbital in a column packed with the the material shown in A. Adapted from ref. [81] with permission. Copyright Elsevier 1998. structure of such phases and an example for the separation of an enantiomeric mixture of an anti-depressant in a column packed with the Whelk-O type phase, showing an unprecedented resolution with efficiency of 200,000 plates/m. A comprehensive list of References pp. 159-164

122

Chapter 4

A :

o2~

N

o

/\

NO2

O

(3R, 4S)-Whelk-O 1 ChiralPhase

(s)-Naproxen-derivedchiralphase

B

mAu

\s,/~

60 50 40 30 20 10

0

2

4

6

'8

I~lin.

Fig. 4.10. (A) The chiral stationary phases (S)-naproxen-derived and (3R,4S)-Whelk-O 1. (B) CEC enantiomeric separation of an antidepressant (N-[1-(4-bromo-phenyl)-ethyl]2,2-dimethyl-propionamide) on a column packed with (3R,4S)-Whelk-O 1 chiral phase immobilized on 3 t.tm silica. Adapted from ref. [79] with permission. Copyright Elsevier 1997.

materials used in CEC, including specific applications, is shown in Table 4.2. CEC applications have been discussed in details by Robson, et al. [86] and by Dermaux and Sandra [87].

B

TABLE 4.2

3

2

CEC PACKING MATERIALS AND APPLICATIONS USING PACKED BEDS

C

%

cn

n

k.

Stationary phase (size)

Components separatedlparameters investigated Mobile phase

Ref.

Hypersil C18 (3-5 pm)

Effect of applied voltage on EOF

MeCN-10 mM phosphate 60:40 vlv, pH 7 M e C N 4 mM phosphate 80:20 viv, pH 8

[631

Hypersil C 18 (5 pm)

Evaluation of non-aqueous mobile phase

MeCN-water 65:35 v/v, also DMSO, MeOH, DMF, formamide

[146]

Hypersil C 18 (5 pm)

Effect of applied voltage and buffer concentration on current

MeCN-1-4mM sodium tetraborate 80:20 v/v, pH 9.2

[147]

Hypersil C 18 (3 pm)

Study of column lifetime

MeCN-50 mM TRIS 75-80:25-20 vlv

[991

Hypersil C 18 (3 pm)

Testosterone, progesterone, androstenedione, norethindrone

MeCN-MeOH-20 mM TRIS-HCl37.5:37.5:25 vivlv, PH 8

[148]

Hypersil C 18 (3 pm)

Retinyl esters

2.5 mM Lithium acetate in DMF-MeOH 99: 1 vlv

[I491

Hypersil C 18 (3 pm)

Triazine herbicides

MeCN-25 mM NaOAc 5050 vlv

[651

Hypersil C 18 (3mm)

Cannabinoids

6-25 mM Phosphate 65-75:35-25 v/v, pH 2.57

Hypersil C18 (3 pm)

Barbituates (barbital, butethal, phenobarbital, amylobarbital, secobarbital, hexobarbital)

MeCN-50 mM MES pH 6.1-water 60-40:20:20-40 V/V/V;MeOH or MeCN-50 mM phosphate, pH 4.5water 40-50:20:40-30 v/v/v

[1501 [62, 1511

Hypersil C18 (3 pm)

Diuretics (metolazone, epitizide, bendrofluazide)

MeCN-5 mM ammonium acetate in water 50:50 - 20:80 viv

'a 4

E:

z

h

[152, 1531

TABLE 4.2 (continued)

Stationary phase (size)

Components separatedfparameters investigated Mobile phase

Ref.

Hypersil C18 (3 pm)

Steroids (hydrocortisone, prednisolone, betarnethasone, adrenosterone, fluocortolone, ...)

MeCN-5 mM ammonium acetate 1733 - 38:62 v/v

[152, 1531

Hypersil C18 (3 pm)

Steroids (aldosterone, testosterone, hydrocortisone)

M e C N 4 mM sodium tetraborate 20:80 vlv, pH 8

[211

Hypersil C18 (3 pm)

Steroids (aldosterone, testosterone, dexamethasone, beta-estradiol)

MeCN-20 mM sodium acetate-acetic acid pH 5, 70:30 v/v MeCN-25 mM TRIS pH 8,70:30 vlv

[I331

Hypersil C18 (3 pm)

Steroids (hydrocortisone, prednisolone, betamethasone, betamethasone dipropionate, clobetasol butyrate, ...)

MeCN-2 mM phosphate 80:20 v/v, pH 7.8

[I541

Hypersil C18 (3 pm)

Benzodiazepines (nitrazepam and diazepam)

5 mM Ammonium acetate in MeCN-water 50:50 - 80:20 v/v

[I531

Hypersil C18 (3 pm)

S-oxidation compounds

MeCN-25-55 mM TRIS-HCI 65-45:35-55 vlv

11551

Hypersil C18 (3 pm)

Isradepin and by-products

MeCN-2 mM sodium tetraborate 80:20 vlv, pH 8.7

[81

Hypersil C18 (3 pm)

Textile dyes (neutral azo and anthraquinone)

M e C N 4 mM borate 80:20 vlv, pH 8

[211

CEC Hypersil C 18 (3 pm)

EOF dependence on pH, buffer concentration, temperature, % organic in mobile phase

MeCN-50 mM TRIS-HC1,20-80:20-80 v/v, pH 8-9 (also [156, MES, pH 6, and NaOAC, pH 4) 1571

CEC Hypersil C18 (3 pm)

Column packing by centripetal forces

MeCN-4 mM sodium tetraborate 80:20 v/v pH 9

~1251

CEC Hypersil C 18 (3 pm)

Column packing by gravity

MeCN-50 mM TRIS 80:20 v/v, pH 9

~1581

0

S-

4

3 5: 4

CEC Hypersil C l 8 (3 pm)

Drying columns to improve performance

MeCN-4 mM sodium tetraborate 80:20 vlv, pH 9

CEC Hypersil C18 (3 pm)

Fish and vegetable triglycerides

50 mM ammonium acetate in MeCN-IPA-n-hexane 57:38:5 vlvlv

CEC Hypersil C18 (3 pm)

Derivatives of fish and vegetable oils, fatty acids

MeCN-50 mM MES 90: 10 vlv, pH 6

CEC Hypersil C18 (3 pm)

Macrocyclic lactone, streptomyces S541

MeCN-5 mM borate 80:20 v/v, pH 9

CEC Hypersil C18 (3 pm)

Cephalosporin cefuroxime axetil

MeCN-5 mM borate 80:20 v/v, pH 9

Spherisorb ODs-1 (3 pm)

Retention time reproducibility1parameters effecting

MeCN-50 mM TRIS 80:20 v/v, pH 7.8

Spherisorb ODs-I (3 pm)

EOF dependence on pH, buffer concentration, temperature, % organic in mobile phase

MeCN-50 mM TRIS-HCI, 20-80:20-80 v/v, pH 8-9 (also MES, pH 6, and NaOAC, pH 4)

Spherisorb ODs-I (3 pm)

Short end injection

MeCN-50 mM TRIS 80:20 v/v, pH 7.8

Spherisorb ODs-I (3 pm)

Voltage programming, comparison of ODs phases

MeCN-50 mM TRIS 80:20 vlv, pH 8.1

Spherisorb ODs- 1 (3 pm)

Synthetic nucleosides

MeCN-50 mM TRIS 80:20 vlv, pH 7.8

Spherisorb ODs-1 (3 pm)

C- and N- protected peptides

MeCN-50 mM TRIS 80:20 vlv, pH 7.8

Spherisorb ODs-I (3 pm)

Diuretic (bendroflumethiazide)

MeCN-10-50 mM NaH2P04 70:30 v/v, pH 3.5-9.8

Spherisorb ODs-1 (3 pm)

Steroids (Tipredane)

MeCN-50 mM TRIS 80:20 vlv, pH 7.8

Spherisorb ODs-1 (3 pm)

Steroids (digitoxigenin, digoxigenin, cinobufatalin, cinobufagin, bufalin)

MeCN-4 mM sodium tetraborate 70:30 vlv, pH 8 MeCN-water 0.1% formic acid 70:30 vlv

Spherisorb ODs-I (3 pm)

Steroids (budesonide, Steroid A)

MeCN-50 mM TRIS 80:20 vlv, pH 7.8

% '9

s

TABLE 4.2 (continued)

Stationary phase (size)

Components separatedparameters investigated Mobile phase

Spherisorb ODs-1 (3 pm)

Tricyclic antidepressants

MeCN-10-50 mM NaH2P04 70:30 vlv, pH 5.7-9.8

Spherisorb ODs-1 (3 pm)

Prostaglandins

MeCN-2 mM Na2HP04 75:25 vlv, pH 7.3

Spherisorb ODs-1 (3 pm)

Anti-epileptic drugs (phenytoin, primidon, ethosuccinimide, ...)

MeOH-5 mM borate 60:40 v/v, pH 8.5

Spherisorb ODs-1 (3 pm)

Cephalosporin cefuroxime axetil

Spherisorb ODs (3 pm)

Effect of applied voltage on EOF, comparison of retention between CEC and HPLC

Spherisorb ODs-2 (3 pm)

Study of column lifetime

MeCN-50 mM TRIS 75-80:25-20 vlv

Nucleosil C 18 (3-7 pm)

Effect of applied voltage, particle diameter, pore diameter, and buffer ionic strength on reduced plate hgt.

MeCN-50-100 mM phosphate 80:20 vlv, pH 6.9

Nucleosil 100-C18 (5 pm)

Neutral analytes from in vitro rxn. of carcinogenic tissue

Various mobile phase compositions

Nucleosil 100-C 18 (5 pm)

ATP, ADP, AMP

MeOH-2 mM dibutylamine pH 5, 10:90 vlv

Nucleosil 100-C18 (5 pm)

Antiviral drug suramin

MeOH-2 mM dibutylamine pH 5.0, 10:90 v/v

Nucleosil 100-C18 (5 pm)

Morphine alkaloids

MeCN-2 mM ammonium acetate 40:60 vlv

Nucleosil C 18 (5 pm)

Entrapped columns; efficiency

MeCN-0.1 mM acetate 80:20 vlv, pH 3.0

Nucleosil C 18 (5 pm)

Comparison of EOF between stationary phases MeCN-25 mM CAPS0 70:30 vlv, pH 9.53 and stationary phase surface area

Ref.

Nucleosil 10-5-C 18 (5 pm)

Food colorings

MeOH-10 mM ammonium acetate 20:80 v/v, pH 8.5

[174]

Nucleosil(5 pm)

Bare-silica effect on linear velocity and retention

MeCN-1.25 mM sodium phosphate 75:25 v/v, pH 6.0

[I751

Nucleosil 100-3-C18 (3 pm)

In-column vs. on-column photometric detection, precision and LOD

MeCN-phosphate 80:20 v/v, pH 7.3

~1761

Nucleosil I00 C 18 (3 pm)

1 / 2 ~ e awidth k vs. injection length; Il2peak width vs. volume fraction of water in sample solution

Nucleosil 100 C 18 (3 pm)

Influence of column I.D. and length, effect of applied counter pressure on linear velocity

MeCN-2 mM borate 80-90:20-10 v/v

~781

Nucleosil 100 C 18 (3 pm)

Evaluation of CEC-LIF system; precision determination

MeCN-2 mM borate 95:5 v/v

[I791

Nucleosil 100-3-C18 (3 pm)

Effect of organic modifier on EOF and retention of solutes

MeCN4.1-7 mM phosphate 80:20 v/v, pH 7.2; acetone-1 mM phosphate 80:20 vlv, pH 7.2

[I801

Spherisorb SCX (3 pm)

Comparison of EOF for SCX, C18, and open tube. Effect of organic content, ionic strength, and applied voltage on linear velocity.

MeCN-10 mM phosphate 20-80:80-20 v/v, pH 7.5 MeCN-1-20 mM phosphate 70:30 v/v, pH 7.5

[go]

Hypersil ODS/SCX (4 pm) Spherisorb C6ISCX (3 pm)

Separation of PAHs, parabens at different pH, comparison of SCXIODS and ODs phases, separation of triazines

MeCN-20 mM phosphate 80:20 (TRIS pH 8, MES pH 6, [65] sodium acetate pH 4) MeCN-25 mM TRIS 80:20, pH 8.0 MeCN-25 mM sodium acetate 50:50, pH 8

Hypersil C18ISCX (3 pm)

Separation and detection of dexamethasone, fluticasone propionate, and detarnethasone from plasma

MeCN-25 mM ammonium acetate 80:20 v/v, pH 4.0

[99]

*o

f:

iZ 8 2

TABLE 4.2 (continued)

Stationary phase (size)

Components separated/parameters investigated Mobile phase

Zorbax ODSS*(IO pm), Separation of nucleotides, dinucleotides, transfer-ribonucleic acids Nucleosil ODSS (5 pm) *Octadecylsulfonated silica(0DSS)

MeCN-aqueous phosphate 35:65 vlv, 9.75 mM phosphate, 3.25 mM tetrabutyl ammonium bromide, pH 6.50 MeCN-MeOH-l2mM ammonium phosphate30:40:30, V/V/V,pH 6.0

Ref [931

Zorbax ODSS (10 pm)

MeCN-1.25 mM phosphate 75:25 vlv, pH 6.00 Comparison of ODs with ODSS using EOF velocity, mobile phase pH, and retention factor

[951

Zorbax ODSS (10 pm)

Separation of purine and pyrimidine bases, nucleosides, effect of % organic on EOF

MeCN-4.8 mM sodium acetate 40:60 vlv, pH 4.50

[941

XTec SAX (3 pm)

Separation of inorganic anions (F-, C103-, and HCOO-, C1-, Br-, SO^^‘, SCN-, CIO~*-) aliphatic sulfonates

Varying concentrations of phosphate buffer pH 7.2, sulfate buffer pH 8.2, and nitrate buffer pH 6.8

[911

Spherisorb propyl SCX (3 pm) Spherisorb phenyl SCX (3 pm) Symmetry SCX (3 pm) Spherisrob Mixed Mode (3 pm)

Evaluation of various SCX phases, compariosn MeCN-50 mM phosphate 70:30, pH 2.3 MeCN-20 mM borate 70:30, pH 9.0 of e~ectroosomoticmobility at various pHs, separation of antidepressants, amitriptyline, nortriptyline, neutral test mixture

Packed Bed Columns

129

4.2.2.1 Ion-exchangers and mixed-mode phases CEC chromatographic materials should have relatively high surface charge to afford a strong EOF. The EOF generated in commonly used silica-base packing materials depends on the pH of the mobile phase, requiring pH conditions suitable to deprotonate the silanol groups at the surface. At a pH of 8 or above, high EOF is obtained; however, at a pH of 3 or below the EOF is significantly reduced or non-existent. One approach to minimize the EOF dependence on the pH of the mobile phase is to use packing materials containing a fixed charge at the surface throughout a wide pH range. To this end, one can envision a variety of stationary phases attached onto the surface of the silica support by means of the well-known silane chemistry (see Fig. 4.11A). Such phases can be ion-exchangers or materials specifically designed for

A CH 3

CH 3

Where R is: C 8 , C18, C30, SCX, SAX, etc...

~

so 3-

B

so a-

C

.

.

so aFig. 4.11. (A) Generic representation of silane chemistry used to bond stationary phases onto silica supports. (B) Examples of packing materials that will enhance EOF. (C) Mixed-mode phase designed by E1 Rassi's group. Adapted from ref. [95] with permission. Copyright Wiley-VCH 1998.

Referencespp. 159-164

130

Chapter 4

CEC. Fig. 4.11B shows three possible materials suitable for CEC that provides high surface charge to generate EOF. Ion exchangers (i.e., SCX, SAX) are used in CEC as a way to increase surface charge, hence EOF. Columns packed with these materials have been used to separate charged organic and inorganic solutes [88-92]. Packing materials with cationic [65,90,91,93-106] and anionic [46,102,107,108] exchange moieties are one of the main subjects of study in packed beds for CEC. In principle, these materials should provide for a strong and stable EOF through a wide pH range. A material containing sulfonic groups, like those in SCX (e.g., C6-SCX, phenyl-SCX, C3-SCX) imparts a permanent negative charge at the surface, providing EOF towards the negative electrode. The typical quaternary amines of SAX phases, on the other hand, provide a positive charge at the surface of the packing leading to EOF in the direction of the positively charge electrode. Incorporation of an alkyl chain bonded to the silica in addition to the charged moiety (see Fig. 4.11B) has lead to the so-called mixed-mode phases for CEC, in which the separation mechanism can involve hydrophobic and ion exchange interactions, as well as differential electrophoretic migration [65,66,9397,99,101-103,109]. The EOF generated in these materials has been shown to be higher than in typical reverse-phase [65,95,97,101 ]. However, there is some dependence on the pH of the mobile phase, as shown in Fig. 4.12. Such dependence is attributed to the residual silanols present on the silica support. It has also been reported that rather lower EOF velocities than expected have been observed with the SCX phases [90]. More detailed studies on these types of materials are needed to

3.0 pH vs C 18.ODS 1 2.5 E 2.0

~. 1.5 o 1.0 ~" 0.5 O 0.0

i

4

i

i

i

5 6 7 pH of the mobile phase

i

8

9

Fig. 4.12. EOF velocity as a function of the mobile phase pH for a C18 bonded silica and a mixed-mode bonded silica (SCX/C18). Reprinted from ref. [ 101] with permission. Copyright Elsevier 2000.

Packed Bed Columns

to

(a)

i ~

131

,

,

4

|

8

,

12

t (rain)

3

1

(b) 2

2

6 345

0 0

1

2

3

i

|

4

5

t (min)

Fig. 4.13. Separation of acidic compounds in columns packed with (a) 5 p,m Spherisorb-ODS and (b) 5 pm Spherisorb-SAX materials. In (a) the mobile phase was composed of 60% acetonitrile in 2mM phosphate buffer (pH 2.2) and in (b) the mobile phase was composed of 50% acetonitrile in 20 mM phosphate buffer (pH 2.2). The compounds were: 1, 3,5-dinitrobenzoic acid; 2, p-nitrobenzoic acid; 3, p-bromobenzoic acid; 4, o-toluic acid; 5, benzoic acid; 6, o-bromobenzoic acid. Reprinted from ref. [102] with permission. Copyright Elsevier 2000. completely elucidate their behavior in CEC. Most of the mixed-mode stationary phases in CEC have incorporated two different moieties directly attached to a silica support, one with the SCX group and the other with an octadecyl group, for example (see Fig. 4.11B). One mixed-mode material specifically designed for CEC was introduced by E1Rassi's group [93-95,103], which is bonded to the silica support differently (see Fig. 4.11C). They synthesized a silicabased stationary phase that incorporates a (7-glycidoxypropyl)trimethoxysilane sublayer attached to the silica support and a sulfonated layer is covalently inserted between the sublayer and an octadecyl top layer. Although less popular, packed beds containing anion-exchanger groups have been shown to be useful in CEC. For example, using a commercially available SAX mixed mode phase, Lubman and coworkers [107] have been able to achieve excellent separations of peptides. The selectivity of the mixed-mode stationary phases, as well as the ion exchangers, in CEC have shown References pp. 159-164

Chapter 4

132

6

(a) pH 3.0

2

1

5

3

4

4

3 2

. . . . .

i .

2

.

.

.

.

.

.

.

i

4

,

6

t( rain ) (b) pH6.0

4 2

0 t

0

1

2

4 3 t( rain )

.

l

. .

5

J

6

Fig. 4.14. CEC separation of peptides using the mobile phase 60% acetonitrile in 30 mM phosphate buffer with (a) pH 3 and (b) pH 6. Peaks: 1, benzyl alcohol; 2, Gly-Gly; 3, Gly-Thr; 4, Gly-Gly-Gly; 5, Glu-Glu; 6, Gly-Gly-Gly-Gly-Gly; 7, Glu-Glu-Glu. Reprinted from ref. [108] with permission. Copyright Elsevier 2000. to be significantly different to that obtained in conventional reverse-phase silica materials. This is particularly the case for charged compounds where ion exchange-type of interactions have a significant contribution to the separation [65,94,97,102,103,107,108]. For example, selectivity can be tuned by changing the pH of the mobile phase, which influences ion exchanging. These effects are clearly illustrated in Figs. 4.13 and 14 for the separation of peptides and acidic compounds in a packed bed containing SCX phase.

4.2.2.2 Submicron particulate materials The pressure limitations found in HPLC as the particle diameter is decreased are not present in CEC, since the EOF velocity (u) does not depend on particle diameter as seen in Equation 4.1"

Packed Bed Columns

_

U--

~ o ~ r ~ E _ ~eoE q

133

(4.1)

where eo, er, ~, E, 11, and bteo are, respectively, the permitivity in vacuum, the relative permitivity of the medium, the zeta potential, the applied electric field strength, the viscosity of the medium, and the electroosmotic mobility. Therefore, the use of electroosmosis to drive the solvent through a packed bed should allow for the use of very small particles to generate high separation efficiencies. In principle, the limitation on particle diameter in CEC is imposed by overlapping of the electrical double layer in the flow channels. The reciprocal of the double layer thickness (~c) is known as the Debye-Htickel parameter given by Equation 4.2:

1( =

_1_= F N/ 21 8 R T ~o ~;,-

(4.2)

where 6, F,/, R, and T are, respectively, the thickness of the double layer, the Faraday constant, the ionic strength, the gas constant, and the temperature in K. The other variables are as in Equation 4.1. Equation 4.2 shows that the thickness of the electrical double layer is dependent on the dielectric constant of the mobile phase and the ionic strength of the electrolyte present. According to classical double layer theory, as applied to thin capillaries, the diameter of a channel must be at least 20 times the thickness of the electrical double layer (dc>205) in order to minimize parabolic flow [110]. Under such conditions, 80% of the volume transport due to electroosmosis should be retained [ 110]. Overlapping of the electrical double layer becomes significant in channels with diameters less than 206 and the plug-like profile is lost, decreasing EOF considerably. The double layer thickness has typical values of less than 10 nm, depending on the electrolyte concentration. The thickness of the double layer as a function of acetonitrile concentration at different ionic strengths, using univalent ions, is shown in Fig. 4.15. To at least maintain 80% of the flow transport for the double layer thickness in the typical range of 1-10 nm, the minimum flow channel dimensions must be 20-200 nm. In CEC, the EOF depends on the column packing structure and pore size of the packing material [51,111-116]. In a packed bed, there are many interconnected channels between particles, which leads to a porous packed structure. The porosity of the packed bed dictates the permeability through the column. The average channel size between particles in a CEC column can be estimated if the packed bed is assumed a

References pp. 159-164

Chapter 4

134 --- 40

[

E

~ /,o,,er

L_

(11 >,, J

0.1 mM

30

03 d3 .....

O

C3 2O

1 mM

JZ2

10 mM

100 mM

4,-.

O

o~ 10 09 (11 t',r O

'-" F--

0 I,

I

0

10

....

I

..

I

20

30

.

,~. .....

40

I

I

50

60

J_.

70

I

80

.l .......

90

I,

100

Percent of Acetonitrile Fig. 4.15. Thickness of the double layer as a function of acetonitrile concentration in a water-acetonitrile mixture, as per Equation 4.1. Constants for the mixed solvents were obtained from ref. [49,129].

collection of capillary tubes with an average diameter corresponding to the channels between particles. Equation 4.3 shows a relationship between the mean channel diameter and particle size (alp), accounting for the particle structure through the interparticle porosity [ 113]:

dc = 0.42 alp g 1-e,

(4.3)

where ~ is the interparticle porosity. A fairly well packed column is considered to have a random packing structure with an interparticle porosity of 0.4 [117]; therefore, the channel diameter is given by:

dc- 0.28 dp

(4.4)

A similar channel diameter (i.e., 0.25 dp) was originally suggested by Knox who

Packed Bed Columns

3.530 ~o

2.5

135

9

~

................ --m---- 3 pm

i

iiiiiii

--T--0.2

2.0 O

o

~.5

LL

1.0

0 LU

0.5 00

: ..................... !................. ~;~:: ..........

t

.......... i"<

o

~m

!

.~ J

iiiiii

::

........ .......... ! .......... i .......... ' ........... ,........... "....... '

2OO

4OO

Field

6OO

8OO

strength (V cm -1)

Fig. 4.16. EOF mobility as a function of applied voltage for different particle diameters. Reprinted with from ref. [66] with permission. Copyright Elsevier 2000.

obtained it using the flow resistance parameter of a packed column [ 118]. Using a minimum channel diameter of 206 as the limit to avoid a significant overlap of the electrical double layers, one can estimate the particle size required to preserve most of the volume transport with plug-like profile [11]. For 8 values of 1-10 nm, which corresponds to channel diameters of 20-200 nm, the minimum particle diameter would then be 0.071-0.71 lam, for a column having a randomly packed bed. Unger and coworkers have reported that EOF is independent of particle size down to 0.2 ~tm [66], as predicted by Equation 4.1 (see Fig. 4.16). According to Equation 4, the average interparticle channel diameter for 0.2 ~tm particles is about 0.06 ~m. The experimental conditions reported by Unger and coworkers indicate that 8 is about 3 nm, corresponding to a channel diameter of approximately 0.06 ~m, which satisfies the condition of dc>20 8. EOF linear velocities above 2 mm/s have been achieved in CEC using packed beds with submicron particles and organic/aqueous mobile phases [66,120]. So far, the separation efficiencies reported with the submicron packed beds have not offered a significant improvement over those obtained with particle diameters in the 1 ~m range [66,119-121]. Fig. 4.17 depicts the separation of a test mixture obtained in a packed bed with particles of about 0.5 ~tm in diameter. As reported by Luedtke, et al. [ 121 ], plate heights of about three times the particle diameter (H = 3dp) are achieved. This has been attributed to band dispersion due to temperature effects and instrumental limitations, such as the maximum electric field that can be applied with existing units and detection systems [121]. Plots of plate height versus linear

References pp. 159-164

Chapter 4

136

1) Thiourea

2

2) Naphthalene 3) Ethylnaphthalene 4) Amylbenzene

I

I

I

0.4

I

0.8

I

I

1.2

I

1.6

Time/minutes Fig. 4.17. Separation of a test mixture in a packed bed of 0.5 ~tm (C8) particles. Column: packed bed of 15 cm in a capillary of 35 cm total length; mobile phase: 80:20 acetonitrile-50 mM Tris buffer at pH 8; separation voltage of 30 kV; injection, 3 s at 300 V; UV detection (220 nm).

velocity indicate that a minimum in plate height has not been achieved and higher electric fields are needed to achieve higher velocities (see Fig. 4.18). Nevertheless, the small particles do provide for rapid separations with current systems. Further studies are still required to obtain a complete understanding of the effect of the submicron material in CEC.

137

Packed Bed Columns

3.5 (# c

.~

~, |

3.1

~ X

~ ~

~1'~" C

E v

=L

F- 2.7 "1-

T

(9

0.2

0.6

w -1-

1.0 1.4 Ttme/mln

1.8

LU 2.3 F< J &.

1.9

1.5 ,

0.1

I

,

0.6

I

1.1

~

I

1.6

,

I

2.1

,

I

~

2.6

I

3.1

LINEAR VELOCITY (mm/s) Fig. 4.18. Plate height versus linear velocity for 9-(1-pyrene)nonanol, last eluting peak in electropherogram in the insert, obtained in a column packed with particles of about 0.5 ~m diameter. Column: 12 cm packed, 35 cm total length; mobile phase 80:20 acetonitrile-50 mM Tris pH 8; voltage 27 kV.

4.2.2.3 Highly porous materials

The typical porous silica-base materials commercially available have pore sizes close to 10 nm. Based on our discussion on the channel diameter requirements to support EOF, it is unlikely that EOF through such pores can be generated under normal CEC conditions. However, it is evident that intraparticle EOF exists in materials with relatively

large pores,

exhibiting perfusion

through

the particles

[45,56,116,122,123]. Flow transport through a highly porous particle is schematically represented in Fig. 4.19. With appropriate channel diameters, EOF can be generated within the particle, transporting the solute through. In pressure driven systems, there is no flow through the particle pores and solutes can only have access to the pores by diffusion. Perfusive transport through wide-bore silica particles with nominal pore

Referencespp. 159-164

138

Chapter 4

Deft,~;~',~.~%~, ---;--

Dapp~, ~:. ~?~"'"

9

(a) Pressure-drivenFlow (HPLC) Diffusion only

~o

[:::)m

..~,:~ .......

'" "'~ "

'

(b) Voltage-drlven Flow (CEC) Dlffuslon + Convection

Fig. 4.19. Schematic representation of intraparticle flow. In pressure driven flow there is no flow through the particle (A); in electrically driven flow there is intraparticle transport. In (A) transport of solute into the pores is accomplished solely by diffusion, whereas in (B) the EOF enhances transport through the pores. Reprinted from ref. [125] with permission. Copyright Elsevier 1999.

sizes larger than 200 nm was initially reported by Remcho and coworker [ 123]. Later Stol et al. showed intraparticle EOF in particles with pore sizes between 50-400 nm [122,124]. The EOF transport through wide-pore silica has been reported even in particles entrapped in capillary columns via sol-gel processing [45]. Double layer interactions within the pores are minimized by controlling the concentration of the electrolyte in the mobile phase. EOF through the pores is observed at high concentration of electrolyte, while at low concentration of electrolyte, double layer interaction occurs and transport through the pores can be stopped [122,123]. The effect of electrolyte concentration on separation efficiency for columns packed with 7 ~m (C18) particles containing pores with a diameter of 400 nm (nominal value given by manufacturer) is illustrated in Fig. 4.20 [122]. The flow through the pores provides for an enhanced mass transfer, resulting in improved separation efficiencies. Separation efficiencies of 430,000 theoretical plates/rn have been reported for the columns packed with the 7 ~m (C18) particles having 400 nm pore diameter [122]. The efficiencies and linear velocities observed with the highly porous packing materials are comparable to those obtained with small particle sizes. Therefore, the use of the highly porous large packing materials have been proposed as an alternative to the very small particles to perform fast separations with high efficiency in CEC [122,123,125]. The wide-pore large particles also offer the advantage of easier packing than submicron material, since aggregation is minimal. However, highly porous particles have the disadvantage of being fragile; hence, care must be exercised during packing to avoid damage of the particles. The enhanced particle porosity will also affect sample capacity adversely and the increase in ionic strength to maintain a thin double layer can lead to heating of the column.

Packed Bed Columns

13 9

10

7.5

H (pm) 5

2.5

0

1

2

3

4

u (mm/s) Fig. 4.20. Effect of the electrolyte concentration in the mobile phase on separation efficiency in a packed bed containing 7 pm highly porous C18 particles (Nucleosil 4000-7). Reprinted from ref. [122] with permission. Copyright Elsevier 1999.

4.2.3 Retaining frits The flits retaining the chromatographic packing material inside the capillary column in CEC seem to be the "Achilles heel" of the packed column fabrication process. They are the major problem in column manufacturing and perhaps the most critical parameter influencing column performance in general [5,126,127]. Most typically, the flits are fabricated by sintering silica-base packing material by means of heating. Using this approach, the CEC column becomes fragile at the flit since during its fabrication the protective polyimide coating is removed. There also seems to be lack of reproducibility and reliability in the manufacture of the flits, particularly between laboratories. The characteristics of the packing material at the flit position change as heat is applied to produce the flits. This creates non-homogeneous packing at the flit, having different electrical resistivities when compared to the open and pack segments of the columns. The different electrical properties of the flit can contribute to non-uniformities in EOF, and lead to bubble formation at the boundary between the flit and the unpacked segment of the capillary [ 10,51,127]. Constructing flits with resistivities similar to either the packed or the open segment can minimize discontinuities in the column structure, hence decreasing non-uniformities in EOF. Bubble formation, which has also been attributed to heat generation as the electric field is applied, can be reduced by several means. The most common approach to avoid bubble formation is to pressurize the mobile phase at both inlet and outlet column reservoirs References pp. 159-164

140

Chapter 4

[ 10,14,17,27,128]. Other common practices include the use of well-degassed solvents, low concentrations of electrolytes, a relatively large amount of the organic component in themobile phase, working at reduced temperatures (e.g., 15~ when possible, and the use of low conductivity electrolytes (i.e., zwitterionic buffers). The addition of sodium dodecyl sulfate (SDS) into the mobile phase at low concentrations has also been used to minimize bubble formation [67]. The effects that frits and packing materials have on EOF seem inconsistent. For example, placement of two frits in a capillary column (without chromatographic packing material) has shown to be flow restrictive points, reducing EOF by 35% compare to a capillary without frits [56]. On the other hand, an increase in EOF has been reported for a column packed with ODS material when compared to an equivalent open tube [90,129]. The discrepancies can be attributed to differences in the materials and procedures used to fabricate the frits, since these appear to be the major differences reported [ 130]. The bed-retaining frits must posses high permeability to solvent flow, yet the flits must be mechanically strong to retain the packing material and resist the pressures used to pack and/or rinse the column. The heating conditions and method used to prepare the flits affects such characteristics. Pressure resistance studies have shown that, within certain constrains, correlation between pressure resistance of a frit and its influence on the EOF is likely to be insignificant [ 130]. 4.2.3.1 Silica-base frits

The fabrication of frits has been studied in detail by several researchers [27,55,56,129-131 ]. Behnke, et al. [27] studied the performance of columns fabricated using three different frit fabrication procedures. In one of the procedures, the frits were constructed by sintering (using heat) a plug of silica gel wetted with potassium silicate. The frits were mechanically stable; however, under CEC conditions the columns with these frits showed baseline and electrical current stability problems. Another procedure used the method by Cortes, et al. where the flits were formed by polymerization of a potassium silicate solution containing formamide [132]. The columns fabricated using these frits suffered from similar problems. The third method involved the sintering of a silica gel plug wetted with water. Packed beds retained by these frits showed a stable baseline and current. However, they lacked mechanical stability with relatively large column diameters (150 ~m I.D.); decreasing the column diameter to 50 ~tm I.D., increased the stability of the frit. In a different study, Chen and co-workers optimized the silicate polymerization method [ 131 ]. In their approach, the outlet frit is prepared by first filling the column with a sodium silicate solution. Then, the portion of the column at which the frit is desired is brought in contact to a heating element for a few seconds and the frit is

Packed Bed Columns

141

formed. The polyimide coating is not removed under the heating conditions, reducing column fragility at the flits. The excess of the silicate solution is removed by pressure. The flit is then silanized with a solution of 0.02 mol/1 trimethylchlorosilane in DMF, using imidazole as an acid acceptor. The column is packed and the inlet frit is constructed by a quick dip of the column entrance in another silicate solution and heating. Optimal sodium silicate solutions were found to be 10.8% and 5.4% (w/v) for outlet and inlet flits, respectively. The method uses short heating times: 5-6 seconds for the outlet flit and about 1 second for the inlet one, producing relatively short flits (-~75 ~tm). Columns using these flits have been run without pressurization and showed to be mechanically stable without electrical current stability or bubble formation problems over a wide range of acetonitrile-water mixtures, even under relatively high currents conditions (-27 ~tA) [ 131 ]. The most commonly used approach to fabricate silica-base flits, however, is to sinter the actual chromatographic material in place. Care must be taken, however, in order to minimize degradation of the alkylated silica that will not be part of the flit [24]. Heating time used for sintering depends on column I.D., particle size, and type of stationary phase material to be fritted [133]. During heating, a substantial number of silanol groups can be created, changing the surface charge of the material at the flit [ 130]. Excessive heating also damages the outer side of the capillary column, which can create possible adsorption sites. Fig. 4.21 illustrates the effect of using excessive heating to fabricate the flit. Panel A in Fig. 4.21 shows a magnification of the outer side of the column and panel B shows the entire end of the column. Fig. 4.22 shows an adequate sintered frit. It has been shown that decreasing the created silanols by resilanizing the flits can reduce the likelihood of bubble formation, creating a uniform structure that resembles the packed bed, and leads to an improved separation [55]. Fig. 4.23 depicts electroctrochromatograms for the separation of a test mixture on 3 ~tm Spherisorb ODS 1 packed beds (a) before and (b) after resilinazing the outlet frit with ODS. Silanization deactivates undesirable adsorption sites at the flit with which the analytes can interact [ 134]. The packing functionality can also affect the final properties of the flits. This has been shown on flits formed in open tubes using different silica with different functionalities: bare silica, strong cationic exchanger (SCX), strong anionic exchanger (SAX), and Hypersil ODS silica, without the contribution of any other packing material present [130]. The EOF velocity obtained with each flit is different, depending on residual groups of the different packing. Frits formed with bare silica and SCX showed EOF velocities higher than an open tube and flits of ODS material. The amount of residual groups on the surface of the silica support depends on the heating time applied during the formation of the flit, and can contribute considerably to the overall flow. In a column containing a small flit formed with SAX material a reversal

References pp. 159-164

Fig. 4.21. Effect of excessive heating on the capillary column when fabricating a retaining frit, (A) outer surface of the capillary and (B) entire end of the capillary.

g

42 Y

Q

143

Packed Bed Columns

A

13

Fig. 4.22. Adequate frit without deformities on the capillary, particles are 1 pm in diameter.

of the EOF has been observed, when compared to that in an open tube, under otherwise similar experimental conditions. The temperature of the filament used to fabricate such a frit was 430~

References pp. 159-164

and heating time between 12 and 15 seconds. If the

Chapter 4

144

Before rccoating the fdt

(a)

e

t ,oo-{

b a,

400

I] II

d

em

8l~

1000

1200

Ttme / M.mutes

a Dimethyl phthalate b niU'obmzcne

1400

(b) After recoating the frit a

l l

I

x

5/)0

11 I [

I I

lO~X)

e

{~,_ D~.e~hylph~d~ a

II b

I

15oo

~obe~.e

It c. anisole

~1)o

2~/)o

Tune / 1,~u~es

Fig. 4.23. Electropherograms of a test mixture obtained in 3~m Waters Spherisorb ODS-1 packed bed (a) before and (b) after recoating of the outlet frit with ODS; mobile phase, 70:30 acetonitrile-water. Reprinted from ref. [55] with permission. Copyright Wiley-VCH 1999.

heating time is beyond 15 seconds, the EOF towards the positively charged electrode is reduced considerably, indicating a reduction of the residual positive charge responsible for the EOF reversal [130]. It is important to realize how critical is the heating during frit fabrication, as illustrated with Figs. 4.21; the process must be optimized, requiring experienced individuals. An alternative to sintering frits, which deserves mention here, is to form frits via UV photopolymerization of a glycidyl methacrylate and trimethylolpropane trimethacrylate solution (UV radiation, 365 nm for 1 hour) [135]. The photopolymerization process is similar to that used in the fabrication of monolithic columns (Chapters 5 and 6). Frits fabricated with this method have shown to be reproducible; since there is no sintering of packing material, weakening of the capillary column by removal of the polyimide coating and/or alteration of the stationary phase at the frit are avoided.

4.2.3.2 Fritless packed beds As an alternative to the formation of frits, the packing material can be retained by means of tapers fabricated on the fused silica column. In such a case, the packed bed is completely fritless. Two types of tapered capillary columns have been prepared for CEC: internally [136,137] and externally [133,138] tapered, which have been shown

Packed Bed Columns

145

A

B Fig. 4.24. Schematic representation of (A) external and (B) intemal tapers at the outlet of the CEC column. Adapted from ref. [ 136] with permission. Copyright Elsevier 1998.

to be useful in coupling CEC with mass spectrometry and NMR (Chapters 2 and 8). A schematic representation of the tapers is shown in Fig. 4.24. The internally tapered columns are fabricated by sealing the end of the capillary by means of a high temperature flame; the sealed end is carefully grinded to produce a small opening. A laser-based micropipette puller is utilized to make the externally tapered columns. In an attempt to have a fritless column, a capillary having an external taper was packed and the tapered end served as the entrance to the column; no outlet flit was constructed [133]. The problem with these columns is that they can only be used if the electrophoretic mobility of the packing material is larger than the EOF generated, so it can remain in place. Even in such cases, it can be problematic to keep the packing material inside the capillary. The external tapers are weak points on the column, prone to breakage; hence, externally tapered columns are inherently fragile. The internally tapered columns, on the other hand, are not fragile since only the inner diameter is reduced in dimension not the outer diameter and are easily connected to other tubing via shrink tubing. A manufacturing procedure for an internally tapered column is shown in Figs. 4.25 and 26. In Fig. 4.25, the internal taper is prepared at the outlet end of the packed bed, using a sintered frit at the entrance of the column. Fig. 4.26 describes the procedure for a fritless column. The internally tapered capillary columns can also facilitate the procedure of column packing by obviating the need for having a temporary retaining frit (vide infra). This approach holds great promise since it eliminates the problems associated with frits. Because of the connecting tubing, there is potential for band broadening, although it can be minimized.

4.2.4 Fabricating columns The most commonly used procedure to fabricate a packed capillary column for CEC is depicted in Figs. 4.27 A and B. Although several laboratories may have slight variations, the general procedure is as follows. A piece of fused silica capillary of a

References pp. 159-164

Chapter 4

146

Preparing the tapers A e.g. 60 cm length of fused silica capillary is sealed at the middle with a microflame torch.

fused silica capillary

The seal is cut to yield two tapered capillaries. Non-tapered end to be coupled are ground plane and smooth with P4000 (wet). Tapers are ground to form the desired orifice (i.d. 10-50 lam).

Coupling the segments dead volume free

internal tarter

The ground ends are carefully aligned and pushed together. The dual PTFE/FEP-connector is shrunk onto the junction. Slurry preparation 10 - 20 mg beads are ultrasonicated for 20 minutes in 70 - 150 lal acetone (or methanol). Packing the capillary The slurry is flushed in.

The stationary phase beads are allowed to settle under ultrasonication. Pressure drop to zero over a period of 20 min. The capillary is flushed with water.

e)

dual PTFE/FEP shrink-tube-connector

AP = 500 bar b.~ slurry

30 min. sonication AP = 500 bar acetone (resp.methanol)

30 min. sonication AP = 500 bar H20 r

Frit sintering (T=500~

AP = 500 bar

h~ p~

H20 f)

Burning the detection window

g)

Conditioning

inlet flit Column is flushed for 20 minutes with mobile phase (AP = 150 bar) followed by Electrokinetic conditioning:45 rain. at 10 kV with a 25 min. voltage - ramp 45 min. at 15 kV with a 5 rain. voltage - ramp g)

Storage Column is flushed for 30 rain with iso-propanol (AP = 150 bar) Capillary is stored with both ends immersed in iso-propanol filled vials

h)

~

[

UV - window

/

Internal taner

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

25cm

i

0.5-2cm

i

lOcm

i

Fig. 4.25. Procedure to fabricate a single-flit CEC capillary column. Reprinted from ref. [137] with permission. Copyright Elsevier 2000.

desired length is selected, usually about 10 cm longer than the actual column length wanted. Usually, the capillary tube is rinsed before use; it is our practice to rinse the tube with a sodium hydroxide solution (-0.1 mol/1) and then water. A provisional frit is sintered at one end of the capillary tube. Typically, this is accomplished by tapping onto a pile of wet silica material, allowing a small section of the column end to pack;

Packed Bed Columns

147

fused silica capillary

a) Preparing the tapers

b)

c)

Slurry Preparation

internal taper

Packing the capillary

AP = 500 bar

The slurry is flushed in.

~,~

slurry

The stationary phase beads are allowed to settle.

60 min sonication AP = 500 bar

Pressure drop to zero over a period of 20 rain.

acetone (resp. methanol)

Dry out overnight. d)

dual PTFE/FEP shrink-tube-connector

Coupling the segments dead volume free The ground ends are carefully aligned and pushed together. The dual PTFE/FEP-connector is shrunk onto the junction.

d)

Burning the detection window

e)

Conditioning

Storage g)

T w o - peace Fritless Column internal taper

i

i)

I

25cm

2cm

UV - window

[

10cm

I

T h r e e - peace Fritless Column int~rnnl tarter

I

25cm

internal tarter

I

1-2cm

U V - window

I ~

I

,0om

I

Fig. 4.26. Procedure to fabricate a fritless CEC capillary column. Reprinted from ref. [137] with permission. Copyright Elsevier 2000.

then heat is applied to sinter the material. The amount of heat required to form the flits depends on the column diameter and particle diameter [133], as well as the heating element used. Different heating elements have been used and they vary from

References pp. 159-164

Chapter 4

148

A

B

,

C

D

>

===C,

.

.

.

.

.

.

_1 .

.

.

.

.

/" wet silica

heating element

frit

flush

Fig. 4.27A. Representation of the steps involved in the column fabrication processes: (A) introducing with wet silica material into the end of the capillary, (B) the silica material is ready to fabricate the temporary frit, (C) fabrication of the temporary frit with a heating element, and (D) excess of silica material is flushed out after temporary frit is made.

optical splicers [28,123] thermal wire strippers [28], microtorch [18], burners [56], to heating elements incorporated into more sophisticated assemblies [ 14]. One relatively inexpensive setup used in our laboratory, makes use of a soldering gun fitted with a Nichrome ribbon (1 mm thick) as the heating element. The ribbon is punctured making a small hole (-0.5 mm) through which the column can be inserted; this facilitates heating of a small spot at any desired point of the capillary. Once the temporary frit is in place, the column is flushed to remove the excess material used to fabricate the frits. The mechanical stability of the frit can be tested by applying pressure using a HPLC pump. The frit must resist the packing conditions; yet, it should be permeable enough to allow solvent flow. The temporary flit is eventually removed. An alternative to the temporary frit is to connect the end of the capillary column to a union containing a metallic frit; this will retain the packing material inside the column during the packing process. The capillary is then packed to a desired length (vide infra). Once packed, the capillary column is rinsed with water,

149

Packed Bed Columns

-

2nd

1st

<~

I

H20

temporary frit

B

I

Fig. 4.27B. Formation of retaining frits once the capillary column is packed, (A) packed column is pressurized with water and the retaining frits formed using a heating element and (B) a fabricated column with frits and detection window in place.

and while under pressure the retaining flits are fabricated. Typically, the outlet flit is formed at a predetermined distance from the temporary flit. The inlet flit is then fabricated at a desired length (typically 20-30 cm) from the outlet flit. The use of organic solvents to flush the capillary column during frit formation is not recommended since carbonaceous material, presumably from the solvent, can be formed and remained at the flit. After the frits have been fabricated, the temporary frit is cut off and the excess particles inside the capillary are flushed out using a pump. In most cases, the heat applied to form the outlet flit is sufficient to remove a portion of polyimide coating on the capillary that is extending towards the open segment of the column. This exposes part of the fused silica, serving as the optical window for spectroscopic detection methods. After the flits and the optical window have been fabricated, the column must be rinsed with well-degassed mobile phase prior to connecting it to the CEC system. Once the column is connected to the CEC separation unit, it is a good practice to equilibrate the column with the mobile phase by applying voltage across the column in a stepwise manner. This can aid to consolidate the packed bed that may have voids [139]. The effect of having voids in a packed bed on peak shape is shown in Fig. 4.28. One approach to condition the CEC column is to apply 5 kV with the mobile phase in the column, and then the voltage is increased in 5 kV steps until 25-30 kV is reached. The column is allowed to equilibrate at each step

References pp. 159-164

Chapter 4

150

n'l/~:

0

'

/

4

'

'

I

6

'

'

'

I

8

'

'

'

I

10

'

'

'

""1

rrin

1NI

0~

8

10

Fig. 4.28. Electropherograms showing the effect of having a void in the packed bed (a). The peak shape improves after the bed is consolidated under voltage. Column packed with ODS-1, 3 l.tm, 75 l,tm I.D., 25 cm long (33.5 cm total). Mobile phase was 60:40 acetonitrile-2 mmol/1 KH2PO4. Reprinted from ref. [139] with permission. Copyright Elsevier 2000.

until a stable electrical current is observed. 4.3 PACKING METHODS

Several methods have been used to pack chromatographic particulate into capillary columns for CEC. These packing methods have made use of pressure packing using slurries [ 14,17,20] or using supercritical CO2 [30,43], electrokinetic packing [47,48], pseudo-electrokinetic packing [140], centripetal forces [49,141], and packing by gravity [50]. CEC columns packed by either electrokinetic or slurry pressure packing are commercially available. 4.3.1 Pressure packing

The most commonly used method to pack capillary columns for CEC utilizes the slurry packing methods typically used in HPLC. The capillary tube with a temporary frit is connected to a packing reservoir, such as a short HPLC column or another viable unit, which is connected to a high-pressure pump for solvent delivery. The slurry at a concentration of 50-100 mg/ml is prepared in a suitable solvent. Table 4.3

Packed Bed Columns

151

TABLE 4.3 SELECTED LIST OF SOLVENTS USED TO FORM SLURRY OF VARIOUS PACKING MATERIALS TO PACKED COLUMNS FOR CEC

Packing material

Slurry solvent

Packing solvent

Alltech 1.5 gm silica

Water

Water

Alltech 1.5 ~tm ODS AB

Hexane, THF b

Acetone

Alltech 5 gm SCX/C 18

THF, AcN c

Acetone

Astec 5jam Cyclobond I

AcN, THF, MeOH d

Acetone

Hypersil 3 jam ODS

Hexane, 2-propanol, THF Acetone, hexane

Hypersil 3 jam MOS

AcN, hexane, THF

Acetone

Hypersil 3 jam Phenyl

Hexane, MeOH, THF

Acetone

Micra Scietific 1.5 jam NPS ODS I

Hexane, THF

AcN, acetone

Micra Scietific 1.5 jam NPS ODS II

Hexane, THF

AcN, acetone

Micra Scietific 1.5 jam NPS C 18

Hexane, THF

AcN, acetone

Micra Scietific 1.5 jam NPS TAS-1

THF

AcN, acetone

Micra Scietific 3 jam NPS

AcN, water

Acetone, water

Spherisorb 3 jam ODS-1

Hexane, THF

Acetone

Spherisorb 3 jam silica

AcN, water

Acetone

Spherisorb 5 jam SCX

THF

Acetone

aReproduced from from ref. [ 139] with permission. Copyright Elsevier 2000. bTHF - tetrahydrofuran CAcN- acetonitrile dMeOH - methanol

shows a list of solvents used to prepare slurry of packing materials [139]. Sonication of the slurry is recommended to aid dispersing the packing material before the slurry is placed in the packing reservoir. The packing material is delivered into the capillary column at pressures of about 35-70 MPa (5,000-10,000 p.s.i.). Sonication can also be used during packing [20]. The pump is tumed off once the capillary tube is packed. To ensure that disturbance is not introduced to the packed bed when disconnecting the column from the packing reservoir, the column is allowed to "bleed" for a period of time. This means that the column is not disconnected until the system reaches ambient pressure. Then, the column is rinsed with water and the retaining frits are fabricated. References pp. 159-164

152

Chapter 4

The column is rinsed with the mobile phase by pressure and further equilibrated while applying the electric field.

4.3.2 Packing with supercritical C02 Supercritical CO2 packing was originally used to pack capillary columns for HPLC and supercritical fluid chromatography (SFC) [142,143]. The method is also used to prepare columns for CEC [30,43]. In this approach, one end of the capillary is attached to a pressure reservoir, which can be a short 2 mm I.D. HPLC column containing dry packing material. The other end of the capillary is attached to a connecting union containing a metallic frit; a temporary frit can be used instead. At the other end of the connecting union, a restrictor such as a piece of small fused silica capillary with a diameter of about 10 ~tm is attached to maintain the pressures required for supercritical conditions. During packing, the capillary column is immersed in an ultrasonic bath. The temperature of the bath is maintained at 60-70~

well

above the critical temperature of CO2. The column is packed at a constant pressure above the CO2 critical pressure, typically at 20-30 MPa (about 3,000-4,500 p.s.i.). To avoid disturbances to the packed bed, the capillary column is depressurized over a 4-5 hour long period of time. The column is then rinsed with water and the frits formed as described above. The column is flushed with the mobile phase by pressure and then equilibrated with mobile phase while applying the electric field.

4.3.3 Electrokinetic and pseudo-electrokinetic packing An alternative to pressure packing is the use of an electrically driven flow [47,48] to deliver the packing material. It is claimed that the particles are driven into the capillary column by the EOF, while the column and packing reservoirs are vibrated. Particles are suspended in a methanol-water mixture containing an electrolyte and sonicated before packing. An electrokinetic packing system used in our laboratory is depicted in Fig. 4.29. The slurry containing the suspended particles is placed in the upper vial, maintained in a vertical upside down position. A rubber septum cap is used for the vial, through which the capillary is inserted to reach the slurry. An electrode is also placed in the vial, serving as the anode. The outlet end of the column containing the temporary frit is inserted through a septum cap on the second vial, which also contains an electrode serving as the cathode. To aid the packing process, each vial is mechanically agitated. The agitation can be performed by attaching each vial to a speaker connected to a music system or a frequency generator. The electric field is applied until reaching 30 kV and maintained constant until the column is packed. Once the column is packed, the voltage is turned off, the column removed and pressurized with water. Then, the retaining frits are fabricated and the column equilibrated

Packed Bed Columns

153

Islurry vial electrode

J

capillary column

y

particle flow

/'I"

e,ectrode/-/ solvent reservoir speaker Fig. 4.29. Schematic of electrokinetic packing apparatus. as stated above. One of the advantages of electrokinetic packing is that the use of special fittings and high-pressure pumps are not required. The method can also be carried out at low operational cost and allow for the simultaneous packing of multiple columns. A pseudo-electrokinetic packing procedure has also been utilized to pack CEC columns [ 140]. This approach incorporates the use of a high electric field and hydrodynamic flow to pack the capillary columns. The column, equipped with a temporary frit, is mounted in the CEC unit and flushed with background electrolyte (e.g., 75:25 methanol-water mixture containing 10 mmol/1 Tris buffer). The entrance of the capillary is placed in the slurry, which is made using the background electrolyte. A portion of the slurry is pumped into the column to partially fill the capillary tube. Voltage is applied and the material inside the capillary packs against the temporary frit; packing References pp. 159-164

Chapter 4

154

is completed in approximately 15 minutes. Once the column is packed, it is connected to a conventional HPLC pump, flushed with water, and the permanent frits fabricated. The column is then preconditioned with mobile phase prior to use.

4.3.4 Packing by centripetal forces Packed bed columns for CEC can also be obtained by using centripetal forces [49,141]. Packing of the particles is obtained by centripetal acceleration through the capillary column. The velocity of the particles is given by the sedimentation velocity (used) as follows [49]:

(4.5)

(Pdp -- 90) V 0) 2 r Used =

3 ~ rl dp

where 9dp and Po are the density of the particle and medium respectively, V is the volume of the particle, co is the rotational speed, r is the distance from the center of rotation, 1"1is viscosity of the solvent in which the particles are dispersed, and dp is the particle diameter. A schematic diagram of a system used to pack capillary columns by centripetal forces is presented in Fig. 4.30. Particles are slurried in an appropriate solvent (-10-50 mg/ml) and placed in the slurry reservoir; the velocity of the particles is higher in low viscosity solvents such as acetone. In the apparatus shown in Fig. 4.30, two columns containing temporary frits are connected to the reservoir. Two

cell

slurry reservoir Column

l

i

stainless steel support arm

Motor pu ley

Fig. 4.30. Schematic of apparatus to pack by centripetal forces.

Packed Bed Columns

155

extending arms support the capillary columns. This means that the packed columns are inside the extending arms, which are made of 1 mm I.D. stainless steel tubing to avoid wrapping of the flexible fused silica tubes around the central reservoir during packing. Rotation of the apparatus forces the particles to move into and through the columns, sedimenting at the fritted-end. Columns are packed in about 5 to 15 minutes, depending on the solvent used to prepare the slurry. For example, 25 to 30 cm length of a 50 ~tm I.D. capillary column is packed in 5 min at 2,000 rpm using a 10 mg/ml slurry of 3 ~tm ODS packing material in acetone. Once packed, the column is submitted to the same procedure of rinsing and frit fabrication above-mentioned. Packing by centripetal forces also provides for the packing of multiple capillaries at once. A bed-drying step has been introduced into the packing procedure, once the column is packed by centripetal forces [ 141]; although this can be applied to any other packing procedure. The drying step is performed prior to frit fabrication and after frit formation the packed bed is resolvated. This step increases the column efficiency by about 15-20% and retention by 13%. However, despite the improvements, the drying step adds time to the column fabrication protocol because of both additional drying and rewetting of the capillaries. Perhaps it may not currently be worth to consider such step for a relatively small gain. However, if a more convenient dry-packing procedure is developed and the column preparation time can be similar to the wet procedures, it would be advantageous to use this approach.

4.3.5 Packing by gravity Particles can also be delivered into the capillary column by means of gravitational forces [50], which is also based on sedimentation velocity given by the following expression [ 144,145]:

Used =

(1-~) -k [13o(1-~,) + 19o(ec--1)] ~ g 18q

(4.6)

where Used is the sedimentation velocity, ~ is the volume fraction of the particles in the slurry solution, -k is a particle/solvent-dependent constant, 9p is the density of the skeleton of the particle material, 9o and 1] are the density and viscosity of the liquid, respectively, 8i is the particle porosity, ef is the fraction of the total hydrodynamic particle volume that is filled with suspension liquid, g is the gravitational constant, and dp is the particle diameter. In this method, gravity is used to transport the packing material into the column. The packing device is a 1-ml syringe containing about

References pp. 159-164

156

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100 pl of a 10 mg/ml slurry in acetone, which connects to the open end of the capillary to be packed via plastic tubing attached to the syringe needle. To avoid slurry solvent evaporation, the syringe plunger is attached and secured in place by means of a screw top. Capillary columns with a temporary frit are filled with acetone and connected to the packing apparatus. Sedimentation is allowed to proceed for a period of at least 10-12 h, replacing the slurry with a freshly sonicated one about every 4 hrs. Once packed, the columns are submitted to the rinses and frits fabrication procedures mentioned above. Columns with inner diameter of 50 ~tm and length of 20-28 cm have been packed with 3 ~tm particles [143]. Columns have also been packed with submicron packing material using this method [ 119]. This packing procedure is relatively simple, however, relatively long times of 12 to 48 hours, depending on particle size, are required to fill the capillary columns, before they are submitted to any pressurization step. 4.4 COMPARISON OF PACKING PROCEDURES The reported efficiencies for columns packed by the different methods vary considerably. Many factors can be attributed to this effect: experience of the analyst in making the frits, the use of different packing materials (although with the same nominal particle size), the use of different separations systems such as different sizes of the illumination spot on the optical window where detection is performed, that can be 200 ~m-2 mm long and localized after or before the end frit, and different separation conditions. These many variables make it difficult to assess which packing procedure offers the best-packed column by just examining results described in the literature. The effect of these variables must be minimized in order to achieve a true comparison among packing procedures. Ideally, one should compare columns fabricated using the same packing material under similar conditions, where the only difference is the packing method used. The same analyst should also construct the frits, perform packing and testing, using the same CEC apparatus. For a further comparison, experts in each packing technique can pack columns using the same packing materials and all columns would be then tested for performance by a single individual. This would indicate how much the experience on a particular packing method would influence the performance. In our laboratory, a study has been initiated taking into consideration all the above-mentioned factors. The column fabrication, packing, and testing have been performed by the same individual for columns packed by centripetal forces, electrokinetic packing, and pressure packing (slurry and supercritical CO2 packing). Fig. 4.31 shows electrochromatograms for a test mixture separated under similar conditions using capillaries packed by different protocols. Thus far, the data indicate that slurry

157

Packed Bed Columns

5

A

3 2

4

B

1

5

3 2 6

4

C

3

.......... v~ 5

D

3

.-____k I

0

I

2

I

4

1

6

Time/minutes Fig. 4.31. Separation of a test mixture on capillary columns packed by different methods: (A) pressure packing, (B) by centripetal forces, using supercritical fluid, and by electrokinetic packing. Columns were 50 pm I.D., 20 cm packed (30 cm total length); mobile phase 80:20 acetonitrile-4 mmol/1 aqueous borate. Separation voltage of 20 kV. Solutes: 1, thiourea; 2, benzyl alcohol; 3, biphenyl; 4, dimethylnaphthalene; 5, ethylnaphthalene; 6, amylbenzene.

References pp. 159-164

158

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pressure packing renders column with the lowest efficiency and that the highest retention is associated with columns packed by centripetal forces. Under the experimental conditions used, columns packed by supercritical CO2, centripetal forces, and electrokinetic packing showed similar separation efficiencies, about 216,000 plates/m, while pressure packed column showed about 150,000 plates/m, for the most retained compound. The columns packed by centripetal forces and by supercritical fluid showed retention factors slightly higher than the other two packing methods. This indicates that more retentive material is inside the column, which is the result of a tighter packed bed. From the four methods, electrokinetic packing is the simplest and easiest method to implement and use. A thorough investigation is in progress with all packing procedures at the time of this writings. 4.5 CONCLUSION Slurry pressure packing procedures have been the most commonly used to pack capillary columns for CEC; this departs from the well-established knowledge in packing columns for HPLC. The other methods discussed in this chapter, however, are also suitable to pack columns for CEC. In our experience, the other methods offer higher separation efficiencies. Further, packing of very small packing materials (< 1 lam) can be handled easier by the electrokinetic, centripetal, supercritical fluids and gravitypacking procedures, although with each technique the packing parameters must be optimized experimentally. Thus far, the preference of which method to use seems to depend on the familiarity of a given laboratory with a particular procedure. It is clear that column fabrication in CEC requires skill and experience, particularly constructing the retaining frits, a process that remains as one of the major problems in the column fabrication process, and improvement is still necessary. With the advances in CEC column technology, the problems associated with frit fabrication will be minimized. The monolithic and entrapped structures are clear alternatives, which will develop further. CEC practitioners may opt to obtain packed columns from commercial sources, as typically done with HPLC columns. This will avoid the implementation and optimization of a packing procedure in the laboratory, which can be laborious, time consuming, and require experience. 4.6 ACKNOWLEDGEMENT We acknowledge financial support from The National Science Foundation (CHE9614947).

Packed Bed Columns

159

4.7 ABBREVIATIONS

CEC CE EOF HPLC I.D. LC mL mM ~tm MS NMR ODS PTFE PAH p.s.i. SAX SCX SEM SDS SFC TRIS UV

capillary electrochromatography capillary electrophoresis electroosmotic flow high-performance liquid chromatography inner diameter liquid chromatography milliliter millimolar micrometer mass spectrometry nuclear magnetic resonance octadecylsilane polytetrafluoroethylene polyaromatic hydrocarbons pound per square inch strong anionic exchanger strong cationic exchanger scanning electron micrograph sodium dodecylsulfate supercritical fluid chromatography tris(hydroxymethyl)aminomethane ultraviolet

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97 98 99 100 101 102 103 104 105 106 107 108 109 110 111

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Chapter 5

Capillary Electrochromatograph y on M o n o l i t h i c S i l i c a C o l u m n s Nobuo TANAKA* and Hiroshi KOBAYASHI

Department of Polymer Science and Engineering, Kyoto Institute of Technology, Matsugasaki, Sakyo-ku, Kyoto 606-8585, Japan

CONTENTS

5.1 5.2 5.3 5.4 5.5 5.6

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monolithic silica columns . . . . . . . . . . . . . . . . . . . . . . . . . Preparation procedure of silica monoliths from silane monomers . . . . . Structural properties of monolithic silica columns prepared in a capillary Performance of monolithic silica columns in CEC . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

165 167 169

170 173 180

5.1 I N T R O D U C T I O N

There is much interest in high-efficiency- and high-speed separation media for liquid chromatography. The plate numbers available in practice have been in the range of 10,000-30,000 in HPLC for 20 years or so, but these are low compared to well over 100,000 theoretical plates in capillary gas chromatography or in capillary electrophoresis. This is caused by the limitation in the use of small-sized particles for HPLC, where a particle-packed column is commonly used under a pressure-drop of up to 40 MPa. An increase in column efficiency by using small particles, which is the approach taken in the past, is accompanied by an increase in the pressure-drop, as expected from Eqns. 5.2 and 5.3, below. Eqns. 5.1-3 describe the efficiency (plate height) and flow resistance of a column packed with particles [1-3], where N stands for the

Chapter 5

166

number of theoretical plates; L is the column length; H is the height equivalent of a theoretical plate; cy2, the dispersion of a solute band; dp, the particle size; u, the linear velocity of the mobile phase; Dm the diffusion coefficient of a solute in the mobile phase; Cx, the coefficient for the contribution of each term; % the flow resistance parameter; AP, the column pressure drop; to, the column dead-time; and 1"!,the solvent viscosity.

H~

H=

cs2 L

m

L

(5.1)

N

1 1

Dm

+

CdDm u

Csm4bl

+ - -

(5.2)

Dm

Ce---~p + Cm a~p-~--s

~to4

( h O - -rlL ~ 2

(5.3)

The current target of HPLC column development seems to be the generation of over 100,000 theoretical plates per column, nearly an order of magnitude greater than the column efficiencies currently used. Up to now, the most feasible approach is to use a capillary column packed with small silica particles in CEC. Among the recent high-performance liquid chromatographic methods, CEC has been the most widely studied. As stated in the preceding chapters, the most notable features of CEC can be found in the properties of solvent flow: first, that the solvent flow is provided by an electroosmotic flow (EOF) in the absence of pressure; and that the EOF has a plug-type flow profile. These features allow the use of small particles in CEC. The solvent flow is provided by EOF, which is not affected by the particle size and is free from the problem of pressure drop [4,5]. It also allows the use of a long column. In addition, the contribution of the A-term (the first term of Eqn. 5.2) to the band broadening is known to be much less significant in CEC than in a pressure-driven LC [3,6], because of the plug-type flow profile of EOF, leading to the higher column efficiency. On the other hand, with CEC there are some problems in the preparation and

Monolithic columns

167

operation of a column packed with particles. These problems have been found in the following areas (i) It is hard to pack small particles into a capillary to form a bed producing high efficiency. (ii) The frit can suffer failure, or generate bubbles. (iii) The mechanical stability of a packed bed is not adequate. (iv) It is not easy to operate a column packed with small particles. (A column having low permeability needs a long time for washing or equilibration). (v) Bubble formation during operation is a common problem. (vi) The chemical stability of stationary phase is not adequate. (vii) The reproducibility is often a problem, owing to the (slow) irreproducible EOF. Bubble formation owing to Joule heating, and the instability of the packed bed have been the most serious problems in CEC caused by the non-ideal frits and the structure of a packed bed. The high flow-resistance seen with a column packed with small particles in the pressure-driven mode is also a disadvantage, because it is difficult to wash or equilibrate the column after packing or when the mobile phase is changed. It is also difficult to run pressure-assisted CEC or electric-field-assisted HPLC using a column with low permeability. A monolithic column that possesses a one-piece network structure, similar to a long frit, can solve problems (i)-(iii) in CEC in a straightforward manner. Actually, the effort to prepare stable and high-efficiency columns for CEC has, at least partially, contributed to the recent development of monolithic columns. In addition, monolithic columns with small-sized skeletons and large through-pores can reduce the diffusion path-length and flow resistance. Thus, a support structure having large (through-pore size)/(skeleton size) ratios, which is not possible with a particle-packed column, can provide both a low pressure-drop and high column efficiency in HPLC, and will contribute to the improvement in CEC or CECmHPLC-hybrid operation. Monolithic columns can also solve problems (iv) and (v) given above, which are associated with particle-packed columns having frits in CEC. The problems associated with small particles can also be solved by ultra-highpressure liquid chromatography or open-tube liquid chromatography. None of these methods has yet been widely accepted as a routine separation method. Here we will review silica monoliths that can be prepared from alkoxysilanes, and their performance in CEC. 5.2 MONOLITHIC SILICA COLUMNS Monolithic silica columns can be prepared by bonding silica or ODS-silica particles together, by sintering or by embedding silica particles in a silica gel structure, References pp. 180-181

168

Chapter 5

,.."

!i!ig"i.i:i!i:i~i~

.- .$:~

.

.......

~...

.........("

.......... !i~

~&: i,......~: :!~ii,,: ~i::::~,~.i::!ii~i!i ~

(a)

!!iiiiiii'iiiii!i!! ! iii iiiiiiiiiiii!iii ]!]!iiii!i!!iiiiiiiiiiii !iili ::!ii!i!i i!!!!!ii ii!!iiiii!!iii!iiiiii!ii!!!i!!iiiiii !ii?iiiiiiiiii!!iii!i I if?i;i!?i~!ii i i ~i',i!i !i'~i !i!i i !i i i i i i!~ii i i i i!~,!!ii~ii'i i,!ii!iil !!i'i,ili}i!i !i ! i:i~i~i:!~7i~:i:~i~i~i!~i~iiiii~iii~i~i!~ii~i~i:i .~ii:i: ~ .~,.,i~i!i!i~i~i~!~i:i~i~i~i~i!i}iil}i!i~i~i,':

~:~: .:: ::::::::::::::::::::::::::::: ::::::::::::::::::::::::::::::::::: ::::::::::::::::::::::::::::::::: :::::::::::::::::::::::::::::

:::::::::~ ~::::::~::::.::;:: :~.~: .%:: :.,: ::::::::::::::::::::: ,::::::i ::::::::::::::::::::::::::::::::::::::::

::::::::::::::::::::::::::::::::::::: :::::::::::::::::::::::::::::::::::::::::::::::::::::::::::::::",~::::::.:::~:.:,:.:,:.;.:,:,,.:.:.:.:,:.:,:.:.:.'.:.:,:.:.:,:

isi~isiiii:,?iii: ?iiigs~::!!iisi::sii::i!ii:.::::!;:is!:: ,-,:.:.:.~,~:.:,,:~.-.~,:.:...:.~v 9

..:

::~x:: ~,:: ..: ::

~' ~ ~...

:il!!i!i::![i~ii !g ~;~N ::iii g::iii:.'.ii:::.~ii~ ~ ~ii~? * ::::::::::::::::::::::: ::::::::::::::::::::::::::::::::::::::::::::::::::::

:.:,

:ii:!!)iiii;iiiiiiii!iiiiiiiii!!!iiii!i ! ?ii:

(b) Fig. 5.1. Scanning electron micrographs of continuous-bed columns prepared from ODS-silica particles packed in a capillary. Prepared from (a) 75 ktm capillary packed with silica particles by sintering in the presence of NaHCO3 [9], and (b) large pore (left) and small-pore (right) ODS particles by a sol-gel method [10]. Reproduced from refs. 9 and 10, with permission. after the particles have been packed into a capillary [7-12]. Dulay et al. prepared a silica gel matrix from tetra-ethoxysilane in the presence of ODS particles by using a sol-gel process and they and others have reported on the gel-filled columns [7,8]. Similarly, Tang and his co-workers reported the preparation and evaluation of monolithic columns made by bonding ODS particles [10,11]. Figure 5.1 shows the SEM photographs of monolithic silica columns derived from silica particles. Asiaie and his co-workers obtained a porous-silica-based monolithic column by sintering ODS particles packed in a capillary column, by heating in aqueous sodium bicarbonate solution [9]. The stabilization of a packed bed of silica particles by external heating of the

169

Monolithic columns

packed capillary was reported to be effective [12]. These monolithic columns, prepared by the consolidation of a silica-packed bed, showed high mechanical stability and column efficiencies of up to 66,000 plates/30 cm in CEC [ 10]. So far, good results have been obtained with monoliths prepared from relatively large particles. Ideally, monoliths prepared from small particles will take full advantage of the small A-term and C-term in Eqn. 5.2 without the problems in pressure drop or bed stability in CEC. Monolithic silica can also be prepared by hydrolytic condensation of tetra-alkoxysilanes, a process similar to the preparation of particles [13-19]. The formation of the network structure utilizes spinodal decomposition of the polymerization system of tetramethoxysilane in aqueous acetic acid in the presence of a water-soluble polymer (e.g., polyethylene glycol) [20-22]. The preparation processes starting from monomers seem to be simpler than those starting from a particle-packed column, and the products can have higher permeability. Fields reported that continuous silica xerogels prepared from potassium silicate solutions could be used as highly permeable support media, and exhibit reasonable chromatographic efficiency in HPLC [23]. Minakuchi et al. reported the preparation and evaluation of continuous porous-silica columns that provide a much higher column efficiency in HPLC than do conventional columns packed with particles [13-16,18]. The monolithic columns prepared in a capillary can also be used in CEC. 5.3

PREPARATION

PROCEDURE

OF

SILICA

MONOLITHS

FROM

SILANE M O N O M E R S

The preparation of a monolithic silica from tetra-alkoxysilanes in a capillary is more straightforward than in a mould, because the most difficult step in the monolithic silica column preparation (the cladding of the monolith with engineering plastics such as poly(oxy- 1,4-phenyleneoxy- 1,4-phenylene-carbonyl- 1,4-phenylene) (PEEK) after the preparation in a mould) can be avoided [17,18]. The preparations in a capillary do, however, have some difficulties. Voids can develop between the silica structure and the capillary wall, owing to shrinkage of the silica during the preparation [24]. (In the case of the preparation of silica monoliths in a mould, as much as 30% shrinkage was observed.) The problem of shrinkage associated with the polymerization of tetramethoxysilane must be solved practically by optimizing the preparation conditions, and by attaching the silica structure to the capillary wall. A monolithic silica can be prepared from tetramethoxysilane in a fused silica capillary of 50-100 ~m in diameter by the following method [ 17,18], 1. Treat a fused-silica capillary tube (50-100 ~tm I.D., 370 ~tm O.D.) with 1M NaOH solution at 40~ for 3 h, prior to the silica preparation.

References pp. 180-181

170

Chapter 5 2. Add tetramethoxysilane (4 mL) to the aqueous solution of poly(ethylene glycol) (1.06 g, Mw = 10,000, Aldrich) in 0.01 M acetic acid (10 mL). 3. Stir the mixture at 0~ for 45 min. 4. Force the resulting mixture into the fused-silica capillary tube, and allow it to react overnight at 40~ 5. Wash the monolithic silica thus formed with water, then treat it with an aqueous ammonium hydroxide solution (0.01 M) at 120~ for 3 h. 6. Wash the monolith with ethanol and dry it with air. 7. Treat the monolith after drying at 330~ for 24h. 8. Let the monolith react with octadecyldimethyl-N,N-diethylaminosilane in toluene at 50~ for 2 h. 9. After the preparation, cut off portions (10-15 cm) which have large voids from each end of the capillary. 10. Prepare a detection window by using a heating device made of a coiled nickel-chrome wire.

Usually, two 100 cm long monolithic columns were prepared from the same reaction mixture, and two-four 33 cm long columns were obtained from the two 100 cm long silica capillaries containing silica monolith. The capillary columns (100 ~tm I.D.) showed 10,000-12,000 theoretical plates for the effective length of 25 cm under optimized conditions in a pressure-driven mode, and up to 40,000 plates in the CEC mode. The use of smaller-sized capillaries, e.g., 50 ~tm I.D., and the modification of the preparation method of mesopores, resulted in a monolithic silica column of higher efficiency and higher mechanical stability [25-27]. Under optimized conditions, 80,000 plates were obtained with a 25 cm column in CEC. 5.4

STRUCTURAL P R O P E R T I E S OF M O N O L I T H I C SILICA COLUMNS PREPARED IN A CAPILLARY The SEM photographs in Fig. 5.2, of the monolithic silica prepared in a fused

silica capillary show that the silica morphology is somewhat similar to an aggregated structure of uniformly-sized globular silica (a corpuscular system) [28,29], and different from the smooth cylindrical structures forming a network (a spongy system) that was observed for a silica monolith having a larger-sized skeletons and through-pores, prepared in a mould. The morphology is known to be affected by the relative onset of gelation and phase-separation that must be controlled to minimize the shrinkage. Figure 5.2 also shows that the attachment of the silica skeleton to the capillary wall prevented the formation of large voids along the wall. At each end of the tube, however, the bonding of the silica to the wall could not completely prevent the shrinkage, resulting in voids.

Monolithic columns

171

iiiiiiiii!iiiiiiiiiiiiiiii!!! ,.:.iiiii!ii!!iiiiiiii[!iiiiiii!i!i!ili! .......i.:i............ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . (a) '~ ~ .~.:~i~ii~iJiJi:~!!~i!~!!!i!!~!!~i~!!~i~i~~ !~!~!i:--: i!JJ~!!~i .~~:~ii~i~!i~ii~i~i~;~;~.~ 9~.~i i'~:~i:~;ii ~ i~Y~iii!ii!'~ii~~iiiiii}~..,.~,,~ :~il ~:~iiii~;:,d:~":~ :

' ~:~!!~ i'~~;~iii i i i i!i i i !i!!ii i !i{i ',!iil}:i~:,,,~ii 'i,iii ili!!ii~i i!~;ii{i i!i

9!ii:!:i:!iii iii:iiii

:iiiiiiiiii:iii!iiiiiiiiiiiiiii!!!!!!!!!!!iiil)!!!!!! ii!iii!:iii!i!!!i!iiii:iiiiiiiiiiiii!!!!iiiiii ?i (b) Fig. 5.2. (a), Scanning electron micrograph of a continuous monolithic silica prepared in a fused silica capillary (100 ~m) [16]; (b), SEM photograph of monolithic silica prepared in a capillary (75 ~m). Reproduced from ref. 19, with permission.

The ratio between the through-pore size (ca. 8 ~m) and the skeleton size (ca. 2.2 ~tm) shown in Fig. 5.2a is much greater than in a packed bed of a particle-filled column. Figure 5.3 shows the plots of skeleton size against the through-pore size in a column for a silica monolith prepared in a capillary or in a mould, as well as in a particle-packed column. The through-pore size/skeleton size ratios observed with the References pp. 180-181

Chapter 5

172

E cO

nm

1 41 2

m

-H

9 O

m H

3

i---! i----!

5 6

0

I

I

I

I

I

I

I

I

1

2

3

4

5

6

7

8

dpore [ Fig. 5.3. Plots of the skeleton size against the through-pore size of the continuous monolithic silica prepared in a capillary ( 9 and the larger-sized silica rod columns (7 mm x 83 mm) having constant through-pore size/skeleton size ratio (m) [15]. Also plotted are the particle size (vertical axis) against the size of interstitial voids (25-40% of dp as indicated by the bars) found with a conventional particle-packed column.

monolithic silica capillary are 3-5, the plots being located close to the upper end of Fig. 5.3, representing an open-tube column. The size of interstitial voids in a conventional particle-packed column is reported to be 25-40 % of the particle size [29], as indicated by the bars in Fig. 5.3. The through-pore-size/skeleton-size ratios of the monolithic silica columns prepared in a mould were 1.0-1.5, as a result of the shrinkage of the silica network structure during preparation [15]. The large through-pore size/skeleton size ratios in the monolithic silica in capillary were provided by the lack of shrinkage of the whole silica structure. The amount of silica portion in the preparation mixture was similar for the monoliths in a capillary or in a mould. The specific surface area of the silica monolith is similar to that of silica particles, at 300 m2/g. The silica monolith in a capillary possesses extremely high porosity, as shown in Table 5.1. Although the phase ratio is considerably smaller than a particle-packed column, it is much greater than a column packed with non-porous particles.

173

Monolithic columns

TABLE 5.1 POROSITY OF SILICA MONOLITH COLUMNS [17]

Porositya Column type

Particle packedb Monolith (PTFE) c

Monolit~ (PEEK) '~

Monolith (FS) e

Column diameter 4.6 mm

7 mm

4.6 mm

100 ~m

Column length

15 cm

8.3 cm

10 cm

25 cm f

Total porosity

0.78 (0.66)

0.86 (0.80)

(0.87)

0.96 (0.90)

Through-poreg

0.39 (0.39)

0.62 (0.63)

(0.69)

0.86 (0.85)

Mesopore

0.40 (0.27)

0.24 (0.18)

(0.18)

0.10 (0.05)

a The porosities in parentheses were obtained with a C18-bonded phase b Develosil-C18 particles packed in a column, 4.6 mm diameter, 10 cm in length c Monolithic silica column prepared in a mould and inserted into a PTFE tube d Monolithic silica column prepared in a mould and PEEK-resin clad e Monolithic silica column prepared in a 100 ~tm capillary f Effective length between the inlet and the detection window. Total length, 33.5 cm g External porosity

5.5 P E R F O R M A N C E OF M O N O L I T H I C SILICA C O L U M N S IN CEC

Figure 5.4a shows the electro-driven elution of alkylbenzenes and polyaromatic hydrocarbons (PAHs) in 80% acetonitrile. Figure 5.4a shows that the 250 mm column (100 ~tm diameter) produced ca. 40,000 and 35,000 theoretical plates for thiourea and hexylbenzene (k value = 0.8), respectively, at 1.1 mm/s linear velocity. Because of the presence of the very large through-pores at 8-10 ~tm, low column efficiency would be expected in pressure-driven elution, and this was actually the case. Interstitial voids of that size would be found in a column packed with ca. 30 ~tm particles. Figure 5.4b shows that the same column produced 5,000 and 6,000 theoretical plates for thiourea and hexylbenzene, respectively, under the pressure-driven conditions, with a split injection HPLC system at a similar linear velocity. The monolithic silica showed much higher performance in CEC than in the pressure-driven mode.

References pp. 180-181

Fig. 5.4. Chromatograms obtained for alkylbenzenes polyaromatic hydrocarbons [PAHs (b), (41 in electro-driven [CEC (a), (b)] elution, and in pressure-driven conditions [HPLC (c), (d)] [17]. PAHs: N, naphthalene; F, fluorene; Ph, vhenanthrene: A, anthracene; P, pyrene; T, triphenylene; B,: benzo[a]py rene. o m n size: I 00 t n I.D. x 33.5 cm (effective length 25 cm). Mobile phase: (a), (b) acetonitrile-Tris. HC150 mM, pH 8 (80:20); (c), (d), 80% acetonitrile-water. Applied voltage, (a), (b), 900VIcm. Pressure (c), (d), 0.13 MPa.

CEC

HPLC

(

(b)

A

Monolithic columns

175

The Van Deemter plots obtained with thiourea and a retained solute (hexylbenzene) in 80% and 90 % acetonitrile with a 50 gm capillary are shown in Figure 5.5. The lower plate-height and a slight decrease in plate-height with the increase in the linear velocity were observed in CEC, with an Hmin of ca. 4.5 lam at the highest linear velocity, ca. 2 mm/s, with an applied electric field of 30 kV across the 33.5 cm silica monolith (25 cm effective length). The maximum efficiency, 56,000 theoretical plates for hexylbenzene, is remarkable for a column that can be eluted at less than 0.2 MPa in a pressure-driven mode to produce similar linear velocities. The high permeability is advantageous, because the replacement of mobile phase, and HPLC--CEC hybrid operation (or pressure-assisted CEC), would be facilitated. In pressure-driven operation, considerable band broadening was observed at high linear velocity, although the separation impedance was much lower than that of a particle-packed column owing to the much lower flow resistance. The separation impedance (E = AP tO / qN 2 = (AP / N) (tO / N) (1/q)) expresses the total column performance in terms of the reciprocal number of theoretical plates per unit time and pressure drop. Because the contributions of the B- and C-terms are expected to be similar for a pressure-driven mode and an electro-driven mode, the difference in performance can be attributed to the greater contribution of the A-terms in Eqn. 5.2 in the pressure-driven mode. The contribution of the A-term is known to be less in CEC than in HPLC [6]. Monolithic silica columns in capillaries can provide good performance in CEC, because the effect of large pores can be minimized in the presence of a plug-type flow. In this sense, CEC seems to be an attractive way of using monolithic silica columns in capillaries that possess excessively large through-pores. In addition it is free from problems of bubbles and back-pressure. The large through-pores provide high permeability at the expense of the increased A-term contribution to band-broadening in HPLC. In CEC, the contribution is less than in HPLC, although the column efficiency is still poorer than expected from a consideration of the silica skeleton size. The performance of the monolithic silica in a capillary seems to be dominated by the size of the large through-pores and slow EOF. In the van Deemter plot for opentube capillary chromatography, a plate height (H) partly depends on the square of dc as in Eqn. 5.4 (Ds, diffusion coefficient in the stationary phase; dc, inner diameter of the capillary; dj; thickness of the stationary layer) [30]. This also explains the reduction in the number of theoretical plates with a solute having the longer retention.

H = 2 D m + l + 6 k + l l k 2 9~~ u + ~2k u u 96(1 + k) 2 D~ 3(1 + k) 2 D~

References pp. 180-181

(5.4)

Chapter 5

176

30

E z:l.

(a)

25

9

9

0 20 c'-0") .....

0

0

15 "0

(D "1" 10 ,.4--J

i

I

I

I

I

1

2

3

4

Linear Velocity, u, mm/s 30

E

::L

(b)

25 20

c-" 03 (D

"1(D

15 - 0

00 10

0

0 9

0

,..l.-a

0

0

00o i

i

i

i

1

2

3

4

5

Linear Velocity, u, mm/s

Fig. 5.5. Van Deemter plots obtained for C18 monolithic silica in a capillary in CEC (open symbols) and HPLC (solid symbols) with thiourea (A) and hexylbenzene (O,O) as a solute. Mobile phase: acetonitrile-water (HPLC), acetonitrile-Tris.HC1 buffer, 50 mM pH 8: (CEC), (a), 80:20; (b), 90:10. Column size: 50 ~tm I.D. x 33.5 cm (effective length 25 cm).

177

Monolithic columns

20

E ::3.

15-

::!::

,,,...

c--

t~

0 0 <>0

lo

0

5

0

I

I

I

0,5

1

1.5

k Fig. 5.6. Plots of plate heights against k values obtained in CEC with alkyl-benzenes (O,O) and PAHs ( ~ , O ) as solute. Mobile phase: acetonitrile-Tris.HC1 (buffer 50 ~ pH 8) 90:10 (solid symbols) and 80:20 (open symbols). Column size: 100 ~m I.D. x 33.5 cm (effective length 25 cm). Plate heights observed at around u= 1 mm/s were plotted.

Figure 5.6 shows the plots of plate height against k values for the present monolithic silica column in CEC. The results seem to show the contribution of the through-pores as large as 10 lam in the monolithic silica capillary column. The through-pore size seems to be too large even for CEC application. Such a contribution is much less for a particle-packed column that possesses much smaller through-poresize / skeleton-size ratios. Knox recently reported the dominant contribution of the slow mass-transfer in the mobile phase to band broadening in LC [31 ]. Table 5.2 lists the efficiency of various monolithic columns. The generation of 100,000-200,000 plates, or even greater, with 25-40 cm silica-packed columns has been reported. The results cannot be compared rigorously, because the column efficiency is affected by many factors including the column length, the composition and linear velocity of the mobile phase, the temperature, the electric-field gradient, and the solute retention. Other practical advantages associated with monoliths should also be taken into account. The results show the utility of monolithic columns, although most of the monoliths reported so far showed lower performance than a column packed References pp. 180-181

Chapter 5

178

TABLE 5.2 EFFICIENCY OF HIGH-PERFORMANCE COLUMNS

Mode

Medium

CEC

Silica particles bonded

7

Silica particles sintered

6

HPLC

Particle size (~tm)

L (cm)

Ref.

66,000

30

10

40,000

23

9

Na

Silica monolithb

2.2

56,000

25

27

Silica monolith

u

28,000

25

19

Silica-coated open tube

w

115,000

30

34

Polymer monolith c

w

80,000

20.5

32

Polymer monolith d

~

19,000

Polymer monolithe

~

150,000

120

33

Polymer monolithf

~

55,000

27

36

Silica monolith g

1.8

12,000

Silica monolithb

2.2

100,000

12.5

8.3 130

35

15 25

a Number of theoretical plates b FS-type silica monolith (50 ~m FS tube) with 2.2 ~tm skeletons and 8 ~tm through-pores c Prepared from acrylamide, butyl acrylate, acrylic acid, and bis-acrylamide d Prepared from methacrylamide and piperazine diacrylamide e Prepared from butyl methacrylate, ethylene dimethacrylate, and 2-acrylamido-2-methyl-1propanesulfonic acid f Prepared from vinylbenzyl chloride and divinylbenzene followed by reaction with N,N-dioctylamine g PTFE-type silica monolith with 1.7 ~tm skeletons and 2.2 ~tm through-pores

with 1-2 ~m silica particles. Consolidation of a packed bed has been examined successfully to increase the stability by sintering or agglomerating silica particles [7-12]. Monolithic columns prepared by forming a silica-gel matrix from a sol around ODS particles packed in a capillary gave efficiencies of up to 220,000 plates/m in CEC [10]. Loose packing of particles (external porosity of greater than 50%) [11 ], which is possible with a monolithic structure, can lead to high permeability leading to easier operation, a column

Monolithic columns

179

wash, the solvent change, or the equilibration. Polymer monoliths showed efficiencies up to 400,000 plates/m or 80,000 plates per column in CEC [32]. Svec and his co-workers reported that a polymer monolith of 120 cm produced 150,000 plates in CEC [33]. For CEC, organic-polymer-based monolithic columns gave high efficiencies, partly because they can be designed to generate high EOF. Because of the high chemical and mechanical stability, and easy design of surface functionalities, the development of polymer monoliths can be the key for the progress of CEC separations. Monolithic silica columns can also be improved to give greater EOF and greater chemical stability, and at the same time to have a smaller domain size, leading to a smaller A-term contribution. Actually, the results with a silica monolith in a 50 ~tm capillary were much better and more reproducible than those with 100 ~tm capillary monoliths [27]. While a 25 cm long capillary column produced 12,000 theoretical plates in HPLC in 90% acetonitrile, it produced 56,000 theoretical plates in CEC. The monoliths prepared in a 50 ktm capillary showed much higher mechanical stability than those prepared in a 100 ~tm capillary, presumably owing to the better attachment to the wall, based on the greater surface area/volume ratio of the smaller-sized capillary tubes. Monoliths with low surface coverage by C18 groups showed particularly high performance with a high EOF velocity, although solute retention was small, resulting in a k value of about 0.2 for hexylbenzene. The column efficiency of 80,000 plates can be obtained for an unretained peak with a 25 cm column [27], although a 50 cm column produced only 100,000 plates, owing to the limitations in the EOF velocity. The capillary monolith with a high coverage also showed the lower performance, because of the lower EOF velocity. This shows a problem of a monolithic silica column in a capillary for CEC application. The slow electro-osmotic flow is presumably a consequence of the high purity of silica materials. The addition of metal impurities will increase the ionic sites, and result in a greater EOF. This approach, however, is opposite to the direction of the development of stationary phases for RPLC in the past, because it would increase the secondary retention effects. The major advantage of the monolithic silica structure is the high permeability. Even if the application of silica monoliths to CEC is accompanied by difficulties, high permeability can contribute to high performance in pressure-driven or CEC/HPLC-hybrid operation. Hyphenation to a mass spectrometric detector would be much easier for HPLC than for CEC. High column efficiency under modest pressure was observed in the HPLC mode, owing to the small-sized skeletons and large through-pores. With their extremely high permeability, long silica monoliths may be able to provide the desired column efficiency in a pressure-driven mode. Actually, a monolithic silica in a capillary (130 cm) generated 100,000 plates per column under 0.4 MPa [25,27]. The preparation procedure needs further improvement to produce the smaller-sized

References pp. 180-181

180

Chapter 5

through-pores and to prevent gel-shrinkage in order to obtain high-performance monolithic silica capillaries. The monolithic columns can overcome some of the limitations associated with particle-packed columns. Features of monolithic columns in CEC include the facts that there is: (i), no need for a frit, no bubble problems, and high bed-stability based on the single-piece structure; (ii), the high efficiency based on the small skeleton size leads to fast internal diffusion (especially for polymer gels); (iii), the preparation of a long column is easy; (iv) there is easy operation based on high permeability (with a fast wash and equilibration with new mobile phase under pressure-driven conditions); and (v), with polymer monolith columns there can be easy chemical design for fast EOF, with the utility of swollen polymer gels. Another great benefit of monolithic columns is that it is easy to prepare a stable, high efficiency column to examine various monomers or three-dimensional structures. Polymer and silica monoliths from monomers that can be studied by many researchers have a good chance of being successful as popular separation media for a wide range of applications. 5.6 R E F E R E N C E S

1 J.C. Giddings, Dynamics of Chromatography, Part 1, Principles and Theory, Marcel Dekker, New York, 1965. 2 P.A. Bristow, J.H. Knox, Chromatographia, 10 (1977) 279. 3 H. Poppe, J. Chromatogr. A, 778 (1997) 3. 4 J.H. Knox, I.H. Grant, Chromatographia, 24 (1987) 135. 5 S. Ludtke, T. Adam, K.K. Unger, J. Chromatogr. A, 786 (1997) 229. 6 M.M. Dittmann, G.P. Rozing, J. Chromatogr. A, 744 (1996) 63. 7 M.T. Dulay, R.P. Kulkarni, R.N. Zare, Anal. Chem. 70 (1998) 5103. 8 C.K. Ratnayake, C.S. Oh, M.P. Henry, J. High Resolut. Chromatogr., 23 (2000) 81. 9 R. Asiaie, X. Huang, D. Farnan, C. Horvath, J. Chromatogr. A, 806 (1998) 251. 10 Q. Tang, M.L. Lee, J. High Resolut. Chromatogr., 23 (2000) 73. 11 Q. Tang, B. Xin, M.L. Lee, J. Chromatogr. A, 837 (1999) 35. 12 Y. Adam, K.K. Unger, M.M. Dittmann, G.P. Rozing, J. Chromatogr. A, 887 (2000) 327. 13 H. Minakuchi, K. Nakanishi, N. Soga, N. Ishizuka, N. Tanaka, Anal. Chem. 68 (1996) 3498. 14 H. Minakuchi, K. Nakanishi, N. Soga, N. Ishizuka, N. Tanaka, J. Chromatogr. A 762 (1997) 135. 15, H. Minakuchi, K. Nakanishi, N. Soga, N. Ishizuka, N. Tanaka, J. Chromatogr. A 797 (1998) 121. 16 H. Minakuchi, N. Ishizuka, K. Nakanishi, N. Soga, N. Tanaka, J. Chromatogr. A 828 (1998) 83. 17 N. Ishizuka, H. Minakuchi, K. Nakanishi, N. Soga, H. Nagayama, K. Hosoya, N. Tanaka, Anal. Chem., 72 (2000) 1275.

Monolithic columns

181

18 N. Tanaka, H. Nagayama, H. Kobayashi, T. Ikegami, K. Hosoya, N. Ishizuka, H. Minakuchi, K. Nakanishi, K. Cabrera, D. Lubda, J. High Resolut. Chromatogr., 23 (2000) 111. 19 C. Fujimoto, J. High Resolut. Chromatogr., 23 (2000) 89. 20 K. Nakanishi, J. Porous Mater., 4 (1997) 67. 21 K. Nakanishi, N. Soga, J. Am. Ceram. Soc., 74 (1991) 2518. 22 K. Nakanishi, N. Soga, J. Non-Cryst. Solids, 139 (1992) 1 and 14. 23 S.M. Fields, Anal. Chem., 68 (1996) 2709. 24 N. Ishizuka, H. Minakuchi, K. Nakanishi, N. Soga, K. Hosoya, N. Tanaka, J. High. Resolut. Chromatogr., 21 (1998) 477. 25 H. Kobayashi, H. Nagayama, T. Ikegami, K. Hosoya, N. Tanaka, N. Ishizuka, H. Minakuchi, K. Nakanishi, International Symposium on Advances in Chromatographic and Electrophoretic Separations, Bayreuth, April 2000. 26 K. Nakanishi, H. Shikata, N. Ishizuka, N. Koheiya, N. Soga, J. High Resolut. Chromatogr., 23 (2000) 106. 27 N. Tanaka, H. Kobayashi, H. Nagayama, T. Ikegami, K. Hosoya, N. Ishizuka, H. Minakuchi, K. Nakanishi, 24th International Symposium on High Performance Liquid Phase Separations and Related Techniques, Seattle, June, 2000. 28 A.P. Kamaukhov, in S. Modry (Editor), Proceedings of Rilem/IUPAC International Symposium on Pore Structure and Properties of Materials, Vol. I, Academia, Prague, 1974. 29 K.K. Unger, Porous Silica; Elsevier, Amsterdam, 1979. 30 P.P.H. Tock, C. Boshoven, H. Poppe, J.C. Kraak, J. Chromatogr., 477 (1989) 95. 31 J.H. Knox, J. Chromatogr. A, 831 (1999) 3. 32 A. Palm, M.V. Novotny, Anal. Chem., 69 (1997) 4499. 33 E.C. Peters, M. Petro, F. Svec, J.M.J. Frechet, Anal. Chem., 70 (1998) 2296. 34 A.L. Crego, J. Martinez, M.L. Marina, J. Chromatogr. A, 869 (2000) 329. 35 A. Maruska, C. Ericson, A. Vegvari, S. Hjerten, J. Chromatogr. A, 837 (1999) 25. 36 I. Gusev, X. Huang, C. Horvath, J. Chromatogr. A, 855 (1999) 273.

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Chapter 6

Capillary Column Technology: Continuous Polymer Monoliths Frantigek SVEC

Department of Chemistry, University of California, Berkeley, CA 94720-1460, USA

CONTENTS

6.1 6.2 6.3 6.4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Replaceable polymeric stationary phases . . . . . . . . . . . . . . . . . Polymer gels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Highly crosslinked acrylamide-based monoliths . . . . . . . . . . . . . . 6.4.1 Polymerization in aqueous solutions . . . . . . . . . . . . . . . . 6.4.2 Polymerization in organic solvents . . . . . . . . . . . . . . . . 6.5 Imprinted enantioselective monoliths . . . . . . . . . . . . . . . . . . . 6.6 Polystyrene-based monoliths . . . . . . . . . . . . . . . . . . . . . . . . 6.7 Methacrylate ester-based monolithic columns . . . . . . . . . . . . . . . 6.8 Assessment of porous structure . . . . . . . . . . . . . . . . . . . . . . 6.8.1 Control of porous properties . . . . . . . . . . . . . . . . . . . . 6.9 Effects of properties on the separation ability . . . . . . . . . . . . . . . 6.9.1 Surface chemistry . . . . . . . . . . . . . . . . . . . . . . . . . 6.9.2 Pore size . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.9.3 Flow of the mobile phase . . . . . . . . . . . . . . . . . . . . . 6.10 Other applications of porous polymer monoliths in CEC column technology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.10.1 Immobilization of beads in porous polymer monolith . . . . . . 6.10.2 Monolithic frits . . . . . . . . . . . . . . . . . . . . . . . . . . 6.11 A c k n o w l e d g m e n t . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.12 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

184 185 187 189 189 200 206 207 212 220 221 223 225 228 230 232 232 234 237 238

184

Chapter 6

6.1 INTRODUCTION CEC is often presented as a hybrid method that combines the capillary column format and electroosmotic flow typical of capillary electrophoresis (CE) with the use of a solid stationary phase and a separation mechanism characteristic of HPLC based on specific interactions of solutes with a stationary phase. Therefore CEC is most commonly implemented by means typical of both HPLC (packed columns) and CE (use of electrophoretic instrumentation). As described in another chapter of this book, both commercial columns and instrumentation manufactured specifically for CEC remain scarce. Although numerous groups around the world prepare CEC columns using a variety of approaches, the vast majority of these efforts described in the previous chapter mimic in one way or another standard HPLC column technology. However, some aspects of this technology have proven difficult to implement on the capillary scale. Additionally, the stationary phases packed in CEC capillaries were at the very beginning standard commercial HPLC-grade beads. Since these media were tailored for regular HPLC modes, and their surface chemistries were optimized accordingly, their use incorrectly treated CEC as a subset of HPLC. Truly optimized, however, CEC packings should play a dual role: in addition to providing sites for the required interactions (as in HPLC), they must also be involved in electroosmotic flow. As a result, packings that are excellent for HPLC may offer limited performance in the CEC mode. This realization of the basic differences between HPLC and CEC [ 1] has stimulated the development of both specific particulate packings having properties tuned for the needs of CEC as well as alternative column technologies. The technical difficulties associated with packed columns have spurred the development of a number of different approaches. For example, one of these competing technologies - in situ polymerized organic separation media - was adopted from a concept developed for much larger diameter HPLC columns. Historically, Kubin et al. were most likely the first who published the preparation of a continuous polymer matrix for chromatography while attempting to replace natural polysaccharide gel beads with a highly swollen poly(2-hydroxyethyl methacrylate) gel for the low pressure size-exclusion chromatography of proteins [2]. However, the permeability of this continuous bed gel was far too low to make the material useful. The first operating monolithic columns were prepared from open-pore polyurethane foams in the early 1970s and used as stationary phases for both liquid and gas chromatography. However, these materials were found to suffer from excessive swelling and softening in some solvents and the technology has almost been forgotten [3-6].

Monoliths

185

Successful approaches towards continuous media for "classical" liquid chromatography emerged only in the late 1980s, and included macroporous disks, rolled woven matrices, compressed poly(acrylamide) gels forming continuous bed, and rigid polymer monolithic columns. The genesis, properties, and applications of these novel separation media have recently been detailed in a series of excellent review articles [7-10]. Inorganic silica-based analogs of these columns were subsequently reported by several groups starting in 1996 [11,12]. As a result of their unique properties, these monolithic materials have recently attracted considerable attention from a number of different research groups. This popularity is also why monolithic CEC columns are the subject treated in several chapters in this book. Polymer based monolithic CEC columns were detailed in two our recent reviews that were modified and updated for the following text [13,14]. 6.2 R E P L A C E A B L E P O L Y M E R I C S T A T I O N A R Y PHASES

Although it may be argued that linear polymers do not belong to the category of organic monoliths, it should be noted, that as employed, these materials exist as physically entangled polymer chains that effectively resemble highly swollen, chemically crosslinked gels. Additionally, these materials effect separations in an identical manner, using EOF as the driving force. The use of capillaries filled with solutions of linear polymers is an interesting column technology originating directly from the field of electrophoresis. In contrast to the polymer gels that will be discussed later, the preparation of these pseudostationary phases needs not to be performed within the confines of the capillary. Several research groups have already reported the use of specifically designed copolymers [15-22] and modified dendrimers [23,24] containing both charged and hydrophobic moieties as alternatives to micelles in micellar electrokinetic chromatography (MEKC) for the separation of neutral analytes. Fujimoto et al. [ 16] realized, that in contrast to polyacrylamide homopolymers, the migration of neutral compounds through a capillary column would be achieved more easily if charged functionalities were incorporated into the neutral polyacrylamide chains. Indeed, acetone and acetophenone may be baseline separated using a 10% aqueous solution of an acrylamide copolymer containing 2.4% 2-acrylamido-2methyl-1-propanesulfonic acid. However, nearly 100 min were required to effect this separation at a field strength of 260 V/cm, making this approach impractical even with the use of higher voltage. Alternatively, Tanaka et al. [17] alkylated commercial polyallylamine with C8-C16 alkyl bromides, followed by a Michael reaction with methyl acrylate and subsequent hydrolysis of the methyl ester to obtain free carboxyl functionalities. Although difficult to prepare, this polymer effected efficient separations of ketones and aromatic References pp. 238-240

186

Chapter 6

hydrocarbons in less than 20 min at 400 V/cm. Similarly, Kenndler's group (Poto~ek et al. [25]) has demonstrated the separation of phenols using a partially hydrolyzed polyacrylamide solution.

12H2s 1

2

3

Schure et al. [22] published an extensive study employing a pseudostationary phase prepared by a modified emulsion polymerization of methacrylic acid 1, ethyl acrylate 2, and dodecyl methacrylate 3. He also related the physical properties of the polymer solution to the quality of CEC separation obtained. For example, increasing the concentration of the linear polymer solution increased the number of interacting moieties, improving the efficiency to a maximum of 293 000 plates/m in a 3.72% polymer solution. Increasing the polymer concentration also led to an increase in the number of carboxyl functionalities available in the capillary column, thus boosting electroosmotic flow and reducing the time required for analysis. Rheological measurements covering the range of polymer concentrations from 0 to 4% (upper solubility limit) indicated that the dissolved pseudostationary phase afforded the best separation for concentrations at which the viscosity of the solution was the highest, and the polymer chains were most entangled. These soluble polymers were shown to effect the efficient separations of alkyl benzoates, aromatic ketones (Fig. 6.1), alkylbenzenes, and polyaromatic hydrocarbons. Columns filled with polymer solutions are extremely simple to prepare, and the "packing" can easily be replaced as often as desired. These characteristics make the pseudostationary phases excellent candidates for use in routine CEC separations such as quality control applications where analysis and sample profiles do not change much. However, several limitations limit their widespread use. For example, sample capacity is typically very low, pushing standard detection methods close to their sensitivity limits. Additionally, migration of the pseudostationary phase itself represents a serious problem for separations utilizing mass spectrometric detection well known from micellar electrokinetic chromatography. However, the most significant limitation involves the solubility profiles of the polymers. It is generally recognized [17,22] that these amphiphilic polymers are rarely soluble in the broad range of organic solvent/water mixtures typically used for reversed-phase separations, a shortcoming that greatly reduces their operational scope.

187

Monoliths

r r~

r

r

0

I

I

2

4

I

6 min

Fig. 6.1. Separation of alkylphenones using replaceable polymer solution as a separation medium (Reprinted with permission from [22]. Copyright 1997 American Chemical Society). Conditions: Column 50 cm (42 cm active) x 50 ~tm i.d., 4.00% linear polymer retentive phase, mobile phase 40% acetonitrile in buffer pH 9.1; voltage, 30 kV; injection, 5 kV for 15 s. Peaks: acetophenone (1), butyrophenone (2), hexanophenone (3), and octanophenone (4).

6.3 P O L Y M E R GELS The first monolithic CEC columns contained swollen hydrophilic polyacrylamide gel, mimicking those used for capillary gel electrophoresis [26]. Typically, the capillary was filled with an aqueous polymerization mixture containing monovinyl and divinyl (crosslinking) acrylamide-based monomers as well as a free radical redox initiating

system,

such

as

ammonium

peroxodisulfate

and

N,N,N',N'-tetra-

methylethylenediamine (TEMED). Since initiation of the polymerization process begins immediately upon mixing of all the components at room temperature, the reaction mixture must be used immediately. The polymerization process is allowed to proceed overnight to afford capillaries filled with continuous gel beds. It should be noted that these gels are loose, highly swollen materials that may contain no more than 5% solid polymer.

References pp. 238-240

188

Chapter 6

4

5

6

For example, Fujimoto et al. [ 16] polymerized an aqueous solution of acrylamide 4, methylenebisacrylamide fi, and 2-acrylamido-2-methyl-l-propanesulfonic acid 6 within the confines of a bare capillary. Despite the lack of chemical attachment to the inner wall of the capillary, these crosslinked gels showed fair physical stability. Although column efficiencies of up to 150 000 plates/m were observed for acetophenone, retention times on these columns were prohibitively long. This behavior was probably due, in part, to the relatively high background buffer concentration of 0.1 mol/L employed. This concentration is at least one order of magnitude higher than that typically used in current CEC studies. Based on his results, Fujimoto concluded that the prevailing mechanism of the separation was sieving rather than an interaction of the solutes with the matrix [ 15].

O~NH"~ O~OJ~/~Si(OCH3) 3 7

_8

Replacement of the hydrophilic acrylamide with the more hydrophobic N-isopropylacrylamide 7, in combination with the pre-functionalization of the capillary internal surface with 3-(trimethoxysilyl)propyl methacrylate 8, afforded a continuous gel covalently attached to the capillary. The electrochromatographic elution of hydrophobic analytes from this column required the use of aqueous buffer/acetonitrile mixtures [27]. In contrast to the previously described work, improvements in the separations were observed using these "fritless" hydrogel columns. For example, Fig. 6.2 shows the successful separation of a series of aromatic compounds using the soft gel column. These improvements consist of both shorter retention times and column efficiencies as high as 160,000 for various steroids [27]. The separations ofhydrophobic compounds obtained using this polymer gel stationary phase exhibit many of the attributes typical of reversed-phase chromatography, including a linear dependence of the retention factor k' on the composition of the mobile phase. This led Fujimoto to

189

Monoliths

3

4

0I

5

10

15 min

Fig. 6.2. Electrochromatographic separation of benzyl alcohol (1), resorcinol (2), methylparaben (3), and 13-naphthol (4) using a soft gel column (Reprinted with permission from [27]. Copyright 1998 Wiley-VCH). Conditions: Column 48.5 cm (24 cm active) x 75 lam i.d., stationary phase 4.1% T, 9.7% C, 0.7% S poly(2-acrylamido-2-methyl-1propanesulfonic acid-co-N-isopropyl acrylamide-co-methylene bisacrylamide); mobile phase 20:80 acetonitrile and 2.5 mol/L phosphate buffer pH 6.8; 16 kV.

the conclusion that, in contrast to his original polyacrylamide-based gels from 1995, size-exclusion was no longer the primary mode of separation. 6.4 HIGHLY CROSSLINKED ACRYLAMIDE-BASED MONOLITHS 6.4.1 Polymerization in aqueous solutions

Another approach towards continuous CEC beds involved highly crosslinked acrylamide polymers and was reported by Hjert6n et al. [28] at about the same time as Fujimoto's gels. An outstanding review [9] describes this development in detail. The original approach required a multiplicity of steps including the modification of the capillary surface with 3-(trimethoxysilyl) propyl methacrylate 7, two polymerization

References pp. 238-240

Chapter 6

190

O•OI•/OH O

~

9

O

10

11

12

steps, and a chemical functionalization [29]. The initial polymer matrix was formed by copolymerizing a dilute aqueous solution of 2-hydroxyethyl methacrylate 9 and piperazine diacrylate 10 using a standard redox system in the presence of a high concentration of ammonium sulfate. The pores of this matrix were then filled with another polymerization mixture containing allyl glycidyl ether 11 and dextran sulfate, and the second polymerization proceeded within the pores of the initial matrix leading to the "immobilization" of the charged dextran within the newly formed composite. Eventually, reaction of both epoxide and hydroxyl functionalities with 1,2-epoxyoctadecane 12 led to the covalent functionalization of the matrix with a number of C18 chains. Several chromatographic measurements were performed using these capillaries, with retention times in excess of 20 min being monitored for the elution of aromatic hydrocarbons [29].

~~'SO3H

13

o~o~ 14

In order to simplify the previous preparation method, a more straightforward procedure was later developed by the same group [30]. The polymerization mixture consisted of an aqueous solution of acrylamide 4, piperazine diacrylamide 10, and vinylsulfonic acid 13 with added stearyl methacrylate 3_ or butyl methacrylate 14 to control the hydrophobicity of the gel. Since neither of these non-polar monomers is soluble in water, a surfactant was added to the mixture, followed by sonication to form an emulsion of the hydrophobic monomer in the aqueous solution. Once initiated, the mixture was immediately drawn into an acryloylsilanized capillary, where the polymerization was completed. The presence of the strongly acidic sulfonic acid functionalities afforded EOF that remained constant over a broad pH range.

Monoliths

191

Although the initial separations performed using these continuous gel beds were good, an ingenious trick involving changes in the strength of the mobile phase also enabled further improvements in the resolution of various polycyclic aromatic hydrocarbons. A solution containing the analytes dissolved in 50/50 acetonitrile/aqueous buffer was injected electrophoretically into the capillary column, and the separation was started. After a short period of time, the solvent in the inlet vial was replaced with a 70/30 acetonitrile/buffer mixture, and the elution was completed under these conditions, leading to peaks sharper than those obtained using standard isocratic elution. The authors assumed that this improvement resulted from the gradient of the mobile phase generated by diffusion across the strong solvent/weak solvent interface. Addition of sodium dodecyl sulfate at levels below the critical micelle concentration was also reported to improve isocratic CEC separations [30]. Fig. 6.3 shows the remarkable difference between two separations of aromatic hydrocarbons in the presence and in the absence of sodium dodecylsulfate. The same group very recently described another interesting method of preparation of a monolithic capillary column that was used for CEC gradient separation of proteins [31]. The first step involved a polymerization initiated by ammonium persulfate/TEMED in a two-phase system: aqueous phase, consisting of a solution of acrylamide 4 and piperazine diacrylamide 10 in a mixture of a buffer solution and dimethylformamide, and immiscible highly hydrophobic phase containing octadecyl methacrylate. Continuous sonication was applied for 40 min in order to emulsify the hydrophobic monomer and form a dispersion of fine polymer particles. In the remaining steps of this preparation process, another portion of initiator was added to the system to restart the polymerization of two newly added monomers, dimethyldiallylammonium chloride .1.$. and piperazine diacrylamide 10. The resulting partly polymerized dispersion was then forced into a methacryloylsilylated capillary using pressure and, finally, the polymerization process was carried out to completion.

12

Understanding and controlling the forces that drive the movement of charged molecules during CEC separations is extremely important. Proteins may be particularly difficult to separate, since, depending on their net charge as determined by both

References pp. 238-240

Chapter 6

192

1

== 3 4

5

10

ZO

rain

i 0

ZO

rain

Fig. 6.3. Effect of sodium dodecylsulfate in the mobile phase upon electrochromatography of polycyclic aromatic hydrocarbons on a C18-derivatized continuous bed containing sulfonic acid groups (Reprinted with permission from [30]. Copyright 1996 American Chemical Society). Capillary: 14 cm (10 cm effective length) x 100 ~tm i.d. Applied voltage: 3.0 kV Eluent: 60% (v/v) acetonitrile in 4 mmol/1 sodium phosphate (pH 7.4). (a) 1.0 mmol/1 SDS added to the eluent; (b) without addition of SDS (control). Peaks: naphthalene (1), 2-methylnaphthalene (2), fluorene (3), phenanthrene (4), anthracene (5).

the protein pI and the pH value of the mobile phase, the same molecule can move electrophoretically towards either the anode or the cathode. In contrast, EOF proceeds in only one direction as determined by the charge of the mobile counter ions in the Stern layer. Thus, in Hjert6n's approach, using immobilized quaternary ammonium cations, EOF always proceeds from the anode to the cathode. He also used electroosmotic flow to introduce the mobile phase gradient (generated using an HPLC instrument to mix the solvents) into the capillary. Two overall flow scenarios shown schematically in Fig. 6.4 were discussed [31 ]:

193

Monoliths

Normal-Row Gradient (Driven by EOF) Hlgh-EOF Column

Counter-Flow Gradient (Driven by EOF) Moderate-EOF Column ! ..... I

.

.

.

.

.

.

.

.

.

.

1

.

I

I_.1~

"t'l~ i IG~

1~ I ~

Velph

_~....... ~,. r z j ~

i 8

T

T

T T

z

....

---

i

Q

!

!

'

!

x-

lID

inlet

outlet

inlet

oultet

a

(~= protein mr

Vmigr= IVeo-Veiphl Fig. 6.4. Diagram illustrating the principle for counterflow (a) and normal-flow (b) gradients (Reprinted with permission from [31]. Copyright 1999 American Chemical Society). The direction of electroosmotic flow is opposite to that of the electrophoretic movement in both methods and is opposite to the net migration velocity, Vmigr (= Veo - Velph), in (a) and coincides with the net migration direction in (b). Veo is not constant along the capillary, and Velph is higher in the direction of the electrophoretic migration.

(i) In the monolith containing a high level of charged moieties, electroosmotic flow outweighs the electrophoretic migration. In this "normal flow gradient" situation, both the EOF and the net migration of the protein molecules occur in the same direction, provided that all of the proteins have net charges of equal sign. This was achieved by using a mobile phase consisting of 80% acetonitrile and 20% buffer with a pH value of 2. This pH is well below the pI of the proteins, ensuring that the biopolymers were positively charged. Samples were injected at the cathode, and as with other typical gradient systems, the percentage of acetonitrile was higher at the

References pp. 238-240

Chapter 6

194

R

~CL

L

CL

C

Ch

Ch

Ch

ws

2 i

0

I

I

i

i

i

5 10 15 20 25 Time [mln] a

0

I

l

l

/

i

5 10 15 20 25 Time [mini b

0

5 10 15 20 25 Time [mini c

Fig. 6.5. Separation of proteins using gradient electrochromatography (a, b) and reversed-phase gHPLC (c) (Reprinted with permission from [31 ]. Copyright 1999 American Chemical Society). Peaks: ribonuclease A (R); cytochrome c (C); lysozyme (L); chymotrypsinogen (Ch). Protein concentration: 0.6 mg/mL of each protein except for ribonuclease A (1.8 mg/mL); mobile phase: a linear gradient from 5 to 80% acetonitrile in 5 mmol/L sodium phosphate pH 2.0; Detection at 280 nm. Columns: 8 cm (6 cm effective length) x 50 gm i.d.; voltage, 5.5 kV (700 V/cm); (a) Moderate-EOF column, (b) high-EOF column, (c) conventional capillary RP-HPLC; pressure 5 MPa.

capillary inlet (cathode) than at the outlet, with detection occurring at the anodic end of the column. (ii) In contrast, in the monolith containing a moderate number of charged moieties, electromigration of the charged proteins was faster than the electroosmotic flow. In this "counter flow gradient" system, the EOF proceeded in the same direction. However, proteins were injected at the anode, and migrated in a direction opposite to the gradient due to their electrophoretic mobilities. Accordingly, the detector was positioned at the cathodic end of the capillary column. Fig. 6.5 shows excellent separations of four proteins in both moderate (Fig. 6.5a) and high EOF columns (Fig. 6.5b) using electrically driven flow, and compares the separation with that achieved using standard HPLC methodology (Fig. 6.5c). Since the separations were primarily governed by the nature of the mobile phase gradient, all three chromatograms are very similar. This comparison also demonstrates that the separation is achieved via reversed-phase partitioning rather than electrophoresis. It is worth noting that this successful approach to monolithic CEC column for the separation of proteins employs a polymeric matrix containing positively charged

Monoliths

195

Fig. 6.6. Effect of ammonium sulfate concentration in the polymerization mixture on morphology of a monolithic CEC column prepared from a solution of piperazine diacrylate, N,N-dimethylacrylamide, and vinyl sulfonic acid (52% C, 29% T) in 20 mmol/L phosphate buffer pH 7. Ammonium sulfate concentration 0 (1), 20 (2), and 120 mg/mL (3). (Reprinted with permission of authors from [32]). moieties [31]. Since the overwhelming majority of proteins also possess net positive charges at pH=2, the danger of undesired electrostatic interactions is greatly diminished. The occurrence of very strong electrostatic interactions indicates why, to date, no successful attempts to separate proteins on columns containing carboxylic or sulfonic acid moieties at pH values below the pI were demonstrated. Hoegger and Freitag [32] also prepared acrylamide-based monoliths using polymerization in aqueous solutions. However, their typical polymerization mixture contained a much higher concentration of monomers (up to 29%) including piperazine diacrylamide 10 (52% in respect to total monomers), dimethylacrylamide 16, and 2-acrylamido-2-methyl-l-propanesulfonic acid 6 dissolved in an aqueous phosphate buffer pH 7. Porous properties of this monolithic polymer were controlled by addition of ammonium sulfate. Micrographs shown in Fig. 6.6 clearly demonstrate the changes in References pp. 238-240

Chapter 6

196

=

O

9

0

4

6

8

min

Fig. 6.7. Separation of a mixture of polar and nonpolar aromatic compounds by CEC. Conditions: capillary 27 cm x 75 ~tm i.d.For details on monolith column see Fig. 6.6; mobile phase: 1:3 methanol/acetonitrile, 30kV, 25~ injection 10 kV for 3s; UV- detection at 200 nm. (Reprinted with permission of authors from [32]). Peaks: pyrene 1, phenanthrene 2, anthracene 3, phenol 4, hydroquinone monomethylether 5, 2-naphtol 6, catechol 7, hydroquinone 8, resorcinol 9.

~ N I

I 16

morphology affected by addition of the inorganic salt. An optimized monolithic column enabled the separation of a number of aromatic compounds in the non-aqueous mobile phase consisting of methanol and acetonitrile with an efficiency of up to 80,000 plates/m (Fig. 6.7). The acrylamide-based monolith originally developed by Hjert6n for CEC capillary columns was very recently used by the same group to fill a 30.6 cm long 40 mm wide serpentine channel etched in flat quartz substrate shown in Fig. 6.8 [33]. The channel

197

Monoliths

.v Pow.,ie r ~ Detection Slit in Supporting

~,~,.

---,.-

|

Iiill

c,.o~, o~ Outlet Reservoir

~Alll

In!et Reservoir

O

|

c'~mp'cL ....

m m

Fig. 6.8. Experimental setup and scanning electron micrograph of morphology of the continuous bed prepared within the channel of a quartz chip. (Reprinted with permission from [33]. Copyright 2000 American Chemical Society).

walls were first functionalized by treating the surface with sodium hydroxide and hydrochloric acid followed by the reaction with [3-(methacryloyloxy) propyl] trimethoxysilane 8 in acetone at 150~ for 1 h. The channel was completely filled through an external reservoir at one end with the polymerization mixture and then pushed back by an aqueous poly(ethylene glycol) (MW 8,000) solution introduced from the other end to remove the polymerization mixture from part of the channel

References pp. 238-240

Chapter 6

198 TABLE 6.1

COMPOSITION OF POLYMERIZATION MIXTURES USED FOR THE PREPARATION OF MONOLITHIC SEPARATION MEDIUM WITHIN CHANNELS [33]

PDA a

MA b

IPA c

VSA d

(NH4)2SO4Buffer e

g

g

g

~tL

g

mL

Contin. bed 1

0.30

0.14

0.18

8

0.01

1.7

Contin. bed 2

0.30

0.14

0.26

70

0.18

2.5

apiperazine diacrylamide bMethacrylamide CN-isopropylacrylamide dVinylsulfonic acid esodium phosphate (50 mmol/L, pH 7.0). Initiating system consisting of 12 ~tL of 5% aqueous solution of N,N,N',N'-tetramethylethylenediamine and 12 ~tL of 10% aqueous solution of ammonium peroxodisulfate was admixed to each mixture.

laying between the detection window and the outlet. Two different polymerization mixtures shown in Table 6.1 were prepared and polymerized. N-isopropylacrylamide "7 is added to the polymerization mixture to increase hydrophobicity of the monolith required for the separations in reversed phase mode. Vinylsulfonic acid 13 provides the chargeable functionalities that afford electroosmotic flow. Since the gelation occurs rapidly already at the room temperature, the filling of the channel must proceed immediately after the complete polymerization mixture is prepared. The methacryloyl moieties attached to the wall copolymerize with the monomers in the liquid mixture. Therefore, the continuous bed fills the channel volume completely and does not shrink even after all solvents are removed. Fig. 6.8 also shows scanning electron micrograph of the dry monolithic structure that exhibits features typical of macroporous polymers [34]. Direct comparison of column efficiencies of the continuous separation medium prepared in 4.5 cm long both capillary and linear channel as measured using non retained marker acetone did not reveal any difference and an identical plate height of 4-5 ~tm was observed for both at flow velocities of 0.5-1.5 mm/s. This indicates that there is again no substantial difference between beds formed in both formats. Obviously, the channel behaves as a capillary although it does not have the circular cross section.

199

Monoliths

r~

r~

I,i

O_L.

0

........

~ 2

.... / 4 Time

I 6

| 8

...... i 10

[min]

Fig. 6.9. Electrochromatogram of two antidepressant drugs (amitriptyline 1, nortriptyline 2) and a related quaternary ammonium compound (methyl amitriptyline 3) obtained by isocratic elution in 30.6 cm long channel. Conditions: mobile phase, 5 mmol/L sodium phosphate, pH 2.5, containing 70% acetonitrile (v/v); flow rate of the mobile phase through the inlet reservoir, 50 p,L/min; applied voltage, 12 kV (400 V/cm); electrokinetic injection, 1 kV, 5 s; UV detection at 239 nm; sample concentration, 0.10-0.15 mg/mL. (Reprinted with permission from [33]. Copyright 2000 American Chemical Society).

Hjert6n's group achieved with this device very good CEC separations. Figure 6.9 shows as an example the separation of tricyclic antidepresant drugs. These basic compounds were well separated in narrow peaks within 8 min in the long serpentine channel using a buffer of pH 2.5 to reduce their ionization, and higher percentage of acetonitrile (70%) in the mobile phase. Similarly, using the same bed, acetone, aniline, and five phenones were baseline separated in 17 min. The most impressive result of this study is the separation of uracil, phenol, and benzyl alcohol monitored by a UV detector that is achieved in 17 s using 18 mm long channel filled with continuous bed containing a high concentration of vinylsulfonic acid (bed 2 in Table 6.1). In contrast to work concerned with the CEC separations in open-tubular chip formats that often suffer from both very low loading capacity requiring extremely sensitive laser induced

References pp. 238-240

Chapter 6

200

fluorescence detection and limited variability in surface chemistries, Hjert6n's study represents an important step towards analytical microchips and systems with channels containing genuine separation medium. This allows a precise tuning of the interactions, controlling EOF, and using more common and less sophisticated detectors.

6.4.2 Polymerizationin organicsolvents Despite the undeniable success, the use of purely aqueous-based polymerization systems for the preparation of monolithic capillaries for CEC discussed above also has some limitations. Perhaps the greatest limitation is that the typical nonpolar monomers such as butyl methacrylate 14 and stearyl methacrylate 3 that were used to achieve the necessary hydrophobicity for a reversed-phase CEC bed are insoluble in water. In contrast to the "fixed" solubilizing properties of water, the wealth of organic solvents possessing polarities ranging from highly nonpolar to extremely polar enables the formulation of mixtures with solvating capabilities that may be tailored over a very broad range. An additional feature of organic solvents is their intrinsic ability to control the porous properties of the monoliths.

O~OH 17 In contrast to the process of sonication used originally by Hjert6n's group to disperse hydrophobic monomers in an aqueous buffer [30,31], Palm and Novotny simplified the incorporation of highly hydrophobic ligands into acrylamide-based matrices by using mixtures of aqueous buffer and N-methylformamide to prepare homogeneous polymerization solutions [35]. The overall concentration of the monomers (acrylamide 4, methylene bisacrylamide fi, acrylic acid 17, and C4, C6, or C 12 alkyl acrylate) in solution was kept constant throughout the study at the level of 5%. The composition of the mixed buffer/methylformamide solvent depended on the type of alkyl methacrylate used, and ranged from 50% N-methylformamide for butyl acrylate to 95% for dodecyl acrylate. Columns with high efficiencies were only obtained when the polymerization was performed in the presence of poly(ethylene glycol) (Mw = 10 000) dissolved in the polymerization mixture. Poly(ethylene glycol) is known to induce lateral aggregation of acrylamide chains, thus contributing to the formation of more porous structures [36]. Polymerization was achieved using the usual peroxodisulfate/TEMED initiating system within acryloylsilylated capillaries,

201

Monoliths

affording monoliths possessing an opaque appearance characteristic of macroporous polymers. However, no characterization of the pore structures was performed. Once the polymerization was complete, the poly(ethylene glycol) and other low molecular weight compounds were washed out of the column using electroosmotic flow. This preparation method is remarkably reproducible. All of the monoliths containing the various alkyl acrylates behaved as typical reversed-phase stationary phases, as evidenced by the linear decrease in their retention factors in response to an increasing percentage of organic solvent in the mobile phase. Column efficiencies calculated for on-column detected peaks of phenylketones used as model analytes were in the remarkable range of 300 000-400 000 plates/m. These monolithic columns easily tolerated rather high loading levels without concomitant loss of efficiency, though excessive tailing of the peaks was observed under overload conditions. In contrast to the typical hydrophobic aromatic model compounds often used, Novotny's group extended the range of potential analytes to include sugars, oligopeptides, steroids, and bile acids [35,37,38]. Since sugars are best separated at low pH values, vinylsulfonic acid 13 was incorporated into the monolith rather than acrylic acid in order to provide moieties that would support EOF under these conditions. Figure 6.10 shows a typical separation of a oligosaccharide ladder. Since oligosaccharides do not adsorb light in the UV range, aminobenzamide tags were attached to the analyte molecules prior to the separation, and laser-induced fluorescence was used for their detection. In different run, column efficiencies for glucose, maltose, and maltotriose were all found to be in the range of 190 000-230 000 plates/m. A monolithic CEC column incorporating dodecyl acrylate 18 was also successfully used for the isocratic separation of charged molecules - oligopeptides (di-, tri, penta-, and hexapeptide). The baseline separation was achieved in isocratic mode in less than 5 minutes at 900 V/cm. The elution pattern and the efficiency of the separation were found to strongly depend on both the percentage of acetonitrile and the pH of the mobile phase, suggesting that a gradient elution method would have been even more appropriate. Larger proteins could not be eluted isocratically [35].

O~OC12H25 18

References pp. 238-240

Chapter 6

202

25 Ol~?

~176 l Ol~

GI~

0 (D

O]~lO

0 GI~

.8

I

18

.....

I

18

!

I

1

20 22 24 Time (min)

f-

26

I

28

I

30

Fig. 6.10. Isocratic electrochromatography of an oligosaccharide ladder in a capillary filled with a macroporous polyacrylamide/poly(ethylene glycol) matrix, derivatized with C4 ligands (15%) and containing vinylsulfonic acid (10%) (Reprinted with permission from [35]. Copyright 1997 American Chemical Society). Conditions: capillary length, 50 cm (40 cm effective length) x 100 ~tm i.d.; mobile phase, 0.1% aqueous acetic acid containing 5% (v/v) acetonitrile; field strength, 600 V/cm,; injection, 100 V/cm for 5 s; sample concentration, 30 mg/mL in derivatization solvent and thereafter diluted 1:100 in the mobile phase.

Another successful example is the separation of a series of steroids listed in Fig. 6.11 using a monolithic capillary column prepared by redox initiated polymerization of a solution of acrylamide 4, methylene bisacrylamide fi, vinylsulfonic acid 13, and dodecyl acrylate 18 in N-methylformamide/TRIS-boric acid buffer (pH 8.2) to which poly(ethylene glycol) (MW 10,000) was added (overall composition: 5% T, 60% C, 10% vinylsulfonic acid, 15% lauryl acrylate, 3% poly(ethylene glycol)). The capillary tube was first vinylized and its part beyond the detection window was coated with linear polyacrylamide to avoid band broadening. Since laser induced fluorescence was used to decrease the detection limit of the method to about 100 attomoles for neutral steroids, all of the analytes were first tagged with dansylhydrazine. Fig. 6.12 shows an

203

Monoliths

0

H

0

H

Androsterone

Dehydroandrosterone

5-13-Androstan-17-one

0

Estrone

0

H

11-13-Hydroxyandrosterone o

0

19-Hydroxy-4-androstene-3,17-dione

0

Equiline

Progesterone

Fig. 6.11. Structures of steroids separated using monolithic poly(acrylamide-co-methylene bisacrylamide-co-vinylsulfonic acid) capillary column.

excellent separation of an artificial mixture of eight steroids in less than 30 min using the gradient elution [37]. The potential of the method developed in Novotny's group is demonstrated on the separation of steroids extracted from a real world sample of pregnant human urine (Fig. 6.13). Using retention times, spiking, and mass spectroscopy, a number of peaks could be safely assigned to specific compounds. The same monolithic column was also used for the separation of free and glycine bile acids shown in Fig. 6.14. A very high column efficiency of up to 300,000 plates/m enabled the separation of chenodeoxycholic and deoxycholic acids at a 40 femtomole level [371. In contrast to free and glycine conjugated bile acids, negatively charged taurine conjugates could not be injected electrophoretically and migrated towards cathode in a direction opposite to the electroosmotic flow. Therefore, a new monolithic column was prepared by in situ polymerization of a solution of acrylamide 4, methylene bisacrylamide 5, 3-aminopropane vinyl ether (APVE) 19, [2-(acryloyloxy)-ethyl]trimethyl ammonium methyl sulfate (AETMA) 20, and poly(ethylene glycol) (MW 10,000) in formamide and TRIS-boric acid buffer (5% T, 60% C, 30% AETMA, 30% APVE, 3% poly(ethylene glycol)). It is worth noting that this mixture does not contain any strictly hydrophobic component and the polarity of this polymer resembles that of materials typically used for the separations in normal-phase mode. Indeed, Figure 6.15 documents that the less polar compounds elute prior to more polar analytes containing a larger number of

References pp. 238-240

Chapter 6

204

$ 6 r r r

7b

3

=

2

9

l

5

10

7a

I

! |

ii

15 20 Retention time, min

25

30

Fig. 6.12. Gradient CEC separation of derivatized neutral steroids. (Reprinted with permission [38]. Copyright 2000 Elsevier). Conditions: Column 35 cm (active length 25 cm) x 100 ~tm i.d., mobile phase gradient of acetonitrile-water-240 mmol/L phosphate buffer pH 3 from 35:60:5 to 65:30:50 in 15 min; 600 V/cm; injection 100 V/cm for 10 s. Peaks: labeling reagent 1, progesterone 2, 1l[3-hydroxyandrosterone 3, dehydroisoandrosterone and equiline 4, estrone 5, androsterone 6, 19-hydroxy-4-androsterone-3,17-dione 7, 5-~-androstan- 17-one 8.

O'~O

~N(CH3)3"SO2CH2

19

~,,,,.O~

NH2 20

hydroxyl groups. However, the elution order of cholic acid, taurine conjugate, and glycine conjugate indicate that a mixed mode is most likely operative in this separation [38]. Under the separation conditions using a mobile phase with a pH value of 3, taurine conjugate is a strong anion since its pKa = 1.5 and exhibits high electrophoretic mobility, glycine conjugate with pKa = 4 is a weak anion and its mobility is

205

Monoliths

2.5

2!

0.3

!

|

5

|

10 15 Retention time, min

|

2O

Fig. 6.13. Gradient CEC separation of derivatized urinary neutral steroids extracted from pregnancy urine. (Reprinted with permission from [37]. Copyright 2000 Elsevier). Conditions: Column 35 cm (active length 25 cm) x 100 gm i.d., mobile phase gradient of acetonitrile-water-240 mmol/L phosphate buffer pH 3 from 35:60:5 to 65:30:5 in 15 min; 600 V/cm; injection 100 V/cm for 10 s. Peaks: labeling reagent 1, 11-[3-hydroxyandrosterone 2, dehydroisoandrosterone 3, estrone 4, spiked androsterone 5.

slow, and uncharged cholic acid (pKa = 6) moves with the electroosmotic flow. Peak sharpening, originally observed by Smith for the separations of basic tricyclic antidepressants using capillary columns packed with strong acid cation exchangers [39], leads to apparent column efficiencies in the range of 600,000 plates/m. Although both reproducible preparation and operation of CEC columns are extremely important issues that will further stimulate the development and the acceptance of this technique, only a few groups have reported data on column-to-column, run-to-run, and day-to-day reproducibility of monolithic capillary columns. Palm and Novotny showed reproducibility data for migration times tr, efficiencies, and retention factors k' for a number of analytes on acrylamide-based monoliths [35]. The relative standard deviations (RSD) were smaller for run-to-run compared to dayto-day measurements. For example, the average run-to-run RSD for 6 analytes was

References pp. 238-240

Chapter 6

206

O

OH' NH OH' NH O

"NH

i

O

.

i

/

S~~M :,,,,~.,../i..,./"

HO,,F/~x./i...... / S ~ O 3 H / ~ o... H S~O3H H H H Taurolihtocholicacid(TLCA) Taurodeoxycholicacid(TDCA) Taurocholicacid(TCA)

O ~~~'~NH

HOf'''1 ''''I" H

Glycolithocholicacid(GLCA)

O

O

_OH" Y ~ N H

HOf~J'~"~ij H

OH~ ~ " ~ N H

HOf~

Glycodeoxycholicacid(GDCA)

H9

H

....OH

Glycocholicacid(GCA)

OOH

HOf~'~OH H Cholicacid(CA)

Fig. 6.14. Structures of bile acids separated using monolithic poly(acrylamide-co-methylene bisacrylamide-co-vinylsulfonic acid) capillary column.

0.8% for tr, 2.6% for k', and 4.3% for the efficiency, while the average day-to-day RSDs for the same variables were 2.1%, 6.1%, and 4.6%, respectively. 6.5 IMPRINTED ENANTIOSELECTIVE MONOLITHS

Molecular imprinting has recently attracted considerable attention as an approach to the preparation of polymers containing recognition sites with predetermined selectivity. The history and specifics of the imprinting technique pioneered by Wulff in the 1970s have been detailed in brilliant review article [40]. These materials, if successfully prepared, are expected to find applications in numerous areas such as the resolution of racemates, chromatography, substrate selective catalysis, and the production of "artificial antibodies". Imprinted monoliths have also recently received

207

Monoliths

100%

< O O

< r.a I-. O

< r I-

l

< O _l

O

< O

I--

0

3.42

<

0

<

I

7.19 Time, min

i

I

10.56

i

14.33

Fig. 6.15. Normal-phase CEC negative ion ESI-MS separation of a mixture of free bile acids, glycine, and taurine conjugates using monolithic poly(acrylamide-co-methylene bisacrylamide-co-3-aminopropane vinyl ether-co- [2-(acryloyloxy)-ethyl]-trimethyl ammonium methyl sulfate) capillary column. (Reprinted with permission from [38]. Copyright 2000 Elsevier). Conditions: Column 30 cm x 100 pm i.d., mobile phase 35:60:5 acetonitrile-water-240 mmol/L phosphate buffer pH 3; 400 V/cm; injection 6 kV/cm for 5 s. For peaks assignment see Fig. 6.14.

attention as stationary phases for capillary electrochromatography [41-48]. Due to their specifics, these monolithic materials are detailed in a specific chapter of this book. 6.6 POLYSTYRENE-BASED M O N O L I T H S Horv/tth's group has recently reported the preparation of porous rigid monolithic capillary columns for CEC by polymerizing mixtures of chloromethylstyrene 21, divinylbenzene 22 and azobisisobutyronitrile in the presence of various porogenic solvents such as methanol, ethanol, propanol, toluene, and formamide [49]. The capillary

wall

was

silanized

using

a

50%

dimethylformamide

3-(trimethoxysilyl)propyl methacrylate 8 at a temperature of 120~

solution

of

for 6 hours. In

order to avoid the spontaneous polymerization of the functional methacrylate, a stable free radical (DPPH) was added to the solution. The SEM micrographs of Fig. 6.16

References pp. 238-240

Chapter 6

208

Fig. 6.16. SEM micrographs of poly(styrene-divinylbenzene) based monolithic capillary. (Reprinted with permission from [49]. Copyright 1999 Elsevier).

I

21

22

209

Monoliths

m

E

E

=,,=

0

3 4

o

0

.......

0

|

5

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

|,, ,

10

minutes

A

15

Fig. 6.17. Electrochromatogram of acidic and basic peptides (Reprinted with permission from [49]. Copyright 1999 Elsevier). Column 31 cm (21 active length)x 75 gm, porous styrenic monolith with dimethyloctylammonium functionalities; mobile phase 25% acetonitrile in 5 mmol/1 phosphate buffer pH 3.0 containing 50 mmol/l sodium chloride; reversed polarity, electrokinetic injection for 2 s, 5 kV. Peaks: angiotensin II (1), angiotensin I (2), [Sar 1, AlaS]-angiotensin II (3), insulin (4).

show that the resulting monolith exhibits a "classical" macroporous structure "wrapped" in a thin outer layer of apparently nonporous polymer. The reactive chloromethyl moieties incorporated into the monolith served as sites for the introduction of quaternary ammonium functionalities. The pores of the monolith were filled with N,N-dimethyloctylamine, and after a suitable reaction period, the column was washed with methanol and equilibrated with the mobile phase. Unfortunately, only very limited information concerning the nature and extent of modification was presented. These capillary columns possessing positively charged surface functionalities were used for the reversed-phase separations of basic and acidic peptides. Figure 6.17 shows the separation of three angiotensins and insulin with plate numbers as high as References pp. 238-240

210

Chapter 6

200,000 pl/m using a mobile phase consisting of acetonitrile and phosphate buffer (pH = 3). Surprisingly, the retentions of both angiotensins I and II increased as the percentage of acetonitrile in the mobile phase was increased from 25 to 45%, and no elution was observed at higher percentages of organic solvent. Good separation of chemically similar tripeptides (Gly-Gly-Phe and Phe-Gly-Gly) was also observed in a pH 7 buffer using unfunctionalized poly(styrene-co-divinylbenzene) monoliths devoid of charged functionalities. In this case, the driving force for movement of the analytes through the column is their electrophoretic migration, while separation results from their interactions with the stationary phase [49]. However, the addition of acetonitrile to the mobile phase significantly decreases the analytes mobility, making this approach less attractive. Huang et al. also produced macroporous polymer layer open tubular capillary columns (PLOT) for CEC similar to those employed in gas chromatography [50]. Narrow bore capillaries (20 ~tm i.d.) are required in order to achieve high efficiency. The preparation process is closely related to that described previously. A vinylized capillary was filled with a polymerization solution of chloromethylstyrene 21, divinylbenzene 22, and 2-octanol as a porogen, and an in situ polymerization was performed. The resulting porous polymer layer was functionalized by reaction with N,N-dimethyloctadecylamine and any residual chloromethyl groups quenched by reaction with a solution of sodium hydroxide. Although no experimental data concerning the extent of functionalizations or process was published, the surface coverage of positively charged functionalities was sufficiently large to afford EOF velocities of 2.1-2.5 mm/s that did not change significantly over the broad range of 0-60% acetonitrile in the mobile phase. Horv~th's group also demonstrated the complex nature of CEC separations that often involve the interplay of EOF, electrophoretic migration, and chromatographic retention through several examples of peptide and protein separations using the PLOT capillary columns. For example, the retention of proteins was seen to increase with increasing percentage of organic solvent in the mobile phase. Even more intriguing is the demonstrated isocratic separation of proteins shown in Fig. 6.18. Although the window of mobile phase compositions within which this separation could be achieved was narrow, this approach deserves further study since it may eliminate the need for the rather complex gradient elution instrumentation. Gusev et al. monitored the conductivity of his modified monolithic polystyrenebased columns for over 3 months and observed no changes [49]. Similarly, the electroosmotic mobility was measured over a number of days and again almost no changes were found [50]. This demonstrates excellent stability of the polystyrenebased monolithic column.

Monoliths

211

r~

I. I.

! |

I

!

6

8

10

lime [mini

Fig. 6.18. Electrochromatogram of four basic proteins obtained by isocratic separation using a modified polychloromethylstyrene-based PLOT column (Reprinted with permission from [50]. Copyright 1999 Elsevier). Column 47 cm (active length 40 cm) x 20 gm, inner polymer layer 2 ~tm; mobile phase 20% acetonitrile in 20 mmol/1 phosphate buffer pH 2.5; voltage -30 kV; EOF velocity measured with dimethylsulfoxide (DMSO) -3.46 x 10-8 m2V-ls-1, migration time for DMSO 3.10 min. Peaks: a-chymotrypsinogen (1), ribonuclease (2), lysozyme (3), cytochrome C (4).

Zhang's group in China developed monolithic poly(styrene-co-divinylbenzene) CEC column in which EOF is supported by carboxyl groups of polymerized methacrylic acid units (Xiong et al. [51 ]). In a typical procedure, vinylized 75 mm i.d. capillaries were filled with a mixture of 5% styrene 23, 10% divinylbenzene 22, 5% methacrylic acid 1, and 80% toluene containing 1% azobisisobutyronitrile (in respect to monomers) and polymerized at 70~ for 24 h. The pore volume of 0.098 mL/g and mean pore size of 40 nm determined for this monolith appear to be rather small and do not correspond with the published SEM pictures that reveal existence of large pores, and the chromatographic performance of the columns in CEC mode.

23 References pp. 238-240

Chapter 6

212

1

0

1

2

3

3

4 min

Fig. 6.19. CEC separation of phenylenediamines using monolithic poly(styreneco-divinylbenzene-co-methacrylic acid) capillary column. (Reprinted with permission from [50]. Copyright 1999 Wiley-VCH). Conditions: Column 23 cm (active length 17 cm) • 75 ~tm i.d., mobile phase 70:30 acetonitrile-4 mmol/L TRIS buffer solution pH 8.8; injection 1 kV for 1 s; UV detection at 200 nm.

Using benzene as a probe, column efficiencies of 90,000-150,000 were observed within a flow velocity range of 1-10 cm/min (0.2-1.7 mm/s). Using mobile phases with different percentages of acetonitrile in aqueous buffer solution (pH 8.82), linear dependency of retention factors for a series of alkylbenzenes on the volume fraction of organic modifier was found. This indicates again that the reversed phase separation mechanism is operative for this type of monoliths. Different families of compounds such as phenols, chlorobenzenes, anilines, phenylendiamines, and alkylbenzenes were well separated typically in less than 5 min using 20 cm long columns and a field strength of 20-30 kV. Fig. 6.19 shows the separation of phenylenediamines as an example of the column performance [51 ]. 6.7 METHACRYLATE ESTER-BASED MONOLITHIC COLUMNS In contrast to the acrylamide and styrene-based monoliths that have largely been characterized by their chromatographic performances, extensive materials development and optimization have been performed for monolithic CEC capillaries prepared

Monoliths

213

from methacrylate ester monomers. These investigations made use of the concepts developed from our original work with the macroporous discs and molded rigid monolithic HPLC columns that we introduced in the late 1980s and early 1990s [7,52,53]. The experience acquired earlier with these materials proved helpful in investigating the interrelated effects of morphology and composition on the overall CEC process.

O

o 24

Production of these monolithic capillary columns is simple (Fig. 6.20) [54]. Either a bare or a surface treated capillary is filled with a homogeneous polymerization mixture consisting of porogenic solvents, butyl methacrylate 14, ethylene dimethacrylate

24,

and

2-acrylamido-2-methyl-l-propanesulfonic

acid

6,

and

radical

polymerization is initiated only when desired using either a thermostated bath [54] or UV irradiation [55] to afford a rigid monolithic porous polymer. Once the polymerization is complete, unreacted components such as the porogenic solvents are removed from the monolith using a syringe pump or electroosmotic flow. This simple method for preparing monolithic capillary columns has numerous advantages. For example, the fused silica tubing may be used either directly as supplied without first performing any chemical modification of its internal surface or after its functionalization using a suitable vinyl containing moiety. All of the chemicals may be used as supplied, although careful purification contributes to better batch-to-batch reproducibility (vide infra). Additionally, the final polymerization mixture contains free radical initiators such as benzoyl peroxide or azobisisobutyronitrile, ensuring its stability and easy handling for several hours at room temperature or for days in the refrigerator without risking the onset of polymerization. In optimizing the process, specific attention was paid to the design of the porogenic mixtures. Ideally, this system had to enable (i) the preparation of a homogeneous, single phase polymerization mixture from the charged, water soluble monomer that supports the EOF, and the hydrophobic monomers that affect the separation without using additional compatibilizing agents; (ii) the direct uniform incorporation of these monomers with widely differing polarities into a macroporous polymer monolith; (iii) the fine control of the porous properties of the resulting monolith over a broad range; and finally, (vi) the facile initial washing and equilibra-

References pp. 238-240

214

Chapter 6

BARE C A P I L L A R Y .....]

A) Filling POLYM. MIXTURE FILLED CAPILLARY

B) Polymerization 1 THERMAL INITIAT I ON

I

_

~C)

UV INITIATI ON ]

Washing

l

Mechanical pumping

EOF

I

I

MONOLITHIC COLUMN

Fig. 6.20. Schematics for the preparation of monolithic capillary columns. First, the bare capillary is filled with the polymerization mixture (step a) that contains functional monomer, crosslinking monomer, initiator, and porogenic solvent. Polymerization (step b) is then initiated thermally or by UV irradiation to afford a rigid monolithic porous polymer. The resulting monolith within the capillary is washed (step c) with the mobile phase using a pump or electroosmotic flow and used as for the CEC separations.

Monoliths

215

1 2

9

~...~ '

0

I

2

'

.....~ ~..../L.........I

4

'

I

6

8 min

Fig. 6.21. Electrochromatographic separation of benzene derivatives on monolithic capillary column prepared by UV initiated polymerization. Conditions: capillary column, 100 lam i.d. x 25 cm active length; stationary phase poly(butyl methacrylate-co-ethylene dimethacrylate) with 0.3 wt. % 2-acrylamido-2-methyl-l-propanesulfonic acid; pore size, 296 nm; mobile phase, 75:25 vol./vol mixture of acetonitrile and 5 mmol/L phosphate buffer pH 7; UV detection at 215 nm; 25 kV; pressure in vials, 0.2 MPa; injection, 5 kV for 3 s. Peaks: thiourea (1), benzyl alcohol (2), benzaldehyde (3), benzene (4), toluene (5), ethylbenzene (6), propylbenzene (7), butylbenzene (8), and amylbenzene (9).

tion of the capillary column by being miscible with the mobile phase used for electrochromatography. An extensive study led to the development of a ternary porogen system consisting of water, 1-propanol, and 1,4-butanediol in various proportions [54]. Monolithic capillary columns prepared using this porogen system and photochemical initiation possessed efficiencies of over 210 000 pl/m for the separation of a model mixture of aromatic compounds shown in Fig. 6.21 [55]. Similarly, peptides were separated on this capillary column using a mobile phase containing 1-octanesulfonic acid (Fig. 6.22). This ion-pairing alkylsulfonic acid additive likely affects the separation of the peptides both by associating with the terminal amine groups of the peptides and preventing them from interacting with the negatively charged surface functionalities of the monolith as well as by increasing the hydrophobicity of the analytes [55].

References pp. 238-240

216

Chapter 6

~D r~

r~

2

i '

I

I

'

I

2 4 6 Retention time, min

'

l

8

Fig. 6.22. Electrochromatographic separation of Gly-Tyr (1), Val-Tyr-Val (2), methionine enkephalin (3), and leucine enkephalin (4) on monolithic methacrylate capillary column with a pore size of 492 nm. (Reprinted with permission from [55]. Copyright 1999 Wiley-VCH). Conditions: Mobile phase 10% of aqueous 10 mmol/L sodium 1-octanesulfonate and 90% of a 2:8 mixture of 5 mmol/L phosphate buffer pH=7.0 and acetonitrile. UV detection at 215 nm. Total sample concentration 1 mg/mL.

In contrast, recently we prepared enantioselective monolithic columns consisting of ethylene dimethacrylate 24, O-[2-methacryloyl oxy)ethylcarbamoyl]- 10,11-dihydroquinidine 25, and glycidyl methacrylate 26 or 2-hydroxyethyl methacrylate 8 [56]. Since the monomer mixture does not involve any highly hydrophilic component such as 2-acrylamido-2-methyl-l-propanesulfonic acid, we could use the traditional mixture of cyclohexanol and dodecanol as the porogen that proved in the past to be a very powerful allowing fine control of porous properties of monolithic HPLC columns [57,58]. Changing proportions of these solvents in the polymerization mixture, it is possible to adjust the pore size of the monolithic materials in a very broad range. Typically the nature of the comonomer as well as the initiation method affects the porous structure. In contrast to monoliths prepared with 26, a much higher percentage of dodecanol in the dodecanol/cyclohexanol mixture is required to obtain polymerized 2-hydroxyethyl methacrylate units containing monoliths with sufficiently large pores. Fig. 6.23 shows the effect of dodecanol content in the porogenic solvent mixture on

Monoliths

E

217 2000 -

(a)

3000

c

2000

E

.$ "o

1000

o I:1..

0

1000

-,'=

0

20

40

60

0

,,

20

,

40

I

60

Dodecanol (wt %) Fig. 6.23. Effect of thermal (a) and UV initiation (b), type of comonomer, and percentage of 1-dodecanol in the polymerization mixture on the mode pore diameter of quinidine-functionalized chiral monoliths. (Reprinted with permission from [56]. Copyright 2000 American Chemical Society). Reaction conditions: polymerization mixture, chiral monomer 25 8 wt%, glycidyl methacrylate (D) or 2-hydroxyethyl methacrylate (I) 16 wt%, ethylene dimethacrylate 16 wt%, porogenic solvent 60 wt% (consisting of 1-dodecanol and cyclohexanol), polymerization time 20 h at 60~ (a) and 16 h at room temperature (b).

ocH3

o

mode pore size (pore diameter at the maximum of the distribution curve) for monoliths prepared from mixtures containing 26 or 9 using either thermal or UV initiations. For example, use of 20% dodecanol and 40% cyclohexanol for the preparation of glycidyl methacrylate monoliths by thermal polymerization at 60~

affords material

with a mode pore size of 1,000 nm. In contrast, a mixture containing 50% cyclohexanol and 50% of the less polar dodecanol is required for the preparation of 2-hydroxyethyl methacrylate-containing monoliths with a similar pore size.

References pp. 238-240

218

Chapter 6

Photoinitiated polymerization of the same mixtures at 20~ generally yields monoliths with larger pores compared to those initiated thermally. Thus, reduced contents of dodecanol in the polymerization mixture has to be used for UV initiated polymerizations in order to obtain pore sizes comparable to those of their thermally polymerized analogs. For example, a polymerization mixture containing only 30% dodecanol can be used to produce a 2-hydroxyethyl methacrylate monolith with 1,000 nm pores by UV polymerization at 20~

These shifts can readily be explained by the

effect of the polymerization temperature, since the creation of larger pores is favored at lower temperatures [59]. Similarly, the porous properties of the monoliths also depend on the percentage of chiral monomer 25 in the polymerization mixture. An increase in the percentage of 25 in the polymerization mixture at a fixed composition of porogen leads to a significant decrease in the pore size for both photoinitiated and thermally initiated polymerizations. For example, a photopolymerized monolith with a mode pore diameter of 1,600 nm is obtained using a mixture consisting of 4% 25 and 20% 2-hydroxyethyl methacrylate, in addition to 16% ethylene dimethacrylate, 35% 1-dodecanol, and 25% cyclohexanol. Monoliths with smaller pore diameters of 1,400 and 600 nm are obtained if the percentage of 25 is increased to 8% and 12% respectively, with a concomitant decrease in percentage of 2-hydroxyethyl methacrylate [60]. The methacrylate-based polymers are stable even under extreme pH conditions such as pH 2 or 12. Fig. 6.24 shows the CEC separations of aromatic acids and anilines at these pH values [14]. The sulfonic acid functionalities of the monolithic polymer remain dissociated over the entire pH range creating a flow velocity sufficient to achieve the separations in a short period of time. In contrast to the stationary phase, the analytes are uncharged, yielding symmetrical peaks. Needless to say that typical silica-based packings may not tolerate such extreme pH conditions. Tests of the reproducibility of retention times, retention factors, separation selectivities, and column efficiencies for our methacrylate monolithic capillary columns are summarized in Table 6.2. This table shows averaged data obtained for 9 different analytes injected 14 times repeatedly every other day over a period of 6 days, as well as for 7 different capillary columns prepared from the same polymerization mixture. As expected, both injection-to-injection and day-to-day reproducibilities measured for the same column are very good. Slightly larger RSD values were observed for column-to-column reproducibility. While the selectivity effectively did not change, larger differences were found for the efficiencies of the columns.

Monoliths

219

pH=12

1

~D r~

m.

1

5

4 e~ cD

,

0

5

10

0

[

4

'

8

R e t e n t i o n time, min Fig. 6.24. Electrochromatographic separation of aromatic acids (a) and anilines (b) on monolithic capillary columns. (Reprinted with permission from [14]. Copyright 2000 Elsevier). Conditions: monolithic poly(butyl methacrylate-co-ethylene dimethacrylate) stationary phase with 0.3 wt. % 2-acrylamido-2-methyl-l-propanesulfonic acid; pore size, 750 nm; UV detection at 215 nm; voltage, 25 kV, pressure in vials, 0.2 MPa; injection, 5 kV for 3 s. (a) capillary column, 100 ~tm i.d. • 30 cm (25 cm active length); mobile phase, 60:40 vol./vol mixture of acetonitrile and 5 mmol/L phosphate buffer pH 2.4. Peaks: 3,5-dihydroxybenzoic acid (1), 4-hydroxybenzoic acid (2), benzoic acid (3), 2-toluic acid (4), 4-chlorobenzoic acid (5), 4-bromobenzoic acid (6), 4-iodobenzoic acid (7). (b) capillary column, 100 ~tm i.d. x 28 cm (25 cm active length); mobile phase, 80:20 vol./vol mixture of acetonitrile and 10 mmol/L NaOH pH 12. Peaks: 2-aminopyridine (1), 1,3,5-collidine (2), aniline (3), N-ethylaniline (4), N-butylaniline (5).

References pp. 238-240

Chapter 6

220 TABLE 6.2

REPRODUCIBILITY OF THE ELECTROCHROMATOGRAPHIC PROPERTIES OF METHACRYLATE-BASED MONOLITHIC CAPILLARIES

Variable

RSD % Run-to-run n=14

Day-to-day n=3

Column-to-column n=7

Retention time

0.18

1.19

3.50

Retention factor

0.21

0.30

1.43

Selectivity

0.05

0.10

0.11

Efficiency

1.50

4.30

7.80

Conditions: capillary columns, 100 pm i.d. x 30 cm active length; stationary phase poly(butyl methacrylate-co-ethylene dimethacrylate) with 0.3 wt.% 2-acrylamido-2-methyl-l-propanesulfonic acid; mobile phase, 80:20 vol./vol, mixture of acetonitrile and 5 mmol/L phosphate buffer pH 7; UV detection at 215 nm; voltage 25 kV; pressure in vials 0.2 MPa; sample concentration 2 mg/mL of each compound; injection 5 kV for 3 s. Data shown are average RSD values obtained for thiourea, benzyl alcohol, benzaldehyde, benzene, toluene, ethylbenzene, propylbenzene, butylbenzene, and amylbenzene.

6.8 A S S E S S M E N T OF THE P O R O U S S T R U C T U R E

The ability of a liquid to flow through the network of channel-like pores that traverse the length of these monolithic materials is essential to all of their applications. In addition to providing permeability, the porous structure also accelerates the rate of mass transfer within the separation medium as a result of convection [61 ], since all of the mobile phase flows through the pores [62]. Despite this important fact, only a limited number of studies directly assessing the effect of pore size of the monolithic CEC media have been published [54]. The absence of data for other monolithic systems is probably due both to the limited means available to control their porous structures during preparation as well as to difficulties in determining their actual pore structure in the swollen state. It should be emphasized that the standard methods typically used for the visualization and measurement of porous properties such as scanning electron microscopy, mercury intrusion porosimetry and nitrogen absorption/desorption, are performed on materials in the dry state, while the columns actually operate in the presence of a solvent. As a result, the data measured in the dry

Monoliths

221

state may not completely reflect the operational pore size of the capillaries during the chromatographic process. A number of authors demonstrated the porous structures of their monoliths using scanning electron micrographs (for example [14,32,33,49,63]. Some of these pictures are also shown throughout this chapter. Although these micrographs are impressive and visualize the macroporous structure, they do not enable a quantitative determination of parameters of the porous structure. In a recent study, Gusev et al. used three methods to determine the porosity of monolithic capillary columns in the "solvated" state [49]. First, the elution time of a low molecular non-retained tracer in ~HPLC was used to calculate the total porosity. The second method afforded an estimation of the porosity from the conductivity ratio. Monolithic and empty capillaries were filled with an electrolyte and their conductivities were measured. Although several equations relating conductivity ratio to total column porosity have been derived, Archie's equation appeared to provide the best fit of the experimental data. The last method was gravimetric, using the weight difference between a dry and acetone filled monolithic column. Since none of these three methods affords information about pore size distribution, liquid extrusion porosimetry with hexadecane was used to determine the integral pore volume distribution. However, since this technique requires samples larger than those available from a capillary column, it was performed using a material prepared via a larger scale bulk polymerization [49]. Similarly, we polymerized the same mixture used for the preparation of capillary columns in glass vials and used the product for mercury intrusion porosimetry. Since we found that a strong correlation exists between the "dry" porous properties of the monoliths and their chromatographic performance, even dry porosity measurements may be used to tailor column performance. 6.8.1 Control of porous properties

The novel ternary porogenic system that we have developed enables precise control of porous properties over a broad range [64]. For example, the percentage of 1-propanol in the porogenic solvent exerts an enormous effect on the pore diameter at the maximum of the distribution curve (mode pore diameter) as documented in Fig. 6.25 for the UV initiated polymerization system. Based on these results, monoliths of any pore size within the broad range of 250-1300 nm can easily be produced by simply changing the ratio of propanol to butanediol in the porogenic mixture. It should be noted that the window of weight percentage of 1-propanol that brackets this wide range of pore sizes is sufficiently large to obtain polymers of any mode pore diameter with an accuracy of 25 nm with respect to the targeted value. Despite the fact that these monoliths are prepared from a polymerization mixture containing monoReferences pp. 238-240

Chapter 6

222

1500IM

t~ lO00e~

0

s0o"-II

58

59

60 61 l-Propanol, %

62

Fig. 6.25. Effect of the percentage of 1-propanol in the porogenic mixture on the porous properties of monolithic polymers (Reprinted with permission from [64]. Copyright 1998 American Chemical Society). Reaction conditions: polymerization mixture: ethylene dimethacrylate 16.00 wt.%, butyl methacrylate 23.88 wt.%, 2-acrylamido-2-methyl-1propanesulfonic acid 0.12 wt.%, ternary porogen solvent 60.00 wt.% (consisting of 10 wt.% water and 90 wt.% of mixtures of 1-propanol and 1,4-butanediol), azobisisobutyronitrile 1 wt.% (with respect tomonomers), polymerization time 20 h at 60~

mers of very different polarities, all of the mercury porosimetry profiles seen in Fig. 6.26 exhibit distribution curves similar to those found for polymers prepared from mixtures of fully miscible monomers [58], as well as the CEC systems described initially [64]. Such precise control of porous properties is expected to be very useful in the design of specialized CEC columns for separation in modes other than reversed-phase. For example, size exclusion chromatography (SEC) is an isocratic separation method that relies on differences in the hydrodynamic volumes of the analytes. Because all solute-stationary phase interactions must be avoided in SEC, solvents such as pure tetrahydrofuran are often used as the mobile phase for the analysis of synthetic polymers, since they dissolve a wide range of structures and minimize interactions with the chromatographic medium. Despite the reported use of entirely non-aqueous eluents in both electrophoresis and CEC [65], no appreciable flow through the methacrylate-based monoliths was observed using pure tetrahydrofuran as the mobile phase. However, a mixture of 2% water and tetrahydrofuran was found to substan-

223

Monoliths

12

,

i

i

,

,,,,,

4

3 2

i

61

w

10

100 1 000 Pore diameter, nm

10 000

Fig. 6.26. Differential pore size distribution profiles of porous polymeric monolithic capillary columns with mode pore diameters of 255 (curve 1), 465 (2), 690 (3), and 1000 nm (4) (Reprinted with permission from [64]. Copyright 1997 American Chemical Society).

tially accelerate the flow velocity, while still capable of dissolving polystyrene standards with molecular weights as high as 980 000 [66]. Fig. 6.27 shows the SEC separation of polystyrenes in the CEC mode using a methacrylate-based monolithic capillary column. The rel. mol. mass of the peaks was assigned by injections of the individual standards. The elution order of the polystyrene standards and toluene confirms that size exclusion is the prevailing separation mechanism. Although the porous properties of the monolithic column used for this experiment were not optimized for SEC separations, these results demonstrate that CEC is not limited to the reversedphase mode of chromatography. Extensive studies of SEC separations of polystyrenes in the CEC mode using packed capillary columns and dimethylformamide as the solvent has recently been published by a Dutch group [67-69]. 6.9 EFFECTS OF THE POROUS P O L Y M E R P R O P E R T I E S ON THE

SEPARATION The ability to achieve precise and independent control over both the porous properties as well as the level of charged moieties of the rigid monolithic stationary phases opened new avenues for studies focusing on the effects these properties exert on the References pp. 238-240

Chapter 6

224

80-

2 o',,

40

-

4 1

r~

1

I

0

25 Retention

50 time, min

Fig. 6.27. Electrochromatographic size-exclusion chromatography of polystyrene standards (Reprinted with permission from [64]. Copyright 1998 American Chemical Society). Conditions: monolithic poly(butyl methacrylate-co-ethylene dimethacrylate-co-2acrylamido-2-methyl-l-propanesulfonic acid) capillary column, 100 ktm i.d. x 30 cm active length; stationary phase, 59.7 wt.%; pore size 750 nm; mobile phase, tetrahydrofuran containing 2 vol.% of water; UV detection at 215 nm; voltage, 25 kV; pressure in vials, 0.2 MPa; sample concentration, 2 mg/mL of each compound; injection, 5 kV for 3 s. Peaks: polystyrene standards with a molecular mass of 980,000 (1), 34,500 (2), 7,000 (3), and toluene (4).

chromatographic process. Both of these variables were found to be extremely important in controlling the flow velocity and efficiency of the monolithic capillary CEC columns. In addition to these materials properties, CEC separations are also affected by the conditions under which they are performed, including the applied voltage, and both the pH and elution strength of the mobile phase.

Monoliths

225

6.9.1 Surface chemistry Electroosmotic flow velocity is directly proportional to the zeta potential z that, in turn, is directly related to the surface charge. In contrast to silica-based CEC media, the ability to easily control the level of charged functionalities that support the electroosmotic flow is a major advantage of the polymeric monolithic capillaries. This variable can easily be adjusted by changing the percentage of charged monomer in the polymerization mixture. For example, linear increases in migration velocity paralleling increases in AMPS monomer content were observed by Fujimoto in both 6% crosslinked polyacrylamide gels [15] as well as 9.7% crosslinked N-isopropylacrylamide 7 polymers [27]. Similarly, increasing the content of sulfonic acid groups within the methacrylate ester monoliths significantly increased the flow velocity, thus reducing the overall analysis time. Similar chromatographic performances were maintained in these higher flow capillaries by making concomitant changes in the composition of the porogenic mixture in order to keep the pore sizes of the monoliths effectively constant. In contrast, the monomer 25 in addition to its role of the chiral selector it also provides the driving force for electroosmotic flow. Therefore, an increase in its loading should result in an increase in the electroosmotic flow velocity. Surprisingly, this effect was not very dramatic. Only a minor increase in flow velocity from 0.97 to 1.12 mm/s is observed for monoliths as the loading with the chargeable monomer 25 tripled [56]. Monoliths containing two significantly different percentages of dimethyldiallylammonium chloride 15 were recently prepared in order to control the EOF component of the overall migration rate of proteins [31 ]. These charged moieties were incorporated into the monolith during a later stage of the preparation process. This process appeared to be well suited to achieve monolith with properties required for the desired separations (vide supra). The majority of CEC separations reported to date have been performed in the reversed-phase mode. Under these conditions, the hydrophobicity of the stationary phase determines the selectivity of the separation, and retention can easily be controlled by adjusting either the composition of the mobile phase or the hydrophobicity of the surface, with the first option being easier to implement. However, in contrast to the rich variety of solvents available for use in HPLC, acetonitrile-based solvent systems are employed in most CEC applications due to their high dielectric constant and low viscosity [30,35,51,64,70]. Little has been done to date to tailor the surface chemistry of the stationary phase in CEC. Novotny prepared a series of monolithic columns incorporating a variety of monomers that differ in the length of their pendant alkyl chains [35]. Unfortunately, the performance of these monoliths containing butyl, hexyl, and dodecyl methacrylate

References pp. 238-240

Chapter 6

226

moieties has not been demonstrated in comparative separations, preventing an assessment of the effect of such changes in surface chemistry for a homologous series of monoliths. The effect of surface polarity is even more important in separations where two or more simultaneous interactions must occur in order to achieve the desired selectivity. This is particularly true in chiral separations. Since aqueous buffer systems are almost universally used as CEC mobile phases, enantioseparations are often run under reversed-phase conditions as opposed to the normal-phase mode typically used in chiral HPLC. Therefore, non-specific hydrophobic interactions would be highly detrimental to the discrimination process that involves subtle differences between the enantiomers. The importance of tailoring surface chemistry was first demonstrated by three different monolithic capillary columns that were prepared by directly incorporating of the chiral monomer 2-hydroxyethyl methacrylate (N-L-valine-3,5-dimethylanilide) carbamate 27 [71]. These columns were tested for the enantioseparation of a model

.J~ o

NH o

racemic compound, N-(dinitrobenzoyl)leucine diallylamide. Fig. 6.28 compares the chiral separations achieved using the various columns. Although the column containing butyl methacrylate 14 as a hydrophobic comonomer did resolve the racemic analyte, the peaks were very broad and tailed severely. The efficiency of this system was poor, with plate counts of only 600 and 160 plates/m obtained for the respective enantiomers (Fig. 6.28a). However, when pure acetonitrile was used as the mobile phase, a narrowing of both peaks and a concomitant increase in column efficiency to 2,500 and 540 plates/m were observed. These improvements indicate that the originally observed tailing probably resulted from non-specific hydrophobic interactions between the chiral analyte and the relatively hydrophobic surface of the separation medium. Therefore, new capillary columns were prepared substituting the more hydrophilic glycidyl methacrylate 26 for the highly hydrophobic butyl methacrylate. Although this capillary column exhibited a surprisingly high efficiency of 210,000 plates/m for the unretained peak of thiourea under reversed-phase conditions, the incorporation of the more hydrophilic monomer resulted in a substantial decrease in

227

Monoliths

(b)

(a)

(c)

< E e~

<

0

I

I

5

10

I

I

15 0 5 10 0 Retention time, min

I

I

3

6

Fig. 6.28. Effect of the hydrophilicity of chiral monolithic columns on the electrochromatographic separation ofN-(3,5-dinitrobenzoyl)leucine diallylamide enantiomers (Reprinted with permission from [13]. Copyright 2000 Wiley-VCH). Conditions: monolithic column, 100 ~tm i.d. x 30 cm active length; mobile phase, 80:20 vol./vol, mixture of acetonitrile and 5 mmol/L phosphate buffer pH 7; UV detection at 215 nm; voltage, 25 kV; pressure in vials, 0.2 MPa; injection, 5 kV for 3 s. Stationary phase with butyl methacrylate (a), glycidyl methacrylate (b), and hydrolyzed glycidyl methacrylate (c).

hydrophobic selectivity (OtCH2 =1.08). However, this change in surface polarity resulted in a significantly improved chiral separation (Fig. 6.28b). The peaks for the enantiomers were sharper, and the column efficiencies calculated for this separation increased to 8,100 and 1,900 plates/m. Instead of defining and optimizing new conditions for the direct incorporation of an even more hydrophilic monomer into the monolith, the epoxide rings of the monolith described above that contains glycidyl methacrylate and chiral units were hydrolyzed using dilute aqueous sulfuric acid to afford very hydrophilic diol functionalities 28. This hydrolytic reaction was easily performed in situ within the pores of the monolithic capillary column. After hydrolysis, the diol-functionalized hydrophilic capillary was unable to effect any separation of alkylbenzenes in the reversed-phase mode. However, this monolithic column afforded a significantly improved separation of the enantiomers (Fig. 6.28c). The peaks in this separation were narrow and well resolved (Rs = 2.0). Column efficiencies (61,000 and 49,500 plates/m) were rather high, and even peak tailing was greatly reduced, suggesting that few undesirable References pp. 238-240

228

Chapter 6

OH

OH 28

interactions remained. Unfortunately, this substantial increase in column efficiency was accompanied by a decrease in selectivity [71 ]. In the following study, we prepared monoliths from the quinidine-functionalized chiral monomer 25 and glycidyl methacrylate 26 or 2-hydroxyethyl methacrylate 1t [60]. Figure 6.29 shows the effect of comonomer polarity on the CEC separation of DNB-leucine enantiomers. The polarity of the surface again clearly affects both the enantioselectivity and the efficiency of a monolithic capillary column. The simple replacement of 26 with the more polar It increases the column efficiency of these capillary columns by a factor of 8. Simultaneously, the enantioselectivity for DNBLeu rapidly increases from a selectivity factor et = 1.62 to 3.36. This effect was attributed to the significant reduction of non-specific interactions, as well as the absence of lateral epoxypropyl functionalities with uncontrolled stereochemistry at the central carbon atom. Although the epoxide groups could be again hydrolyzed in situ to obtain more polar diol functionalities shown above, this would represent an additional reaction step that complicates the preparation procedure, and does not address the detrimental effect of the uncontrolled chirality. These examples clearly demonstrate the benefits of the facile tuning of surface chemistry afforded by the monolithic media. The wealth of commercially available monomers possessing a wide variety of functionalities, together with the extreme simplicity of the preparation of the monolithic columns, makes this approach an appealing option for the design of capillary columns with high selectivities. 6.9.2 Pore size

The major advantage of CEC compared to classical HPLC is that much higher column efficiencies can be achieved using identical separation media. For columns packed with beads, the column efficiency of both of these methods is particle size dependent, and increases as the size of the packing decreases [ 1]. Since the monolithic columns are molded rather than packed, issues of particles size become irrelevant, and instead, the size of the pores within the monolithic material is the variable most

229

Monoliths

mAN

I (a)

keff(S ) = 5.51 mAU II(b) keff(R ) = 8.91 ] = 1.62 I

]

I I

]

(s) Rs

= .24

~ N(S) = 5 0 0 m - '

I /

I

II

LI

(s)

] keff(S) = 4.79

[ keff(R ) = 16.10

II ~

= 3.36

II Rs

II N(S)

(R)

0

20

= 7.2

= 4 , 0 0 0 m -~

=3,000m-'

(R)

40

0

20

40

60

70

Retention time (min) Fig. 6.29. Electrochromatographic performance of monoliths prepared by copolymerization of ethylene dimethacrylate and chiral monomer 25 with glycidyl methacrylate (a) and 2-hydroxyethyl methacrylate (b) as comonomers. (Reprinted with permission from [60]. Copyright 2000 American Chemical Society). Conditions: capillary column 335 mm (250 mm active length) x 0.1 mm i.d., pore size 993 nm (a) and 1163 nm (b), analyte DNB-(R,S)-leucine, mobile phase 400 mM acetic acid and 4 mM triethylamine in acetonitrile-methanol (80:20, v/v), 25 kV, temperature 30~

expected to affect chromatographic efficiency. Indeed, initial studies have shown that the size of the flow-through pores dramatically affects separation efficiency [54,64]. Consistent with our previous findings using systems consisting of butyl methacrylate, ethylene dimethacrylate, and 2-acrylamido-2-methyl-l-propanesulfonic acid [54,64,72], the column efficiency of quinidine-functionalized monolithic capillaries again clearly depends on the pore size. Fig. 6.30 illustrates that this holds for chiral monoliths prepared by either thermal or UV initiation [60]. As previously found for the reversed phase separations of alkylbenzenes, the effect of pore size on the separation of enantiomers is also rather complex and subtleties of these effects remain to be explored in more detail. We assume that the unusual shape of these curves might reflect changes in the overall morphology within the monoliths rather than the effect of chemical functionality since the monoliths used in the measurements shown in Fig. 6.30 are chemically identical. The mode pore sizes used for the plot, determined by mercury porosimetry, References pp. 238-240

Chapter 6

230

E 15000.

A

co

o.

10000

r r r

r

5000

b

E 0

0

0

|

0

I

1000

'

I

2000

'

I

'

3000

I

4000

Pore diameter (nm) Fig. 6.30. Column efficiencies for (S) enantiomer determined from DNZ-(R,S)-Leu separations on quinidine-functionalized monoliths as a function of pore diameter. (Reprinted with permission from [60]. Copyright 2000 American Chemical Society). Conditions: polymerization mixture, chiral monomer 8 wt%, 2-hydroxyethyl methacrylate 16 wt%, ethylene dimethacrylate 16 wt%, porogenic solvent 60 wt% (consisting of 1-dodecanol and cyclohexanol in different proportions), UV initiated polymerization for 16 h at room temperature (r-I) and thermally initiated polymerization for 20 h at 60~ (11), capillary columns 335 mm (250 mm active length) • 0.1 mm i.d., mobile phase 0.4 mol/L acetic acid and 4 mmol/L triethylamine in 80:20 acetonitrile-methanol, separation temperature 50~ voltage-25 kV.

indicate only the positions of the maxima (modes) of the pore size distribution curves rather than the frequency of occurrence of all of the pores. Monoliths with smaller and smaller modal pore sizes exhibit considerable differences in their microporous structure as confirmed by an increase in their specific surface areas. Deviations from a plug-like flow profile were found to occur in the smaller transport channels [1,73] leading to decreased efficiencies. These effects would also be expected to be more prevalent for the monoliths with smaller pore diameters.

6.9.3 Flow of the mobile phase Electroosmotic flow is generally reported to be independent of the size of the packing, and consequently the size of the interstitial voids between the particles, unless this size is so small that the electrical double layers overlap [74]. The ability to independently control both the pore size and level of charged functionalities of the methacrylate ester monolithic capillaries enables the direct investigation of the net effect of transport channel size on flow velocity. Recent results clearly demonstrates a

231

Monoliths

16(-

E E

>~ o...

m

o >

12

o q., r

c

. m

o

,

,

i

|

,

,

|

,

lo'oo . . . .

Pore size, nm

ls'oo

Fig. 6.31. Effect of mode pore diameter on flow velocity of the mobile phase through monolithic capillary columns. (Reprinted with permission from [55]. Copyright 1999 Wiley-VCH). Conditions: stationary phase poly(butyl methacrylate-co-ethylene dimethacrylate) with 0.3 wt. % 2-acrylamido-2-methyl-l-propanesulfonic acid; Column 100 ~tm i.d. • 28 cm; mobile phase, 75:25 vol/vol mixture of acetonitrile and 5 mmol/L phosphate buffer pH 7, marker thiourea. The line represents linear fit of experimental data.

considerable increase in flow velocity through the thermally initiated monolithic capillaries with the same level of charged moieties as the pore size increases from 250 nm to several micrometers [64]. A similar increase in flow velocity was observed for monoliths prepared by UV initiated polymerization [60,72]. Fig. 6.31 clearly demonstrates these effects. The range of pore sizes in question significantly exceeds the thickness of a few nanometers at which the electrical double layers would overlap for a system utilizing a mobile phase containing low molarity buffers [64]. Moreover, if it is assumed that the observed decrease in flow rate with decreasing mode pore diameter would simply result from the increasing percentage of pores within which overlap of the electric double layers occurs, then the flow velocity should reach a maximum value for those monoliths having sufficiently large pores, and remain constant thereafter, since the number of pores within which overlap of the electric double layer can occur decreases rapidly as the pore size increases. In practice however, this phenomenon is not observed. The fact that the overall flow velocity increases linearly over a broad range of pore sizes may also support the contention that this increase in flow rate is macroscopically related to a decrease in the resistance to flow through the channels. An additional effect may result from microscopic variations in the strength

References pp. 238-240

Chapter 6

232

Fig. 6.32. Schematic of flow path in monolithic separation media with large (a) and small (2) pores.

of the electrical field in both the small and large pores. The effects of tortuosity and variations in the cross sectional area of a packed structure on the conductance and chromatographic performance of CEC capillaries packed with beads that have recently been discussed in the literature [75] may also play a role. It is likely, that the flow path through a monolith with large pores is rather straight and therefore, the trajectory is short. In contrast, this path may be longer in monoliths with small pores, since the molecules passing through the pores make many detours around a larger number of clusters of microglobules. As shown in Fig. 6.32, the analyte must traverse a longer distance that then results in longer retention times even if the overall electroosmotic velocity does not change. 6.10 OTHER APPLICATIONS OF POROUS POLYMER MONOLITHS IN CEC COLUMN TECHNOLOGY 6.10.1 Immobilization of beads in porous polymer monolith

A number of different approaches were used for monolithization of capillary column packed with silica beads [76-82]. Chirica and Remcho employed sol-gel

Monoliths

233

transition of silicate solution located in interstitial voids of a packed capillary column [83]. Since control of the solid phase formation to avoid cracks is very tricky [78,81], Chirica and Remcho used in situ polymerization of organic monomers [63]. Although the preparation of monoliths by polymerization of a mixture of monomers and porous particles was described in an earlier patent [84], this was the first application of this approach to prepare CEC columns. The process consists of a few steps: First, a very porous temporary frit is sintered at one end of the capillary and the tube slurry packed with porous 5 ~tm octadecyl silica beads. Using positive pressure, the packed capillary is filled with the polymerization mixture consisting of ethylene dimethacrylate, azobisisobutyronitrile, 1-propanol, 1,4-butanediol, and water. The last three compounds are inert solvents added to achieve the required porosity for the flow through. Optionally, an alkyl methacrylate, where the alkyl is methyl, ethyl, or butyl, and 2-acrylamido-2-methyl-1propanesulfonic acid were also added. After 48 h polymerization at a temperature of 60~ both ends of the capillary including that with the original frit were cut to obtain column with a desired length. SEM micrograph in Fig. 6.33 clearly shows the porous polymer within the voids between silica particles. Since the column is packed across its entire length, a piece of open capillary with a detection window was coupled this monolithic column. An addition of polymerizable sulfonic acid to polymerization mixture is a suitable tool for the control of electroosmotic flow. For example, only 0.1% of 2-acrylamido-2-methyl-1-propanesulfonic acid in the polymer increases electroosmotic flow velocity by 10-15% thus accelerating the separations. Despite numerous efforts, use of polymerization mixtures with only ethylene dimethacrylate as a monomer led to immobilized columns with dramatically decreased retentions. Since these mixtures are likely to produce porous polymers with very small pores and the polymer fills also the native pores of silica beds, the access of analytes to the C18 functionalities is restricted. Substitution of approximately half of ethylene dimethacrylate in the mixture with modestly hydrophobic methyl methacrylate returns the retention characteristics back to those of the original C18 silica. However, peak shapes featuring severe tailing were monitored for highly hydrophobic polyaromatic hydrocarbons. Using butyl methacrylate as a comonomer eventually alleviated this problem. As has been shown elsewhere [66], copolymers of ethylene dimethacrylate and butyl methacrylate exhibit hydrophobicity index similar to that of C 18 silica and this match of polarities is favorable for separations in the reversed phase mode. Fig. 6.34 demonstrates the performance of a column prepared using this technology for the separation of a series of drugs.

References pp. 238-240

234

Chapter 6

Fig. 6.33. Scanning electron micrograph of a capillary column packed with 5 pm ODS silica beads and entrapped in porous poly(methyl methacrylate-co-ethylene dimethacrylate-co-2acrylamido-2-methyl-l-propanesulfonic acid). (Reprinted with permission from [63]. Copyright 2000 American Chemical Society).

6.10.2 Monolithic frits

In addition to utilization of monoliths as a column material, two reports describing respectively silicate and synthetic organic polymer based monolithic frits were published recently [85,86]. The conventional method of frit fabrication for a particle packed column usually involves thermal sintering of a section of the packing material, such as bare or octadecyl silica, using a heating device. This approach has several weaknesses such as the lack of control of the temperature and porous properties of the flit that decreases reproducibly of the fabrication process. In contrast, photoinitiated free radical polymerization of glycidyl methacrylate and trimethylolpropane trimethacrylate in the presence of porogenic solvent affords a monolithic plug within the column that serves as a frit. This procedure represents a simple approach to reproducible fabrication of frits even in capillaries with large inner diameters.

23 5

Monoliths

;k- 254 nm

e-,

L

,_____.

~ 220 nm

6

< 2 3

0

I 2

I 4

7

I 6

I 8

min

Fig. 6.34. CEC separation of drugs using a capillary column packed with 5 mm ODS silica beads and entrapped in porous poly(butyl methacrylate-co-ethylene dimethacrylate-co-2acrylamido-2-methyl-l-propanesulfonic acid). (Reprinted with permission from [63]. Copyright 2000 American Chemical Society). Conditions: column 26 cm (active length 17 crn) x 75 p,m i.d, mobile phase 70:30 acetonitrile-10 mmol/L acetate buffer solution pH 3, 20 kV, detection at 2 different wavelengths shown.

The initiation by UV irradiation facilitated localization of the polymer plug in the capillary column. An outlet frit was prepared first by introducing the monomer mixture into the capillary from which about 2 mm section of the polyimide coating was removed and its ends were sealed. The capillary was then masked by aluminum foil, leaving 1 mm of the section without coating exposed to the UV light. The polymerization process was finished in about 1 h, the unreacted monomer solution flushed from the tube, and the column packed with 1.5 lam octadecyl silica beads. Similar procedure was employed to form the inlet frit. Although no specific functionalization of the inner capillary surface preceding the polymerization was carried out, the outlet frit easily withstand a short exposure of a pressure as high as 400 MPa used during the column packing. This demonstrates that the 1 mm thick monolithic frit is strongly bound to the inner surface of the bare capillary wall. Fig. 6.35 shows the inlet frit structure with embedded 1.5 gm beads. This micrograph also suggests that the photopolymer does not exhibit the microglobular morphology typical of standard macroporous polymers (vide supra) if formed in the presence of the silica beads. The silica particles are entrapped in the pores and hold within their domains.

References pp. 238-240

236

Chapter 6

Fig. 6.35. SEM micrograph of a cross section of a photopolymerized 1 mm long inlet frit with embedded 1.5 ~tm ODS beads within a 75 gm i.d. capillary. (Reprinted with permission from [86]. Copyright 2000 American Chemical Society).

In contrast to some other procedures, the UV photoinitiated polymerization does not require elevated temperature for the reaction to be completed. Therefore, the mobile phase used for packing remains in both the outlet frit and the packing during polymerization of the inlet frit. Consequently, the conditioning time for the column prior to its use is shortened significantly. No bubble formation was observed while using packed capillary columns with photopolymer frits. A systematic study of the run-to-run reproducibility for the analysis of a mixture of neutral molecules consisting of thiourea, benzyl alcohol, benzaldehyde, and 2-methylnaphthalene was carried out over a period of 3 days. Figure 6.36 shows electrochromatograms of runs 10, 30, and 50. There is almost no variation in retention times of all test compounds. The relative standard deviations for retention factor, the efficiency, and the resolution, averaged over 60 runs, are 3.5%, 3.3%, and 5.5%, respectively. The integrity of the packed column remained unchanged, and the column

237

Monoliths

Run 10

r Run 30

r

r

~J r

Run 50

/x..__ I 2

,

I 4

I

[

6

I

I 8 min

Fig. 6.36. Selected representative CEC electrochromatograms from 60 runs using column with monolithic flits. (Reprinted with permission from [86]. Copyright 2000 American Chemical Society). Conditions: mobile phase 40% (v/v) 5 mM phosphate buffer with addition of 2 mmol/L SDS (pH 7.0) and 60% (v/v) acetonitrile. Peaks: thiourea, benzyl alcohol, benzaldehyde, and 2-methylnaphthalene (order of elution).

is assumed to be useful for a much longer period of time. The column efficiencies calculated from peaks of thiourea, benzyl alcohol, benzaldehyde, and 2-methylnaphthalene shown in Figure 6.36, are 200 000, 160 000, 60 000, and 20 000 plates/m, respectively. 6.11 ACKNOWLEDGMENT I would like to thank my friend and colleague at the University of California at Berkeley, Professor Jean M. J. Fr6chet for his endless support and helpful advice, and co-workers Dr. Cong Yu, Dr. Michael L~immerhofer, Dr. Eric C. Peters, Dr. Miroslav Petro, Dr. David S3~kora for their most valuable contributions to the research and

References pp. 238-240

238

Chapter 6

development of monolithic CEC columns that are listed in the references. Support of our research by grant of the National Institute of General Medical Sciences, National Institutes of Health (GM-48364) is gratefully acknowledged. Our work was also partly supported by the Division of Materials Sciences of the U.S. Department of Energy under Contract No. DE-AC03-76SF00098. 6.12 R E F E R E N C E S

1 J.H. Knox, J. Chromatogr. A, 680 (1994) 3. 2 M. Kubin, P. ~;pa6ek and R. Chrome~ek, Coll. Czechosl. Chem. Commun., 32 (1967) 3881. 3 W.D. Ross and R.T. Jefferson, J. Chromatogr. Sci., 8 (1970) 386. 4 F.D. Hileman, R.E. Sievers, G.G. Hess and W.D. Ross, Anal. Chem., 45 (1973) 1126. 5 H. Schnecks and O. Bieber, Chromatographia, 4 (1971) 109. 6 T.R. Lynn, D.R. Rushneck and A.R. Cooper, J. Chromatogr. Sci., 12 (1974) 76. 7 D. Josi6 and A. ~;trancar, Ind. Eng. Chem. Res., 38 (1999) 333. 8 K.H. Hamaker, S.L. Rau, R. Hendrickson, J. Liu, C.M. Ladish and M.R. Ladish, Ind. Eng. Chem. Res., 38 (1999) 865. 9 S. Hjert6n, Ind. Eng. Chem. Res., 38 (1999) 1205. 10 F. Svec and J.M.J. Fr6chet, Ind. Eng. Chem. Res., 36 (1999) 34. 11 S.M. Fields, Anal. Chem., 68 (1996) 2709. 12 H. Minakuchi, K. Nakanishi, N. Soga, N. Ishizuka and N. Tanaka, Anal. Chem., 68 (1996) 3498. 13 F. Svec, E.C. Peters, D. S3~kora, C. Yu and J.M.J. Fr6chet, HRC-J. High Resol. Chromatogr., 23 (2000) 3. 14 F. Svec, E.C. Peters, D. S~kora and J.M.J. Fr6chet, J. Chromatogr. A, 887 (2000) 3. 15 C. Fujimoto, Anal. Chem., 67 (1995) 2050. 16 C. Fujimoto, J. Kino and H. Sawada, J. Chromatogr. A, 716 (1995) 107. 17 N. Tanaka, K. Nakagawa, H. Iwasaki, K. Hosoya, K. Kimata, T. Araki and D.G. Patterson, J. Chromatogr. A, 781 (1997) 139. 18 B. Maichel, B. Poto6ek, B. Ga~, M. Chiari and E. Kenndler, Electrophoresis, 19 (1998) 2124. 19 B. Maichel, B. Poto6ek, B. Ga~ and E. Kenndler, J. Chromatogr. A, 853 (1999) 121. 20 B. Poto6ek, E. Chmela, B. Maichel, E. Tesa~ovfi, E. Kenndler and B. Ga~, Anal. Chem., 72 (2000) 74. 21 B. Maichel, B. Ga~ and E.Kenndler, Electrophoresis, 21 (2000) 1505. 22 M.R. Schure, R.E. Murphy, W.L. Klotz and W. Lau, Anal. Chem., 70 (1998) 4985. 23 N. Tanaka, T. Fukutome, K. Hosoya, K.Kimata and T.Araki, J. Chromatogr. A, 716 (1995) 57. 24 N. Tanaka, H. Iwasaki, T. Fukutome, K. Hosoya and T. Araki, HRC- J. High Resolut. Chromatogr., 20 (1997) 529. 25 B. Poto6ek, B. Maichel, B. Ga~, M. Chiari and E. Kenndler, J. Chromatogr. A, 798 (1998) 269. 26 Y. Baba and M. Tsuhako, YrAC-Trends Anal. Chem., 11 (1992) 280. 27 C. Fujimoto, Analusis, 26 (1998) M49.

Monoliths

23 9

28 S. Hjert6n, D. Eaker, K. Elenbring, C. Ericson, K. Kubo, J.L. Liao, C.M. Zeng, P.A. Lindstr6m, C. Lindh, A. Palm, T. Srichiayo, L. Valcheva and R. Zhang, Jpn. J. Electrophor., 39 (1995) 105. 29 C. Ericson, J.L. Liao, K. Nakazato and S. Hjert6n, J. Chromatogr. A, 767 (1997) 33. 30 J i . Liao, N. Chen, Ch. Ericson and S. Hjert6n, Anal. Chem., 68 (1996) 3468. 31 C. Ericson and S. Hjert6n, Anal. Chem., 71 (1999) 1621. 32 D. Hoegger and R. Freitag, HPCE 2000, 13th International Symposium on High Performance Capillary Electrophoresis and Related Microscale Techniques, Saarbri.icken, Germany, February 20-24, 2000, Abstracts p.230. 33 C. Ericson, J. Holm, T. Ericson and S. Hjert6n, Anal. Chem., 72 (2000) 81. 34 J. Seidl, J. Malinsk~,, K. Du~ek and W. Heitz, Adv. Polym. Sci., 5 (1967) 113. 35 A. Palm and M.V. Novotny, Anal. Chem., 69 (1997) 4499. 36 P.G. Righetti, J. Chromatogr. A, 698 (1995) 3. 37 A.H. Que, T. Konse, A.G. Baker and M. Novotny, Anal. Chem., 72 (2000) 2703. 38 A.H. Que, A. Palm, A.G. Baker and M. Novomy, J. Chromatogr. A, 887 (2000) 379. 39 N.W. Smith and M.B. Evans, Chromatographia, 41 (1995) 197. 40 G. Wulff, Angew. Chem.,Intl. Ed. Engl., 34 (1995) 1812-1832. 41 S. Nilsson, L. Schweitz and M. Petersson, Electrophoresis, 18 (1997) 884. 42 L. Schweitz, L.I. Andersson and S. Nilsson, J. Chromatogr. A, 792 (1997) 401. 43 L. Schweitz, L.I. Andersson and S. Nilsson, Anal. Chem., 69 (1997) 1179. 44 L. Schweitz, L.I. Andersson and S. Nilsson, J. Chromatogr. A, 817 (1998) 5. 45 L. Schweitz, L.I. Andersson and S. Nilsson, Chromatographia, 49 (1999) $93. 46 J.M. Lin, T. Nakagama, K. Uchiyama and T. Hobo, Chromatographia, 43 (1996) 585. 47 J.M. Lin, T. Nakagama, K. Uchiyama and T. Hubo, Biomed. Chromatogr., 11 (1997) 298. 48 J.M. Lin, T. Nakagama, X.Z. Wu, K. Uchiyama and T. Hobo, Fresenius J. Anal. Chem., 357 (1997) 130. 49 I. Gusev, X. Huang and C. Horvfith, J. Chromatogr. A, 855 (1999) 273. 50 X. Huang, J. Zhang and C. Horvfith, J. Chromatogr. A, 858 (1999) 91. 51 B.H. Xiong, L.H. Zhang, Y.K. Zhang, H.F. Zou and J.D. Wang, HRC-J. High Resolut. Chromatogr., 23 (2000) 67. 52 F. Svec and J.M.J. Fr6chet, Anal. Chem., 54 (1992) 820. 53 F. Svec and J.M.J. Fr6chet, Science, 273 (1996) 205. 54 E.C. Peters, M. Petro, F. Svec and J.M.J. Fr6chet, Anal. Chem., 69 (1997) 3646. 55 C. Yu, F. Svec and J.M.J. Fr6chet, Electrophoresis, 21 (2000) 120. 56 M. L~immerhofer, E.C. Peters, C. Yu, F. Svec, J.M.J. Frdchet and W. Lindner, Anal. Chem., 72 (2000) 4623. 57 Q. Wang, F. ~;vec and J.M.J. Fr6chet, Anal. Chem., 65 (1993) 2243. 58 C. Viklund, F. Svec, J.M.J. Fr6chet and K. Irgum, Chem. Mater., 8 (1996) 744. 59 F. Svec and J.M.J. Frdchet, Macromolecules, 28 (1995) 7580. 60 M. L~immerhofer, F. Svec, J.M.J. Fr6chet and W. Lindner, Anal. Chem., 72 (2000) 4614. 61 F.E. Regnier, Nature, 350 (1991) 643. 62 D.K. Roper and E.N. Lightfoot, J. Chromatogr. A, 702 (1995) 3. 63 G.S. Chirica and V.T. Remcho, Anal. Chem., 72 (2000) 3605. 64 E.C. Peters, M. Petro, F. Svec and J.M.J. Fr6chet, Anal. Chem., 70 (1998) 2288.

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65 66 67 68 69 70 71

P.B. Wright, A.S. Lister and J.G. Dorsey, Anal. Chem., 69 (1997) 3251. E.C. Peters, M. Petro, F. ~;vec, and J.M.J. Fr6chet, Anal. Chem., 70 (1998) 2296. E. Venema, J.C. Kraak, H. Poppe and R.Tijssen, Chromatographia, 48 (1998) 347. E. Venema, J.C. Kraak, H. Poppe and R. Tijssen, J. Chromatogr. A, 837 (1999) 3. R. Stol, W.T. Kok and H. Poppe, J. Chromatogr. A, 853 (1999) 45. C. Fujimoto, Y. Fujise and E. Matsuzawa, Anal. Chem., 68 (1996) 2753. E.C. Peters, K. Lewandowski, M. Petro, F. ~;vec and J.M.J. Fr6chet, Anal. Commun., 35(1998) 83. C. Yu, F. Svec and J.M.J. Fr6chet, Electrophoresis, 21 (2000) 120. D. Li and V.T. Remcho, J. Microcol. Sep., 9 (1997) 389. J.H. Knox and I.H. Grant, Chromatographia, 32 (1991) 317. G. Choudhary and C. Horv~th, J. Chromatogr. A, 781 (1997) 161. M.M. Dittman, G.P. Rozing, G. Ross, T. Adam and K.K.Unger, J. Capil. Electroph., 5 (1997) 201. R. Asiaie, X. Huang, D. Faman and C. Horv~th, J. Chromatogr. A, 806 (1998) 251. M.T. Dulay, R.P. Kulkami and R.N. Zare, Anal. Chem., 70 (1998) 5103. Q.L. Tang, B.M. Xin and M.L. Lee, J. Chromatogr. A, 837 (1999) 35. S.A. Rodriguez and L.A. Colon, Anal. Chim. Acta, 397 (1999) 207. C.K. Ratnayake, C.S. Oh and M.P. Henry, HRC-J. High Resolut. Chromatogr., 23 (2000) 81. Q.L. Tang and M.L. Lee, HRC- J. High Resolut. Chromatogr., 23 (2000) 73. G. Chirica and V.T. Remcho, Electrophoresis, 20 (1999) 50. J.M.J. Fr6chet and F. Svec, US patent 5,334,310 (1994). Y. Chen, G. Gerhardt and R. Cassidy, Anal. Chem., 72 (2000) 610. J.R. Chen, M.T. Dulay, R.N. Zare, F. Svec and E.C. Peters, Anal. Chem., 72 (2000) 1224.

72 73 74 75 76 77 78 79 80 81 82 83 84 85 86

Chapter 7

Open Tubular Approaches to Capillary Electrochromatography r162

Joseph J. P E S E K

and M a r i a T. M A T Y S K A

Department of Chemistry, San Jose State University, One Washington Square, San Jose, CA 95192, USA

CONTENTS

7.1 7.2 7.3 7.4

7.5

7.6 7.7 7.8

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C h e m i c a l etching process . . . . . . . . . . . . . . . . . . . . . . . . . C h e m i c a l modification process . . . . . . . . . . . . . . . . . . . . . . . Characterization o f etched, c h e m i c a l l y m o d i f i e d capillaries . . . . . . . . 7.4.1 Scanning electron m i c r o s c o p y . . . . . . . . . . . . . . . . . . . 7.4.2 A t o m i c force m i c r o s c o p y . . . . . . . . . . . . . . . . . . . . . 7.4.3 Diffuse reflectance infrared fourier transform spectroscopy . . . 7.4.4 Electron spectroscopy for chemical analysis . . . . . . . . . . . 7.4.5 E l e c t r o o s m o t i c flow m e a s u r e m e n t s . . . . . . . . . . . . . . . . Applications of OTCEC . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5.1 P h a r m a c e u t i c a l s and other small m o l e c u l e s . . . . . . . . . . . . 7.5.2 Peptides and proteins . . . . . . . . . . . . . . . . . . . . . . . 7.5.3 Chiral c o m p o u n d s . . . . . . . . . . . . . . . . . . . . . . . . . 7.5.4 Stability and reproducibility . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

242 244 245 247 249 250 253 255 256 257 258 260 263 264 266 267 268

242

Chapter 7

7.1 INTRODUCTION Applications and development of micro-separation techniques have expanded greatly in the last decade for a number of practical reasons. Among these are the reduced amount of sample needed, particularly important in biological and pharmaceutical applications, the lower volume of solvents used in the analysis, and the ease in which these methods can be interfaced to a mass spectrometer for a detector. Capillary electrochromatography (CEC) is one of the newest micro(~t)-separation formats available. In CEC there are two general approaches to this hybrid technique that combines features of both high performance liquid chromatography (HPLC) and capillary electrophoresis (CE): the packed column configuration that utilizes stationary phases similar to HPLC and the open tubular format where the stationary phase is immobilized on the capillary wall. This chapter focuses on the latter approach with an emphasis on open tubular columns that are fabricated by first etching the inner wall of the fused silica tube. A summary of the continuum of techniques that exist between ~t-HPLC and CE that can be used in the micro-column format are shown in Fig. 7.1. It can be seen that it is possible to make a smooth transition from Ia-HPLC to packed capillary CEC by adjusting the ratio of pressure-driven flow to electrically-driven flow. This adjustment results in an infinite number of hybrid techniques that can be generated from pure ~t-HPLC (no electric field component) to pure packed capillary CEC (no pressure component). The transition region between CE and packed capillary CEC is not so smooth. The bridge is provided by open tubular capillary electrochromatography (OTCEC). There are not, however, the infinite number of choices that can be generated as in the bridge between ~t-HPLC and packed capillary CEC. However, some variations exist by utilizing an etching process, which results in both an increase in surface area and radial extensions of various types and lengths from the surface, as well as the diameter of the capillary that can create a significant array of column formats. The figure also denotes on a relative scale the importance of the major separation mechanisms, solute/bonded phase interactions (as measured by the capacity factor, k') and electrophoretic mobility (laep), that exist in the continuum with k' being dominant at the ~t-HPLC end and ~tep being the only factor in CE. The wall of a fused silica capillary, containing silanol groups, has been identified as a potential site of interaction for solutes in electrophoretic measurements. In particular, adsorption of proteins, peptides and other basic compounds has been more specifically addressed [1-3]. In the ideal situation, the interaction is strictly chromatographic so that the process is reversible with the net result being a loss in the observed efficiency for CE [2,4]. A more serious problem can result when there is irreversible adsorption of the analyte on the capillary wall leading to a lack of repro-

243

Open Tubular C E C

~t-HPLC r

Voltage Assisted r p -HPLC

LAMINAR FLOW

~

~

Pressure Assisted CEC

~

~

~

~

Packed Capillary CEC

~

ELECTRICALL Y DRIVEN FLOW

<::> O T C E C

r

CE

~- ~-- +- ~-- ~- Increasing influence of solute/bonded phase interactions (k') Increasing influence of electrophoretic mobility (~ep) --~ --~ -+ --~ --+ Fig. 7.1. Schematic that summarizes the relationship between the various micro-separation techniques. Line 1 lists the various micro-separation techniques. Below each method in line 2 is the type of flow or combination of flow ( ~ ) that exists for each technique while in line 3 is the relative contribution of solute/bonded phase interactions and electrophoretic mobility to the separation mechanism.

ducibility in crucial measurement parameters such as elution time, efficiency and even mobility. If the interaction is reversible, the specific solute/wall effect can be measured in well-designed capillary chromatographic experiments. Such measurements have been made in both the zonal [5,6] and frontal [7,8] chromatographic modes. The retention factor determined from these experiments can be used to determine the magnitude of the chromatographic interaction that appears in the efficiency term for CE [2,4,6]. The first effective OTCEC separations indicating chromatographic interactions were demonstrated a number of years ago by Tsuda and co-workers [9] using an octadecyl modified 30 pm i.d. capillary. A more definitive way of proving that chromatographic effects are possible in the open tubular format can be provided through the separation of optical isomers. A number of chiral selectors such as cyclodextrin and several cellulose derivatives that were bonded to the inner wall of fused silica capillaries resulted in the separation of enantiomers [ 10-15]. Since the two optical isomers have identical electrophoretic mobilities, separation can only be achieved through differences in solute/bonded phase interactions. It has also been demonstrated that molecularly imprinted polymers bonded to the inner wall of a fused silica capillary can be another approach for a stationary phase in OTCEC [ 16]. Each of these early studies showed that OTCEC was feasible at least for the limited number of samples and experimental conditions tested. Initially, further exploitation of the OTCEC technique was hampered by two fundamental problems that seriously limited its potential as a viable separation method: the low capacity of the column due to the small area available for bonding a stationary phase and the long distance that molecules would have to migrate to interact with the References pp. 268-270

244

Chapter 7

bonded moiety. The latter problem could be addressed by reducing the column i.d. but this is often an unsatisfactory solution because such a decrease further limits the sample size as well as the detection path length in optical systems and hence the detection limit of the analyte. The general approach described below attempts to overcome these two major problems though chemical etching of the inner wall of the capillary. This process produces structural changes within the capillary that are more favorable for OTCEC. First, the etching process increases the overall surface area of the capillary, perhaps by as much as 1000-fold [17], which allows more stationary phase to be attached to the wall thus increasing the capacity of the column. Second, the dissolution and redeposition of silica material during etching creates radial extensions from the wall that decrease the distance a solute must travel in order to interact with the stationary phase. This aspect is especially important for separations of biomolecules where the diffusion coefficient is much lower than for typical small organic molecules or inorganic ions. 7.2 CHEMICAL ETCHING PROCESS The etching process as described in this section was patterned after the original work of Onuska and co-workers for gas chromatography [17] but was modified to accommodate the smaller inner diameters needed for OTCEC [ 18]. The general procedure starts with an approximate 2 m section of bare capillary having a 375 ~tm o.d. and a 20 or 50 lam i.d. that is filled with concentrated HC1, sealed and heated overnight at 80~ The tube is then flushed successively with deionized water, acetone and diethyl ether. The tube is dried for 1 h under nitrogen flow at ambient temperature. The capillary is then filled with a 5% (w/v) solution of ammonium hydrogen fluoride in methanol and allowed to stand for 1 h. The methanol is then removed by nitrogen flow for 0.5 h. After the capillary is sealed at both ends, it is heated in a modified gas chromatography (GC) oven at temperatures between 300 to 400~ for a period of 3 to 4 h. The GC oven provides good temperature control, rapid heating and cooling, and easy accessibility for the lengths of capillary used in this procedure. The exact time and temperature chosen determines the surface morphology that is obtained on the inner wall [18,19]. The general trend observed is that for shorter etching periods (- 2-3 h) at lower temperatures (-300~

the inner wall consists of relatively long

spikes of silica material protruding from the surface. For longer etching periods (- 4h) at higher temperatures (-400~ the structure has fewer long extensions from the surface and becomes somewhat more regular either resembling sand dunes or a sponge-like porous configuration. More detailed studies by atomic force microscopy (AFM) and scanning electron microscopy (SEM) for characterizing etched and etched chemically modified capillary surfaces are described below. Estimates from the AFM

245

Open Tubular CEC

measurements indicate that the area of the inner wall is increased significantly (100 1000 fold), which should facilitate solute/bonded phase interactions after appropriate organic moieties are attached to this etched surface. Additional measurements of the surface area have been made by the BET (Brunauer-Emmet-Teller) nitrogen adsorption method. The data was obtained on 1 cm segments of etched 50 gm capillary with a total length of 2.6 m. The surface area measured was 1 x 10-1 m2/g. The same length of unetched capillary should have a surface area of approximately 8 x 10-5 m2/g. Therefore, the BET and AFM measurements are in good agreement showing an increase in surface area by about a factor of 1000. Additional useful surface morphologies may be possible through manipulation of the various experimental variables involved in the etching process, i.e. etching time, etching temperature, reagent concentration, capillary diameter and capillary configuration in the oven. 7.3 C H E M I C A L M O D I F I C A T I O N P R O C E S S After etching, bonding of an organic moiety to the etched inner wall for the stationary phase takes place using the silanization/hydrosilation process adapted for the capillary format [ 18-20]. This approach for attaching a variety of organic moieties to silica involves first silanizing the surface by reaction with triethoxysilane (TES) to create a hydride intermediate [21 ] as shown in equation 7.1. SILANIZATION OY H+

I

~ Si-OH + (OEt)sSi-H

~

~ ,g

(7.1)

Si-O-Si-H + nEtOH

I OY

First the capillary is treated with a pH 10 ammonia solution (6 mM) for 20 h at a flow rate of 0.1-0.2 mL/h. The capillary is then rinsed with deionized water followed by a wash with 0.1 M HC1 and then a second rinsing with water. The tube is next dried with nitrogen, filled with dioxane and then flushed with a 1.0 M TES solution in dioxane containing HC1 as a catalyst for 90 min at 90~

After the TES treatment, the

capillary was washed with tetrahydrofuran (THF) for 2 h and then with a 1:1 THFwater mixture for 2 h. The capillary is then dried with a flow of nitrogen gas for 0.5 h. The capillary, which now has a hydride surface, is then flushed with dry toluene in preparation for the second modification.

References pp. 268-270

Chapter 7

246

The next step is the hydrosilation reaction that is used to attach an organic moiety [22,23]. An olefin/catalyst (usually hexachloroplatinic acid) solution (pure olefin or olefin dissolved in toulene) is heated to 60-70~

for a period of 1 h. The hydride

capillary filled with toluene is then flushed with the reacting solution for a period of 45 h at 100~

After completion of the hydrosilation reaction, the capillary is washed

successively with toluene and THF for 1 h. After the washing step, the capillary is then dried at 100~ under a nitrogen flow overnight. Normally hydrosilation utilizes as a terminal olefin for creating a Si-C bond between the silicon containing reactant and the organic compound [24] as shown in equation 7.2. HYDROSILATION

I

Si-H

cat.-- I~ S i - C H 2

+ R - C H m CH2

m CH2mR

(7.2)

c a t - catalyst, typically hexachloroplatinic acid (Speier's catalyst) However, it has already been demonstrated that other unsaturated groups (alkynes and cyano) as well as olefins in a nonterminal position can undergo hydrosilation on a silica hydride surface [25,26]. The cyano group is amenable to hydrosilation only in the absence of an olefin and with a free radical initiator as the catalyst giving two possible products as shown below [27].

/0 Ar m

~ N ~Si ~ 0 \

0

O\ / 0

/$i~ Ar

\

/ Si

/\ 0

0

247

Open Tubular CEC

We have also used free radical initiators and other transition metal complexes besides Speier's catalyst to attach some olefins to the silica hydride intermediate [26,28]. The silanization/hydrosilation method is a very versatile approach for attaching a wide variety of organic moieties to silica surfaces. This includes porous silica particles for HPLC and packed capillary electrochromatography, and to the inner walls of fused silica tubes for use in CE and OTCEC. Another advantage of the silanization/ hydrosilation method is that most of the readily accessible silanols are eliminated creating a surface that is particularly suited for the analysis of biomolecules and pharmaceuticals. Finally, when alkenes or alkynes are used in the hydrosilation reaction, bonding to the silica surface results in a highly stable Si-C linkage. The type of bonded phase and hence the specific solute/bonded phase interactions that can be exploited for any particular separation depends on the nature of the organic moiety attached to the surface. The types of groups bonded to the etched inner wall of fused silica capillaries have been hydrophobic (octadecyl, C18) [18-20,29,30], hydrophilic (diol) [30], chiral [31] and liquid crystals [32]. The structures of the molecules attached to the silica hydride surface are shown in Fig. 7.2. In some cases linkers are first bonded to the desired stationary phase moiety before hydrosilation. The versatility of the silanization/hydrosilation process that was proved for the synthesis of bonded phases for HPLC can be readily adapted to OTCEC. As will be shown below, the etching process does make some contribution to the overall chromatographic/electrophoretic behavior of the separation process in OTCEC using the etched chemically modified capillaries. A summary of the complete process for producing OTCEC columns by the two steps (etching and chemical modification) described above is shown in Fig. 7.3. 7.4

CHARACTERIZATION

OF

ETCHED,

CHEMICALLY

MODIFIED

CAPILLARIES

A number of different approaches have been developed for characterizing the etched capillaries. Just as the development of methods for characterizing chemically bonded stationary phases has been useful in understanding their performance, a similar strategy is essential for future development of these separation materials as well as the identification of appropriate applications for OTCEC. In contrast to the porous particles typically used in HPLC where the chemically modified surface is readily accessible to various spectroscopic techniques [33,34], probing the inner wall of a fused silica capillary is not a straightforward process. In most cases, it is necessary to open the capillary so that the inner surface is exposed. This is not a simple procedure. It usually involves carefully breaking the capillary along the longitudinal axis and then examining the fragments under an optical microscope in order to select the most

References pp. 268-270

248

Chapter 7

HYDROPHOBIC: Octadecene CH2~--CH- CH2-- (CH2)I4~CH3

HYDROPHILIC" Diol OH OH

I

I

CH2~-CH- (CH2)4--CH--CH2

CHIRAL: R(+)-l,-(a - naphtyl)ethylamine 0 CH3 Cyclodextrin

..H O"

OH

O

I

II

1~= ~ C H 2 - - C H - C H 2 ~ O ~ C ~ C - - C H 2

I

CH3

OR' LIQUID CRYSTALS: Cholesteryl l O-Undecanoate

C ~ : ~- - 'H O

\

CH3

C H 2 : C H - CH2-- (CH2)6 ~ C H 2 - - C -O

I~I

H 4-cyano-4Ln-pentoxybiphenyl (M) C 5H l 1 - - O ~ ~ ~ ~ -

\~-~/

~

CN

Fig. 7.2. Structures of molecules bonded to the etched inner walls of fused capillaries by the silanization/hydrosilation process. In some cases a linker molecule has been attached to the primary separation moiety in order to facilitate bonding to the hydride surface.

249

Open Tubular CEC

CEC Capillary Derivatization . . . . . . . . . . . . . . . :........

--

-

-

native

~

ylVv

,v

~,

etched

~

yIV"

v-

Vv

hydrided

~

IFlV"

,v

V'

alkylated

I

50

~.tm

20 pm

or

NH4HF2

silanization: SiH(OEt)3

hydrosilation (alkylation): RCH=CH2 + catalyst

Fig. 7.3. Schematic representation of the processes for producing etched, chemically modified capillaries for OTCEC.

suitable samples for a particular characterization technique. Useful fundamental information about the properties of the etched chemically modified surface has been obtained from the five methods described below.

7.4.1 Scanning Electron Microscopy (SEM) SEM provides a picture of the surface morphology that exists on the inner wall of the capillary. This image is particularly useful for comparing the original smooth wall to the greatly roughened surface after etching as well as for comparing the effects of various etching conditions with respect to the type of new three-dimensional structures formed. The sample used is obtained as described above since the inner surface must be accessible to the probing electron beam. After a suitable sample is identified, the surface is gold plated by a sputtering process in order to make the capillary conductive. A typical SEM image showing the inner wall of a fused silica capillary after etching is shown in Fig. 7.4. A variety of 20 and 50 pm etched capillaries have been examined by SEM with respect to trying to correlate the surface morphology obtained as a function of the etching conditions [18,19]. The structures as described above in the section on the ammonium hydrogen difluoride process indicate that for short etching periods, at least two hours and at a temperature of 300~

the surface

appears to have the greatest amount of roughness and longer radial extensions from the surface. Using longer etching times and/or higher temperatures generally results in a diminishing of these features as the original extensions from the inner wall "melt away to a less tortuous morphology. At best, SEM provides a good visual image of the surface but only a semi-quantitative description of the overall roughness that results from the etching process.

Referencespp. 268-270

250

Chapter 7

Fig. 7.4. Scanning electromicrograph of the inner wall of an etched fused silica capillary.

7.4.2 Atomic Force Microscopy (AFM) AFM offers an alternative to SEM in providing a topological description of the etched, chemically modified surface. The two methods in combination provide complementary and confirming characterization of the inner wall that can be used to help understand the separation process in OTCEC using this format. The advantages of AFM over SEM include the ease of sample preparation because a conducting surface is not required, more precise determination of surface roughness from higher resolution in the z-direction (access to height information that is not available in SEM), and the ability to measure localized surface forces in order to identify possible solute/bonded phases interactions. The disadvantages of AFM for the characterization of OTCEC capillaries center on the difficulties in physically accessing regions of convoluted surfaces and uncertainties in rendering non-planar surfaces. The latter problem arises from contributions of the scanning piezo with respect to its movement over a curved surface. However, earlier AFM studies [35,36] have shown that some of the problems associated with non-planar and non-uniform surfaces of capillaries coated with polyacrylamide on the inner wall can be overcome. A comparison of SEM and AFM images of the same surface for the inner bore of native, etched, and etched chemically modified capillaries revealed a good visual

251

Open Tubular CEC

native

etched

hydride

chem-mod

Fig. 7.5. Atomic force microscopy (AFM) images in 3D recorded at two scan sizes of the four stages involved in the etching and chemical modification of capillaries for OTCEC.

correspondence between the electron and force microscopy data [37]. However, there are resolution differences between the two techniques. Subtle features revealed in AFM images require high magnification by SEM that limits the scan size to areas too small to observe the larger surface components. Some examples of the images obtained by AFM of each stage in the production of etched chemically modified capillaries are shown in Fig. 7.5. Two different scan sizes are displayed for the native, etched, etched-hydride and etched-chemically modified capillaries. The higher magnification (smaller scan size) allows for the examination of the more subtle features on each surface. Both scan sizes reveal that the bare capillary is relatively flat without any prominent features while the etched surface displays significant roughening with fairly sharp extensions. The roughness seems to increase further upon silanization to create a hydride layer. The rugged nature of the hydride surface may be due to the fact that more than a single layer of silane may have been deposited during this process. The micrographs of the alkylated surface reveal that considerable filling in and smoothing occurs during the hydrosilation process. Both AFM and SEM can easily detect the differences in surface morphology present in the four types of capillaries that are relevant to this approach to OTCEC. Besides images of the surface, AFM provides additional information that makes it unique as a surface characterization method for the etched, chemically modified capillaries. These measurement features include root means square (RMS) roughness,

References pp. 268-270

252

Chapter 7

surface area and surface-tip forces of attraction. When these AFM measurements are made on the four types of capillaries involved in the fabrication of etched, chemically modified columns for OTCEC, several crucial observations can be made about these unique separation materials. The ordering of the various surfaces for the RMS roughness measurements is native < etched ~ alkylated < hydride. The data is presented graphically in Fig. 7.6. For the surface area determinations the order is as follows:

NATIVE

40

,,,

ETCHED 30

, ,

20

0

40 HYDRIDED 30

. . . . . . .

20 10

li.....

0

.

m i

ALKYLATED

,,,

I

i

ml l,m,,_. 5

.

15

25

35

45

55

RMS ROUGHNESS, nm

65

75

Fig. 7.6. RMS roughness measurements as determined by AFM for the four types of capillaries involved in the etching and chemical modification processes.

Open Tubular CEC

253

native < etched < alkylated < hydride. A close agreement between these two parameters might be expected when comparing surfaces that change as a result of etching and chemical modification since an increase in the surface area should closely parallel an increase in the roughness. The actual order is interesting and might be explained by the following reasoning. The etched surface is greater than the native surface as expected from the etching process. The increase in going from the etched to the hydride surface may be the result of some polymerization of the silanization reagent, TES, giving rise to irregular growths. The smoothing of the hydride surface upon alkylation may be due to the infilling of the pits by alkyl groups attached during hydrosilation [38]. The relative magnitudes of the surface-tip forces of attraction are in the following order: native ~ alkylated ~ hydride I < etched < hydride II. The hydride surfaces have two distinct populations of forces so they are separated in the ordering scheme shown above. The ordering of the surface-tip forces can be qualitatively explained by the relative reactivities of the four surfaces. The native surface has been exposed to atmospheric condition over a long period and thus has fairly low surface energy. The etched surface with increased surface area is significantly more active and hence possesses higher energy. The hydride I surface is similar to native silica, passivated, while hydride II is activated like the etched surface. The alkylated surface has low surface energy similar to most hydrocarbon materials. The increased surface area that is covered by a substantial layer of organic material in the etched chemically modified capillaries can be correlated to the electrokinetic chromatographic behavior of these columns. More specific examples are given in the sections below but some general observations can be made with respect to the AFM data. The elution time for all compounds on the etched chemically modified capillary is significantly longer than on a chemically modified capillary that has not been etched indicating the presence of solute/bonded phase interactions. The peak width of solutes on the etched chemically modified capillary is greater than that for the same solutes that migrate through a native capillary indicating the presence of mass transfer effects. The elution times of solutes are longer as the diameter of the capillary diminishes because of larger k' values that result from the shorter distance a solute must diffuse in order to interact with the bonded moiety on the etched surface [39].

7.4.3 Diffuse Reflectance Infrared Fourier Transform (DRIFT) Spectroscopy Another limitation to the spectroscopic characterization of the inner wall of the etched fused silica capillary is the relatively small surface area. While the etching process is designed to increase the surface significantly with respect to the native material, the area is still small (<1 m2/g) in comparison to common chromatographic supports like porous silica that typically are in the range of 100-500 m2/g. Therefore,

Referencespp. 268-270

254

Chapter 7

insensitive techniques like solid state nuclear magnetic resonance (NMR) that in essence measure the bulk properties of the material are not suitable for characterizing the etched inner wall of the OTCEC capillaries because the surface is very small in comparison to the volume of the sample. However, with proper sample preparation as described above so that the inner bore is readily accessible, surface sensitive techniques offer a means of providing information about the surface that is relevant to the electrophoretic and chromatographic processes taking place inside the capillary. One such technique is DRIFT that has been used extensively to characterize the surfaces of chromatographic stationary phases based on silica and other oxide supports [33,34]. In the DRIFT method, infrared radiation is reflected directly from a surface and contains the spectral information about the moieties (bulk material as well as any molecules chemically bonded or physically adsorbed) that are in the top 1-5 nm thick layer. Sensitivity is enhanced through signal averaging so that many spectra can be rapidly co-added with the S/N ratio improving as the square root of the number of scans. For surface analysis, S/N can also be improved by proper choice of detector. For maximum sensitivity, the mercury-cadmium-telluride (MCT) detector is essential for the relatively low surface area materials such as the OTCEC capillaries. An example of the information obtained by DRIFT for the etched chemically modified inner surface is shown in Fig. 7.7 for a cyclodextrin OTCEC capillary [31]. Two features are important for the characterization of the modified surface. First is the peak near 2270 cm -1 that can be assigned to the Si-H stretching frequency. This band indicates successful silanization of the surface in the first step of the chemical modification process of the etched inner wall. This peak does not disappear during the second step, hydrosilation, because not all Si-H groups react to produce a site with an organic moiety attached, i.e. the reaction is not 100% complete. This is essentially the same result that occurs on porous silica when a hydride surface is modified through hydrosilation [21-23]. The second important feature is the bands in the 2800-3000 -1 cm that can be assigned to carbon-hydrogen stretching frequencies. These infrared absorptions indicate that the second step, hydrosilation, was also successful since these bands can only be present when the organic moiety has been attached to the surface. Neither of these features is observed on an etched capillary that has not been modified by the silanization/hydrosilation process. A similar result (observation of Si-H and C-H stretching bands) was obtained when an etched capillary was modified with a liquid crystal moiety as the bonded phase by the same process [32]. While the overall acquisition of the spectrum for the OTCEC capillaries is not as simple for porous silica materials, the information obtained is comparable and represents a valuable means of characterizing these separation materials.

25 5

Open Tubular CEC

e-

.:E_ E

e" L.

I-4

3~o

2~o

2~o Wavenumbers

24~

2~o

c m -1

Fig. 7.7. DRIFT spectrum of the inner wall of an etched cyclodextrin modified 50 m capillary.

7.4.4 Electron Spectroscopy for Chemical Analysis (ESCA) Photoelectron spectroscopy (ESCA) provides another means of characterizing the inner wall of the fused silica capillary if it is properly opened so that there is easy access to the inner bore. A similar sample preparation procedure to that of the DRIFT experiment is used for ESCA. In the case of ESCA, a beam of x-rays must strike the surface to be analyzed so that the electrons subsequently emitted can be analyzed according to the kinetic energy they possess. The kinetic energy(ies) of the emitted electrons is characteristic of the element(s) present on the surface. Therefore, the ESCA spectrum could be used for verifying the presence of the bonded moiety on the surface as well as the components of the fused silica matrix underneath the organic layer. Indeed, ESCA spectra of etched chemically modified capillaries reveal a carbon (2s) peak in all cases that is indicative of the success of the hydrosilation reaction. Therefore, these results complement the data obtained from DRIFT spectra of the same samples. However, more interesting results are obtained from the ESCA spectra of capillaries that have been etched but not modified. As expected, the spectra contain peaks for silicon and oxygen that are part of the fused silica matrix [32]. In addition to these two peaks, there are also present peaks for the elements F and N. The presence of these elements indicates that some part of the etching reagent (ammonium hydrogen fluoride) has been trapped in the surface matrix during the etching process. An

References pp. 268-270

256

Chapter 7

additional ESCA experiment where the etched surface is bombarded with high energy ions to successively remove the outermost layer of material down to a depth o f - 7 0 nm reveals no substantial change in the elemental composition over this range. Such a result is not surprising since SEM photos reveal surface features are large as 3-5 lam in length as a consequence of the etching process [18,40]. The ESCA data provides unique information about the OTCEC materials and etching process that cannot be obtained from any other method. 7.4.5 Electrooosmotic Flow (EOF) measurements The EOF of the OTCEC capillary provides an indirect means for characterizing the surface of the materials with respect to the sign and magnitude of charge on the inner wall. For example, successful modification of the surface with an organic moiety greatly reduces the number of silanol groups and hence the magnitude of the EOF. In some cases, chemical modification can introduce groups containing a positive charge, such as sulfonic acid, that reverses the direction of the EOF. A typical plot of EOF as a function of the electroosmotic mobility of a neutral marker such as dimethyl sulfoxide (DMSO) for an etched chemically modified OTCEC capillary is shown in Fig. 7.8. First, the overall EOF is quite low so that injection of a neutral marker often requires an extraordinary long period of time to elute from the column. In order to make these EOF measurements, a two marker method is used [41] where after the first injection the electric field is applied for a finite period followed by injection of a second marker and then elution of the two peaks by pressure or vacuum. Knowing the time between the two peaks as well as the period for which the field was applied and the effective length of the capillary is all that is required in order to calculate the EOF. The general shape is always the same for hydrophobic [ 19], hydrophilic [42], chiral [31 ] and liquid crystal [32] bonded phases. At low pH the EOF is anodic instead of cathodic. Then in the pH range of 3 to 4.5 the EOF changes from anodic to cathodic and remains in this direction to the highest pH measured (-8.2). The variation in pH where the EOF changes from anodic to cathodic may be a function of the etching conditions as well as the nature of the organic group bonded to the surface. The most interesting aspect of this determination of EOF as a function of pH is the anodic EOF that occurs at low pH. From the ESCA studies described above that show nitrogen as part of the surface matrix, it is probable that the ammonium moiety from the etching reagent is responsible for the anodic EOF at low pH. Under these conditions, the nitrogen would be protonated giving a positive charge to the surface and hence the anodic EOF. As the pH is raised, the nitrogens and silanols still present lose protons so that the surface changes from a net positive to a net negative charge resulting in the change to a cathodic EOF. The fluorine detected in the ESCA measurements could also be responsible for some negative charge in the surface matrix. Therefore, controlling the pH of

257

Open Tubular CEC

2.00E-08 1.50E-08 1.00E-08 "i, or] "T >

5.00E-09

E

O.OOE+O0

I,i.,. 0 -5.00E-09

pH

I

Y

5

6

I.,IJ

-1.00E-08 -1.50E-08 -2.00E-08

Fig. 7.8. Plot of electroosmotic mobility of an etched chemically modified capillary as a function of pH. Column: etched C 18, i.d. = 20 ~tm. Solute = DMSO.

the running buffer when using the etched OTCEC capillaries could be used to alter either resolution and/or the speed of analysis. 7.5 APPLICATIONS OF O T C E C The OTCEC capillaries described in this chapter have been fabricated in a manner so that the major problems associated with packed capillaries are not present. The open tubular approach greatly reduces the likelihood of bubble formation so that pressurization of the system is not necessary. The other major problem, strong adsorption of basic compounds on the typical support material, is eliminated through the modification scheme, silanization/hydrosilation, that removes silanols and replaces them with hydride groups. This type of separation medium also eliminates the need for any additives in the mobile phase to suppress adsorption of basic compounds, a technique that is often used in packed capillaries as the only means to elute such analytes. Therefore, the bulk of the applications developed to date have centered on the elution characteristics of compounds and separation of mixtures that are difficult to obtain in the packed capillary format. The major exception is the resolution of optical isomers that often can be done equally as well or often better with packed capillaries. The main objective of the chiral separations is to illustrate the presence of Referencespp. 268-270

258

Chapter 7

chromatographic interactions in the open tubular format. The applications developed to date are divided into three major categories and several examples of each type of practical use are provided. In addition, results relating to the stability and reproducibility of these separation materials are also discussed. 7.5.1 Pharmaceuticals and other small molecules

While the primary advantages of CEC (high selectivity with high efficiency) were obvious early in the development of the technique, some significant limitations with respect to the analysis of basic compounds were also identified [43-48]. For the analysis of small molecules, basic compounds such as drugs and metabolites are of considerable interest. Therefore, many OTCEC investigations have focused on these compounds as a possible alternative to packed capillary CEC. Tetracyclines have long been recognized as an important class of antibiotics for both humans and animals [49,50] and HPLC has proved to an excellent separation technique for many mixtures of the parent antibiotic as well as the chemical and physical degradation products. Both standard silica-based [51-57] and polymeric [58] materials have been used as the separation medium. Using OTCEC with etched chemically modified capillaries it was possible to obtain good symmetical peaks (asymmetry factor, AS < 1.5) with reasonable efficiency (50,000-100,000 plates/m) for a variety of tetracyline mixtures [18,29]. In addition it was possible to separate by OTCEC with an etched octadecyl column mixtures of some tetracylines and their degradation products typical of practical samples that were not resolved by HPLC. A physiologically important pair of compounds also anlayzed by HPLC [59], tryptamine and serotonin, were rapidly determined by OTCEC using both C18 and diol modified etched capillaries [20,60]. The effect of column diameter was also documented using this analyte pair. It was found that under identical conditions and using the same stationary phase (C 18) resolution was significantly better on a column with a 20 btm i.d.than one with a 50 btm i.d.[19]. This result also illustrates the chromatographic effect that is present in the etched chemically modified columns since no significant change in resolution would be expected upon a decrease in capillary diameter if the compounds were separated only by electrophoretic differences. In another report, the use of an etched diol modified capillary was shown to be applicable to the analysis of aspartame, the artificial sweetener found in many food product and soft drinks [61]. The stationary phase effect has also been illustrated for another physiologically important pair of analytes, caffeine and theophylline, using two different liquid crystals bonded to the etched wall of the capillary [32]. Both longer retention and better resolution were obtained on a column utilizing a cyanopentoxy (nematic type) liquid crystal bonded material than on a column with a cholesterol moiety (cholesteric type) attached to the etched surface.

Open Tubular CEC

259

Serotonin synthesis and metabolism 5-hydroxyindoleacetic acid

--~-~-~

(5-HIAA)

Tryptophan L-tryptophan HO.

.~o I II

~.

5-hydroxytryptophan

? c

H 5-Hydroxytr~ophlm

E E

(~1

~

..o,on,n J

~

c ~ c ~ ~

H

Serotonin

O 0

B

A i

i

i

i

,,, I

2

3

4

5

20

L. I

I

I

22

24

26

H 5-HydroxytndotelceUc

acid(5-HIAA)

Fig. 7.9. Electrochromatographic separation of a mixture serotonin and its metabolites on an etched liquid crystal modified capillary.

Liquid crystals have proved to be an effective separation medium in this format for a number of other types of small molecules. For example, a pyrimidine/purine basenucleoside mixture was effectively separated by both the nematic and cholesteric type stationary phases with high efficiency [32]. As a further illustration of the chromatographic effect, the elution order changed from thymine < cytosine < adenine < guanine < adenosine on the cholesterol column to thymine < guanine < cytosine < adenine < adenosine on the cyanopentoxy column. A particularly interesting separation is shown in Fig. 7.9 for a mixture of the metabolic components of serotonin on the cyanopentoxy column [32]. This column performs better than one with a cholesterol bonded phase where the last two components are not well-resolved. In addition, the nematic liquid crystal column displays excellent peak symmetry (As = 1 for all peaks) and good efficiency (N = 75,000-100,000 plates/m). In contrast, it has been shown that the cholesterol bonded phase is more suited to the separation of various benzodiazepine mixtures than etched capillaries modified with the cyanopentoxy moiety [32,39,62]. Better separation, although not necessarily better efficiency, was demonstrated for a benzodiazepine mixture on an etched cholesterol modified capillary in comparison to either a bare or unetched cholesterol modified capillary [39]. The presence of solute/bonded phase interactions was also demonstrated with the same benzodiazepine mixture. Retention factors (k) increased significantly on an etched cholesterol modified column with an i.d. of 50 gm in comparison to one Referencespp. 268-270

260

Chapter 7

having an i.d. of 75 ~tm. It was also demonstrated that resolution changed as a function of the etching conditions. Since the only aspect of the capillary that changes as a function of the etching conditions (time and temperature) is the surface area of the inner wall, then this variable will control the amount of stationary phase in the column. If separation was only dependent on electrophoretic effects, then changing the amount of the cholesterol moiety bonded to the wall per unit length of column should have no effect on either retention or resolution. Another comparison showed that using a capillary packed with cholesterol modified particles resulted in different selectivity, resolution and efficiency than an etched open tubular column modified with cholesterol [61]. These results indicate that the two methods, packed capillary and open tubular CEC, have different proportions of electrophoretic mobility and chromatographic effects contributing to the net migration of the same solute in the column under identical mobile phase compositions.

7.5.2 Peptides and proteins Similar to small basic molecules, proteins and peptides present a challenge for obtaining good analytical data in the packed capillary CEC format. The open tubular configuration on the other hand seems to be quite amenable to the elution and analysis of large biomolecules. The first report on the application of etched octadecyl modified capillaries for CEC describes the separation of a five component mixture of basic peptides and proteins with all peaks having good efficiency and symmetry [18]. In addition, a detailed peak shape and migration time comparison for the peptide bradykinin was made on bare, etched, etched/hydride and etched/chemically modified capillaries in order to verify the presence of solute bonded phase interactions. A follow-up investigation described some additional protein separations as well as a comparison of column performance for an angiotensin mixture on bare, etched diol and etched octadecyl capillaries [30]. It was shown that the diol and octadecyl capillaries had distinctly different separation capabilities under identical experimental conditions indicating again the presence of solute/bonded phase interactions. For the separation of basic proteins, the usefulness of the diol capillary was demonstrated for a mixture of four components that strongly adsorb on bare silica surfaces [30]. Fig. 7.10 illustrates a typical separation of this mixture with the peak shapes being symmetrical indicating little or no adsorption of the solutes on the etched chemically modified surface. Efficiencies for the proteins are in the range of 50,000-100,000 plates/m in the OTCEC format that is intermediate between values obtained for HPLC and CE. The improved resolution that can be obtained when reducing the capillary diameter was demonstrated for a mixture of cytochrome c's on a 20 ~m i.d. etched octadecyl modified capillary [ 19]. Mixtures of other basic proteins were also successful in the 20 ~tm i.d. format. An example of a separation with the smaller diameter

Open Tubular CEC

261

A

E

e,1r

(9

r

e',.Q I,,,. 0 U~

<1:

_

-

~

,

~t'

2.

I''' O0

'

,

T

4.

I

O0

.

.

Time

.

.

6.

to

0

8 .!00

(rain.)

Fig. 7.10. Electrochromatogram of protein separation on an etched diol-modified capillary. L (total capillary length) = 45 cm, 1 (effective capillary length) = 25 cm, i.d. = 50 ktm, V = 22 kV, i = 7 ~tm, pH = 4.41. Solutes" 1 = cytochrome c; 2 = lysozyme (turkey); 3 = myoglobin; and 4 = ribonuclease A.

etched capillary is shown in Fig. 7.11. The main problem with using smaller diameter capillaries is detectability for samples with limited mass or volume of material especially with typical UV/visible absorbance detectors. Higher sensitivity detectors such as laser induced fluorescence and mass spectroscopy would be more conducive to using the smaller diameter capillaries. Another possible means for utilizing the etched capillary involves coating a stationary phase on the surface instead of chemically bonding an organic moiety as described above. Such an approach was investigated using Polybrene (hexadimethrin bromide also known as Kieland's reagent) as the coating agent and testing the properties of the new surface for the separation of a protein mixture [63]. The etched Polybrene capillary gave better resolution and higher retention for a protein mixture than a similarly coated unetched capillary. The higher resolution and peak capacity of the etched column could be used when studying the purity of a single protein so that impurity peaks can be more easily identified. The etched liquid crystal modified capillaries were also tested for the separation of protein mixtures [32]. For the two liquid crystal moieties investigated, similar separations of the protein mixtures were obtained. This result is not surprising since previous studies in HPLC have shown that liquid crystals are effective in discriminating between small molecules based on molecular shape [64-66]. The close association of the

References pp. 268-270

Chapter 7

262

E=

,r r

-E 0

2

4

6 8 Time (min.)

10

12

Fig. 7.11. Separation of a cytochrome c mixture on an etched C18 modified 20 pm capillary. Solutes: 1 = horse; 2 = bovine; 3 = chicken; 4 = tuna.

bonded liquid crystal moieties probably helps in the resolution of small molecules but does not assist much in the separation of large molecules such as proteins. The results and conclusions described above have also been confirmed for the separation of proteins in open tubular columns having poly(aspartic acid) acid immobilized on the walls [67]. Due to the nature of the bonded moiety, the mechanism of separation was based on cation-exchange. The elution order observed was similar to that obtained in HPLC so that the pH with respect to the PI of the protein and the ionic strength of the mobile phase control the retention and selectivity. However, column efficiency was found to be 10-100 times higher in the OTCEC format in comparison to HPLC. In the ion-exchange mode with the proteins selected it was determined that optimum separations were achieved at intermediate ionic strength and at high pH. This study also identified one of the main advantages of the low-phase ratio high efficiency OTCEC columns as the ability to do the same or better separations under isocratic conditions that require gradient elution in HPLC.

263

Open Tubular CEC

7.5.3 Chiral compounds

The use of chiral selectors attached to the inner wall of a fused capillary was the first practical demonstration that OTCEC could have some advantages in real applications [10-15]. Most of the separations reported were based on the use of cyclodextrin as the chiral selector for the stationary phase. This choice of chiral selector was the most practical since cyclodextrins and substituted cyclodextrins in the ct, [3 and Y forms have provided the broadest range of enantiomeric separations in HPLC [68]. Therefore, the stationary phase selected for the first etched chemically modified chiral columns were based on 13-cyclodextrin [31] and the enantiomers selected for testing were those that had been successfully resolved by similar bonded moieties in HPLC [69]. Among the compounds that were best resolved in the etched cyclodextrin capillary using the OTCEC format were benzodiazepines. The four compounds that were successfully resolved included oxazepam (separation factor, ot = 1.05), temazepam (ct ~ 1.01), diazepam (or = 1.11) and chlorodiazepam (or = 1.04). The separation for chlorodiazepam is shown in Fig. 7.12 along with the experimental conditions that were found to give the highest resolutions. The three peaks seen in the electrochromatogram are due to the two optical isomers (second and third) and an impuritiy (first). This result was verified by running the same sample on an achiral (octadecyl) column

2

C L_

<

I

I

I

3

4

5

Time (rain.)

Fig. 7.12. Electrochromatographic separation of the optical isomers of D,L-chlorodiazepoxide on an etched cyclodextrin modified capillary, pH = 3.0, V = 20 kV, L = 45 cm, 1 - 25 cm. Peaks: 1 = impurity; 2,3 - D,L isomers of chlorodiazepoxide (individual isomers not identified). References pp. 268-270

Chapter 7

264

where only two peaks were observed. When the other three benzodiazepines were run on the C18 capillary, only a single peak appeared in the electrochromatogram. This data complements the results obtained on the unetched chiral columns [ 10-15] indicating that OTCEC can be a viable alternative in the resolution of enantiomers. As a further test of the etched open tubular approach for the analysis of optical isomers, another column was fabricated based on the selector naphthylethylamine that had been attached to porous silica by the silanization~ydrosilation method for use in HPLC [70]. As in the HPLC experiments, this column was best suited for the resolution of the optical isomers of dinitrobenzoyl methyl esters of amino acids. The best separation (or = 1.14) was obtained for the alanine derivative. In addition, the peak symmetry and efficiency for the naphthylethylamine column was significantly better than that obtained on the cyclodextrin column. However, as shown in HPLC experiments, changes in the amino acid moiety (replacing alanine with valine, etc.) often results in a loss of chiral resolution. In the case of optical isomers, the separation mechanism in HPLC and CEC modes is identical since only interaction between the solute and the bonded phase can result in resolution of the enantiomers. Although it has been demonstrated that successful chiral separations can be achieved in the open tubular format with either etched or unetched capillaries by OTCEC, the high surface area configuration has some potential advantages. The main reasons center on the greater amount of stationary phase that is bonded per unit length of the etched capillary. This higher bonded phase ratio per unit length means that a larger amount of solute (loadability) is possible with the etched column and hence greater sensitivity in detection. The second advantage is that the solute comes into contact with more stationary phase as it moves through the column and hence better resolution is achieved over the same length when compared to the unetched capillary.

7.5.4 Stability and reproducibility For the evaluation of any separation medium, three types of characterization are important in determining the overall usefulness of the material: within column reproducibility, column-to-column reproducibility and stability. All of these factors have been tested for the etched chemically modified capillaries in order to determine the reproducibility of the etching and chemical modification processes as well as the ruggedness of the silanization/hydrosilation method for the attachment of various organic moieties to the roughened inner wall. Several types of etched capillaries have been tested with respect to within column reproducibility. For example, the reproducibility of the migration times for 151 consecutive injections of the proteins lysozyme and ribonuclease A was tested on an etched C18 modified column at pH = 3.0 [20]. It was found that both solutes gave no discernible increase or decrease in migration time (tM) and its overall reproducibility

Open Tubular CEC

6

265

Etched C18 modified capillary, i.d. = 20 l~m

-

,..-7. r .m

g5 .E_ I-r

.9

4

r L ,m

3 2~

I

I

I

I

I

I

286

291

296

301

306

311

Injection #

Fig. 7.13. Reproducibility of 30 consecutive injections of tryptamine at pH - 2.14 on an etched C18 modified capillary after 280 injections.

was + 1.5%. This test was conducted after more than 200 injections of other samples as well as after being mounted and demounted several times that included a methanol wash. A similar test was run on a 20 ~tm i.d. etched C18 capillary using tryptamine as the test solute [19]. After the column had already been subject to more than 300 injections, the variation in the migration time of tryptamine for 30 consecutive runs was + 1.7% (Fig. 7.13). The two liquid crystal columns were also tested after considerable use for reproducibility of migration times for small basic solutes [32]. For the cholesterol capillary after more than 275 runs and for the cyanopentoxy capillary after more than 150 runs, 30 consecutive injections gave a relative standard deviation (RSD) < 2% for the test solute migration time. The column-to-column reproducibility was evaluated by taking columns that were made from different batches of etching and modification as opposed to taking several sections from a single piece of capillary fabricated at one time. This approach should be a more rigorous test of column-to-column reproducibility since it is subjected to more potential errors than cutting several columns from a single length of capillary. Two tests each involving a solute pair were run on etched C18 modified capillaries from four different batches of material. The first test involved the solutes tryptamine and serotonin. The four batches gave a plate count of between 95,000 and 130,000 plates/m for serotonin and the separation factor for the two compounds ranged between 1.057 and 1.070 with an RSD = 0.52%. The second test involved the two proteins turkey and chicken lysozyme. In this evaluation the separation factor between the two solutes was measured and it ranged from 1.07 to 1.10. Therefore, for both large and small basic molecules, column-to-column reproducibility utilizing a stringent set of criteria was excellent.

References pp. 268-270

266

Chapter 7

The column stability can be inferred from some of the same data that was used to evaluate reproducibility. The most direct evaluation is the 151 consecutive injections of the two proteins on the etched C18 capillary [20]. If some degradation of the bonded material had occurred, then an identifiable trend in the migration time (either increase or decrease) should have been evident for one or both of the proteins. The other tests on the 20 gm i.d. etched C18 [19] and liquid crystal [32] capillaries involved fewer injections of the test solute but after many total injections. Again, any serious degradation of the bonded stationary phase would have resulted in either a discernible trend or a much larger % RSD in the migration times. These results concur with the stability data obtained for stationary phases used in HPLC [22,23] and for wall modifications used in HPCE [42,71] produced by the silanization/hydrosilation process. In one of the CE tests, more than 300 consecutive injections of a protein mixture were done at pH = 8.5 [71]. Therefore, as expected, the Si-C linkage produced by the modification method used for the etched chemically modified capillaries has high stability resulting in a long column lifetime. 7.6 CONCLUSIONS The open tubular approach to capillary electrochromatography provides another valuable tool that complements the broad range of micro-separation techniques already available. In contrast to packed capillary electrochromatography, OTCEC at present seems best suited for the analysis of charged compounds so that separation occurs via a mixed mode mechanism: a combination of electrophoretic migration effects and solute/bonded phase interactions. Electrochromatographic capillary columns packed with octadecylsilica (ODS) have a greater bonded phase ratio and therefore higher loadability in terms of the amount of solute that can be injected into the column. This higher loadability results in better sensitivity especially with conventional optical detectors. However, a serious problem encountered in the packed capillary format is the presence of residual silanols on the support surface, many of these accessible to sample molecules as they are driven through the column. The silanols are necessary to generate the electroosmotic flow needed to drive neutral molecules through the system. Many of the most challenging separation problems exist for the analysis of biomolecules and pharmaceuticals. These solutes have a high affinity for the silanols present on most conventional packing materials used for CEC. Additives that suppress the adsorption of basic compounds on the conventional CEC phases can be included in the running buffer but these additional species in the mobile phase often complicate and compromise the analysis. Since many molecules of interest in biological and pharmaceutical analysis are charged under some reasonable pH condition, then the use of OTCEC can provide a means for solving some of the

Open Tubular CEC

267

problems that exist for packed capillary CEC. In addition, the etching process used to increase the surface area and hence the capacity of the system also has some beneficial effects. It creates a small anodic electroosmotic flow at low pH and a small cathodic EOF under neutral to basic conditions. Since some of the cathodic EOF may be due to the presence of fluoride ions in the surface matrix and relatively few silanols, then adsorption of basic compounds is minimized. Further deactivation of the surface occurs through the chemical modification process, silanization/hydrosilation. In this step the remaining accessible silanols are converted to hydrides and then the appropriate organic moiety is attached to the surface via a stable Si-C linkage. The increased surface area created by the etching process is used to improve the capacity of the column as well as to provide greater solute/bonded phase interactions to improve resolution. The open tubular format for CEC utilizing etched chemically modified capillaries is still in its infancy, i.e. less than five years in development [18]. All of the physical and chemical processes involved in the etching, chemical modification and subsequent electrochromatographic behavior of these materials is still not understood and is under investigation. While decreasing the inner diameter of the capillary may increase the amount of solute/bonded phase interactions, detection limits for conventional detectors can be severely strained. Therefore, improved instrumentation is a key to further progress in the development of OTCEC. However, it has already been demonstrated that useful applications for the analysis of biomolecules and pharmaceuticals are possible by OTCEC. As these separation materials become more available and are further developed, new applications will be found. Perhaps the right combination of the etching conditions, the inner diameter of the capillary and the bonded organic moiety may lead to useful separations for some neutral analytes. However, at present there are many challenging separations involving predominantly charged species where OTCEC can be exploited in its present state of development. Such analyses may not be possible by CE since electrophoretic differences alone may not be sufficient to separate some analytes. Similarly, packed capillary CEC may not be suitable because adsorption effects are too great to give reproducible analyses. As is the case with many methods, OTCEC may also supplant existing analytical schemes because it offers a faster or more reproducible alternative to an established protocol. 7.7 A B B R E V I A T I O N S

AS AFM BET C18 CE

separation factor asymmetry factor atomic force microscopy Brunauer-Emmet-Teller octadecyl capillary electrophoresis

Referencespp. 268-270

Chapter 7

268 CEC DMSO DRIFT EOF ESCA GC HPLC k' MCT laep ~t-HPLC N NMR ODS OTCEC RMS RSD SEM TM THF TES

capillary electrochromatography dimethyl sulfoxide diffuse reflectance infrared Fourier Transform electroosmotic flow electron spectroscopy for chemical analysis gas chromatography high-performance liquid chromatography capacity factor mercury-cadmium-telluride electrophoretic mobility micro-high performance liquid chromatography number of theoretical plates nuclear magnetic resonance octadecylsilica open tubular capillary electrochromatography root mean square relative standard deviation scanning electron microscopy migration time tetrahydrofuran triethoxysilane

7.8 REFERENCES

1 J.R. Mazzeo, I.S. Krull in Handbook of Capillary Electrophoresis, J.P. Landers, ed., CRC Press, Boca Raton, 1993, Chapt. 18. 2 S.A. Swedberg in Handbook of Capillary Electrophoresis, J.P. Landers, ed., CRC Press, Boca Raton, 1993, Chapt. 19. 3 Z. Zhao, A. Malik, M.L. Lee, Anal. Chem., 65 (1993) 2747. 4 S.A. Swedberg, Anal. Biochem., 185 (1990) 51. 5 J.S. Green, J.W. Jorgenson, J. Chromatogr., 478 (1989) 63. 6 D. Mcganigill, S.A. Swedberg in Techniques in Protein Chemistry, T. Hugli, ed., Academic Press, San Diego, 1989, Chapt. 45. 7 T. Wang, R.A. Hartwick, J. Chromatogr., 594 (1992) 325. 8 S.A. Swedberg, M. Ditmann, Hewlett-Packard J., 64 (1995) 57. 9 Y. Tsuda, K. Nomura, G. Nakagawa, J. Chromatogr., 248 (1982) 241. 10 S. Mayer, V. Schurig, J. High Res. Chromatogr., 15 (1992) 129. 11 D. Armstrong, Y. Tang, T. Ward, M. Nichols, Anal. Chem., 65 (1993) 1114. 12 S. Mayer, V. Schurig, J. Liq. Chromatogr., 16 (1993) 915. 13 S. Mayer, V. Schurig, Electrophoresis, 15 (1994). 14 S. Mayer, M. Schleimer, V. Schurig, J. Microcol. Sep., 6 (1994) 43. 15 E. Francotte, M. Jung, Chromatographia, 42 (1996) 521. 16 O. Bruggeman, R. Freitag, M.J. Whitcombe, E. Vulfson, J. Chromatogr. A, 781 (1997) 43. 17 F. Onuska, M.E. Comba, T. Bistridcki, R.J. Wilkinson, J. Chromatogr., 142 (1977) 117. 18 J.J. Pesek, M.T. Matyska, J. Chromatogr. A, 736 (1996) 255. 19 J.J. Pesek, M.T. Matyska, S.-J. Cho, J. Chromatogr. A, 845 (1999) 237. 20 M.T. Matyska, Chem. Anal. (Warsaw), 43 (1998) 637. 21 J.E. Sandoval, J.J. Pesek, Anal. Chem., 63 (1991) 2634.

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22 M.C. Montes, C.-H. Chu, E. Jonsson, M. Auvinen, J.J. Pesek, J.E. Sandoval, Anal. Chem., 65 (1993) 808. 23 J.J. Pesek, M.T. Matyska, J.E. Sandoval, E.J. Williamsen, J. Liq. Chromatogr. & Rel. Technol., 19 (1996) 2843. 24 B. Marciniec, Handbook on Hydrosilylation, Pergamon Press, Oxford, 1992. 25 J.J. Pesek, M.T. Matyska, M. Oliva, M. Evanchic, J. Chromatogr. A, 818 (1998) 145. 26 J.J. Pesek, M.T. Matyska, E.J. Williamsen, M. Evanchic, V. Hazari, K. Konjuh, S. Takhar, R. Tranchina, J. Chromatogr. A, 786 (1997) 219. 27 J.J. Pesek, M.T. Matyska, S. Muley, Chromatographia, in press. 28 J.J. Pesek, M.T. Matyska, E.J. Williamsen, R. Tam, Z. Wang, J. Liq. Chromatogr. & Rel. Technol., 21 (1998) 2747. 29 J.J. Pesek, M.T. Matyska, J. Chromatogr. A, 736 (1996) 313. 30 J.J. Pesek, M.T. Matyska, L. Mauskar, J. Chromatogr. A, 763 (1997) 307. 31 J.J. Pesek, M.T. Matyska, S. Menezes, J. Chromatogr. A, 853 (1999) 151. 32 M.T. Matyska, J. J. Pesek, A. Katrekar, Anal. Chem., 71 (1999) 5508. 33 J.J. Pesek, M.T. Matyska, J. Interface. Sci., 5 (1997) 103. 34 J.J. Pesek, M.T. Matyska, in Adsorption and Its Application in Industry and Environmental Protection, A. Dabrowski, ed., Elsevier, Amsterdam, 1999, Vol. I, pp. 117-142. 35 S. Kaupp, H.J. Watzig, J. Chromatogr. A, 781 (1997) 55. 36 A. Cifuentes, J. Diez-Mesa, J. Fritz, D. Anselmetti, A.E. Bruno, Anal. Chem., 70 (1998) 3458. 37 P.E. Pullen, J.J. Pesek, M.T. Matyska, J. Frommer, Anal. Chem., 72 (2000) 2751. 38 E.W. van der Vegte, G. Hadziioannou, Langmuir, 13 (1997) 4323. 39 A. Cataby, H. Sawada, K. Jinno, J.J. Pesek, M.T. Matyska, J. Cap. Electrophoresis, 5 (1998) 89. 40 J.J. Pesek, M.T. Matyska, J. Capillary Electrophoresis., 4 (1997) 213. 41 J.E. Sandoval, S.-M. Chen, Anal. Chem., 68 (1996) 2771. 42 M.T. Matyska, J.J. Pesek, J.E. Sandoval, U. Parkar, X. Liu, J. Liq. Chromatogr. & Rel. Yechnol., 23 (2000) 97. 43 M.M. Dittmann, K. Wienand, F. Bek, G.P. Rozing, LC/GC, 13 (1995) 800. 44 M.M. Dittmann, G.P. Rozing, J. Microcol. Sep., 9 (1997) 399. 45 J.J. Pesek, M.T. Matyska, Electrophoresis, 18 (1997) 2228. 46 L.A. Colon, Y. Guo, A. Fermier, Anal. Chem., 69 (1997) 461A. 47 M.M. Robson, M.G. Cikalo, P. Myers, M.R. Euerby, K.D. Bartle, J. Microcol. Sep., 9 (1997) 357. 48 M.G. Cikalo, K.D. Bartle, M.M. Robson, P. Myers, M.R. Euerby, The Analyst, 123 (1998) 87R. 49 D. Gotlieb, P.D. Shaw, Antibiotics, Vol. 1, Springer Verlag, New York, 1967. 50 J.R.D. McCormick, in Z. Vanek, Z. Hostaled (Editors), The Biogenesis of Antiobiotic Substances, Academic Press, New York, 1965, p.83. 51 K. Wolfs, E. Roets, J. Hoogmartens, H. Vanderhaeghe, J. Chromatogr., 385 (1986) 444. 52 N.H. Khan, E. Roets, J. Hoogmartens, H. Vanderhaeghe, J. Chromatogr., 405 (1987) 229. 53 W. Naidong, E. Roets, J. Hoogmartens, J. Pharm. Biomed. Anal., 7 (1989) 1691. 54 N.H. Khan, P. Wera, E. Roets, J. Hoogmartens, J. Liq. Chromatogr., 13 (1990) 1351.

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55 W. Naidong, K. Verresen, E. Roets, J. Hoogmartens, J. Chromatogr., 586 (1991) 61. 56 W. Naidong, J. Thuranira, K. Vermeulen, E. Roets, J. Hoogmartens, J. Liq. Chromatogr., 15 (1992) 2529. 57 M.T. Matyska, J.J. Pesek, A.M. Siouffi, Chem. Anal. (Warsaw), 40 (1995) 517. 58 P.D. Bryan, J.T. Stewart, J. Pharm. Biomed. Anal., 12 (1994) 675. 59 H. Wakabayashi, K. Shimada, J. Chromatogr., 381 (1986) 21. 60 J.J. Pesek, M.T. Matyska, J. Liq. Chromatogr. & Rel. Technol., 21 (1998) 2923. 61 J.J. Pesek, M.T. Matyska, J. Chromatogr. A, 781 (1997) 423. 62 K. Jinno, H. Sawada, A.P. Catabay, H. Watanabe, N.B.H. Sabli, J.J. Pesek, M.T. Matyska, J. Chromatogr. A, 887 (2000) 479. 63 J.J. Pesek, M.T. Matyska, S. Swedberg, S. Udivar, Electrophoresis, 20 (1999) 2343. 64 A. Catabay, Y. Saito, C. Okumura, K. Jinno, J.J. Pesek, E.J. Williamsen, J. Microcolumn Sep., 9 (1997) 81. 65 A. Catabay, C. Okumura, K. Jinno, J.J. Pesek, E.J. Williamsen, Chromatographia, 47 (1998) 13. 66 A. Catabay, M. Taniguichi, K. Jinno, J.J. Pesek, E.J. Williamsen, J. Chromatogr. Sci., 36 (1998) 111. 67 W. Xu, F.E. Regnier, J. Chromatogr. A, 853 (1999) 243. 68 S.M. Han, Biomed, Chromatogr., 11 (1997) 259. 69 G. Felix, C. Cachau, A. Thienpont, M.-H. Soluard, Chromatographia, 42 (1996) 583. 70 J.J. Pesek, M.T. Matyska, S. Kamath, Analusis, 25 (1997) 253. 71 M. Chiari, M. Nesi, J.E. Sandoval, J.J. Pesek, J. Chromatogr. A, 717 (1995) 1.

Chapter 8

Hyphenation of Capillary Electrochromatography and Mass Spectrometry: Instrumental Aspects, Separation Systems, and Applications Christian G. H U B E R and Georg H O L Z L

Institute of Analytical Chemistry and Radiochemistry, Leopold-Franzens-University, Innrain 52 a, A-6020 Innsbruck, Austria

CONTENTS

8.1 8.2

8.3

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Instrumentation and technology for coupling o f CEC and MS . . . . . . 8.2.1 Instruments for performing capillary electrochromatographic separations . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.1.1 Isocratic and gradient CEC instruments . . . . . . . . 8.2.1.2 Pressure assisted CEC instruments . . . . . . . . . . . 8.2.2 C E C - - M S interfaces . . . . . . . . . . . . . . . . . . . . . . . . 8.2.2.1 C E C - - C F F A B interface . . . . . . . . . . . . . . . . 8.2.2.2 C E C - - A P I interfaces . . . . . . . . . . . . . . . . . . 8.2.2.2.1 C E C - - E S I - M S interface . . . . . . . . . 8.2.2.2.2 Implications o f ESI on interface configuration . . . . . . . . . . . . 8.2.2.2.3 C E C - - A P C I - M S interface . . . . . . . . 8.2.3 Mass analyzers . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.3.1 Linear quadrupole mass analyzers . . . . . . . . . . . 8.2.3.2 Quadrupole ion trap mass analyzers . . . . . . . . . . 8.2.3.3 Time-of-flight analyzers . . . . . . . . . . . . . . . . 8.2.3.4 Comparison o f mass analyzers used for C E C - - E S I - M S 8.2.4 Column technology for C E C m M S ................ Stationary phase-mobile phase systems used for C E C - - M S .......

272 273 275 275 277 277 277 279 280 285 286 286 288 289 291 293 293 296

Chapter 8

272

8.4 8.5

8.6 8.7

8.3.1 Stationary phases . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3.2 CEC eluents compatible with mass spectrometry . . . . . . . . . Optimization of electrochromatographic and mass spectrometric conditions Examples of application . . . . . . . . . . . . . . . . . . . . . . . . . . 8.5.1 Smallmolecules . . . . . . . . . . . . . . . . . . . . . . . . . . 8.5.2 Polypeptides and enzymatic digests . . . . . . . . . . . . . . . . 8.5.3 Nucleic acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

296 297 298 305 305 309 311 312 313

8.1 INTRODUCTION

Capillary electrochromatography (CEC) can be viewed as a hybrid of high-performance liquid chromatography (HPLC) and capillary electrophoresis (CE) [1,2], combining the high separation efficiency of CE with the outstanding separation selectivity of HPLC [3,4]. The major driving force for the development of CEC has been the excellent separation efficiency and high peak capacity offered by electrochromatographic separation systems that can be used to separate the multitude of compounds present in real-life sample mixtures in a short period of time. Due to the short path length of the in-capillary detection cell, CEC using conventional on-column UV detection has the same detection limitations as capillary electrophoresis (CE), resulting in considerably low mass detection limits but relatively poor concentration detection limits [5]. Mass spectrometry (MS) has evolved into a very powerful and sensitive detection technique for liquid phase separation methods since the advent of ionization methods that generate gas phase ions from analytes dissolved in a liquid phase, namely discharge-induced atmospheric pressure chemical ionization (APCI) [6], continuous flow fast atom bombardment (CFFAB) [7], and electrospray ionization (ESI) [8]. Particularly ESI-MS has found broad application in all fields of analytical science, mainly because of its ability to generate intact gas-phase ions from very polar and/or high molecular mass analytes and because of the multiple charging, which allows the mass spectrometric investigation of large biomolecules of molecular masses far beyond 100000 with conventional mass analyzers of only limited mass range (< m/z 4000) [9]. The low flow rates of analyte solution in the microliter or submicroliter per minute range that are required for optimum performance of CFFAB or ESI are most closely matched by miniaturized liquid phase separation methods, such as CE, capillary HPLC, or CEC [10,11]. Consequently, CEC coupled to MS offers a highly efficient, miniaturized separation system coupled to a sensitive detection technique, that allows the positive identification of the separated compounds and additionally yields structurally related information upon fragmentation [ 11-14]. In this chapter, we will focus on the instrumental and technological requirements

Hyphenation

273

that have to be met for the successful hyphenation of CEC and MS. A discussion of the types of CEC instruments, interface designs, and mass analyzers for CEC--MS is followed by some aspects of capillary electrochromatographic phase systems which are compatible with mass spectrometric detection. The chapter is concluded by the presentation of selected examples of application. 8.2 INSTRUMENTATION AND TECHNOLOGY FOR COUPLING OF CEC AND MS Since the first demonstrations of high efficiency separations by CEC [3,4,15,16], the technique has significantly advanced. As new applications of CEC continue to appear, the advantages of coupling CEC instruments with mass spectrometers and the importance of the development of dedicated instrumentation becomes obvious. A schematic diagram of a CEC--MS instrumental arrangement is shown in Fig. 8.1. The two major parts are a capillary electrochromatograph and a mass spectrometer, which are joined together by a suitable interface. The lack of dedicated and automated CEC instrumentation has been one of the limiting factors in the development of CEC, although most commercially available CE instruments can be utilized for CEC after slight alteration. Capillary columns manufactured for CEC with UV detection usually have a detection window right after the packed segment. The retaining frit between the column packing and the ensuing open

capillary electrochromatograph

interface

inlet electrolyte reservoir

electrode

CEC c o l u m n

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e 9

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Fig. 8.1. Generic illustration of a CEC--MS instrumental arrangement.

References pp. 313-316

m/z

274

Chapter 8

capillary segment is believed to be a source of nucleation for the formation of air bubbles, which destabilize and finally discontinue the flow of current through the capillary and thus have an untoward effect on the separation process [17,18]. The application of pressure both to the inlet- and outlet electrolyte reservoirs has therefore been suggested to reduce the risk of formation of air bubbles in the packed and unpacked regions of the separation capillary [16,17]. Accordingly, some of the commercial CE instruments have been modified by the manufacturer to allow application of inert gas pressures up 2 MPa to the electrolyte reservoirs. However, pressurization at the column outlet precludes the interfacing of CEC to MS and therefore, alternative arrangements have to be used for performing CEC--MS. Another problem in the interfacing of CEC and MS is the necessity to conduct the column effluent into the ion source of the mass spectrometer, which frequently entails relatively long distances between the column inlet and the probe tip of the ion source, especially with commercial CEC systems. In order to maintain sufficient electrosmotic flow through such long capillary columns and to keep analysis times in an acceptable range, relatively high voltages have to be applied to the separation columns, eventually exceeding the 30 kV voltage limit of most commercial instruments. The majority of these problems has been circumvented by designing and building laboratory-made CEC units [ 19,20]. The on-line hyphenation of CEC and MS is nontrivial and more complicated than HPLC--MS in that an electrical contact must be established at the column outlet in order to provide a closed circuit for the CEC current flow. Moreover, the interface should also provide an outlet electrolyte reservoir for CEC in order to prevent electrolyte depletion due to electrolysis and electromigration [21]. Since the column dimensions and characteristic flow rates of CE and CEC are very much alike, interface technology for coupling of CEC with MS strongly benefits from the interface designs that have been already developed for CE--MS [22,23]. The earliest CEC MS interfaces were based on CFFAB [24,25], while today the ESI interface [26] is the most commonly used. The principal types of mass analyzers encountered in analytical laboratories are double focusing magnetic/electrostatic sectorfield-, quadrupole-, quadrupole ion trap-, time-of-flight-, and ion cyclotron resonance mass analyzers. Because of their comparatively low cost, easy tuning and operation, and moderate lab space requirements, quadrupole, quadrupole ion trap and time-of-flight mass analyzers keep the widest acceptance as mass analyzers for CEC--MS.

275

Hyphenation

8.2.1 Instruments for performing capillary electrochromatographic separations 8.2.1.1 Isocratic and gradient CEC instruments

Isocratic CEC is frequently performed with CE instruments, that have been modified to allow pressurization of the electrolyte reservoirs. This has the advantage that CE instruments which are frequently found in laboratories involved in separation technology can also be applied to CEC separations. Most CEC separations have been carried out so far by isocratic elution at applied voltages up to 30 kV [ 16,19,27-34]. A modular instrument was built that could be operated at voltages up to 90 kV [20]. To realize the full potential of CEC it is essential to employ instrumentation having gradient elution capability that allows to gradually increase in a controlled manner the eluent strength during a chromatographic run. Moreover, gradient elution is indispensable when a mixture to be separated contains several components, whose elution windows do not overlap under a chosen eluent strength. Electrosmotically driven solvent gradients have been obtained by merging two electosmotic flows that are generated in open fused silica tubes and regulated by computer controlled voltages of 0-50 kV [35]. Other investigators have facilitated gradient elution in CEC by incorporation of gradient forming systems generally employed in HPLC, where the eluent composition in the reservoir at the column inlet is changed by means of two reciprocating displacement pumps [36-40]. Thus, the composition of the mobile phase flowing through the column and driven by electrosmotic forces can be changed in a time controlled manner as customary in gradient HPLC [37]. Fig. 8.2 illustrates a gradient forming system that was applied to the gradient separation of PTH-amino acids with subsequent detection by ESI-MS. Fig. 8.3 com-

inlet resq~rvoir capillary column with tapered end

electrode injection syringe I

'

Illl ~-] sample

I

mobile phase gradient

~l

inject~

ionESo~rce

waste

Fig. 8.2. Schematic illustration of a CEC instrument capable of isocratic and gradient elution and ESI-MS detection. References pp. 313-316

Chapter 8

276 Q 48N

e~-

o

tO

0-

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277N,Q

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0

0

I

5

I

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time [mini

I

15

Fig. 8.3. Isocratic (a) and gradient (b) separation of PTH amino acids. Column, 250 x 0.075 mm i.d. packed with 3.5 ~tm/80 fi, Zorbax ODS; eluents, (A) 2 mmol/1 ammonium acetate, pH 7.0, (B) 2 mmol/1 ammonium acetate, pH 7.0, 90% acetonitrile; isocratic elution with 30% B in (a); gadient elution with 30-80% B in 5 min, followed by 80% for 5 min in (b); flow rate of mobile phase through inlet reservoir, 100 ktl/min; applied voltage, 15 kV; Detection, ESI-MS, m/z 100-2000, 0.5 s/spectrum integration time; sheath liquid, 1 mmol/1 ammonium acetate, pH 7.0, 90% methanol, 3 pl/min; injection, electrokinetic, 2 kV, 2 s; sample, PTH-asparagine, PTH-glutamine, PTH-threonine, PTH-glycine, PTH-tyrosine, PTH-alanine (in order of elution). (Reproduced from ref. [82] with permission of Elsevier Sciences B. V.).

pares the separation pattem of six PTH-amino acids under isocratic and gradient elution conditions with a 250 mm long column at an applied voltage of 15 kV. Under isocratic conditions, the separation of all six compounds took 30 min to complete and the peaks were 30-60 s wide with a mobile phase comprising 2 mmol/1 ammonium acetate, pH 7.0, 27% acetonitrile. However, upon employing a 5 min gradient of 27-72% acetonitrile the separation was completed within 7 min and all peaks except PTH-asparagine and PTH-glutamine were resolved. The data illustrate that analysis times in CEC can be reduced significantly by using gradient elution without untoward effects on the analytical result. Moreover, because of the focusing effect of gradient elution, the peak widths are considerably decreased, resulting in improved signal-tonoise ratio and enhanced detectability, as can be deduced from the absolute signal intensities in Fig. 8.3.

Hyphenation

277

8.2.1.2 Pressure assisted CEC instruments

Pressure assisted CEC alleviates some of the problems experienced in solely electrodriven CEC, such as bubble formation, current instability, and drying out of the column ends [41-43]. Both a pressure- and an electrosmotically driven flow are generated through the separation column by means of a split-flow pumping system and a high-voltage power supply. In this configuration, solvent gradients can be readily formed by using a gradient solvent delivery system rather than an isocratic pump for generating pressure driven flow [43]. Since charged analytes are separated in CEC due to differences in their partitioning between the mobile and the stationary phase as well as due to differences in their electrophoretic mobility, the fine tuning of separation selectivity is feasible during the chromatographic separation with an additionally applied electric field, as demonstrated for oligonucleotides [43] and tryptic peptides [44,45]. Moreover, suppression of bubble formation by hydraulic flow component obviates the need for pressurization of the column outlet so that pressure assisted CEC can be conveniently coupled to mass spectrometry [ 12]. 8.2.2 C E C m M S interfaces

The on-line hyphenation of CEC and MS has several potentially challenging instrumental aspects which complicate the successful combination of these two techniques. The first arises due to the absence of a CEC column outlet electrolyte reservoir and the need to achieve electrical continuity for the CEC system, and, in the case of ESI, also for the ESI ion source. Another consideration is the requirement to efficiently remove the mobile phase and simultaneously generate gas phase ions from the analyte, which have to be transferred with high efficiency into the vacuum of the mass analyzer. Because of this situation, numerous designs have been advanced which solve to various degrees these related problems and are discussed individually below. 8. 2.2.1 C E C - - C F F A B interface

In fast-atom bombardment (FAB), first developed by Barber et al. [46], the analyte is dissolved in a liquid, viscous matrix, typically mixtures of water with 5-90% glycerol, 3-nitrobenzylalcohol, thioglycerol, dithiothreitol/dithioerythritol, or triethanolamine. The analyte-matrix mixture is bombarded at the probe tip with a beam of fast atoms or ions, such as 5-8 kV xenon atoms or 10-40 kV cesium ions formed in a gun. The beam hits the analyte-matrix mixture, inducing a shock wave which desorbs ions and molecules from the uppermost layer of the solution. Ionization of the analytes occurs either in the matrix or in the vapor phase above the matrix, usually by proton transfer reactions between analyte and matrix ions. Subsequently, the gas References pp. 313-316

Chapter 8

278

primary atom or ion beam

CEC column

sheath capillary

stainless steel probe shaft

vespel stainless steel insulator probe tip .

. X

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.

to mass analyzer

syringe pump delivering FAB matrix solution

Fig. 8.4. Schematic of a coaxial CFFAB ion source for CECwCFFAB-MS.

phase ions are transferred into the mass analyzer for m/z determination. The purpose of the matrix is to protect the analyte from decomposition by the atom or ion beam and to allow replenishment of analyte at the surface. FAB is very efficient for producing gas phase ions from thermally labile, highly polar, and high molecular mass (up to Mr 10000) compounds. In continuos flow FAB (CFFAB) [7,24,47] the analyte-matrix mixture is delivered continuously to the probe tip through a fused silica capillary which terminates at the probe tip. This configuration provides a means of coupling liquid phase separation techniques with FAB-MS. Addition of the matrix to the analyte solution is accomplished by one of two methods. (1) The matrix is added at concentrations of 5-10% to the mobile phase, and the column effluent is directly fluxed into the CFFAB ion source or (2) column effluent and matrix solution are delivered independently to the probe tip by a coaxial arrangement of two concentric fused silica capillaries. The schematic of a coaxial CFFAB probe which is compatible with nanoscale liquid phase separation techniques delivering microliter and submicroliter per minute flow rates [48,49] is illustrated in Fig. 8.4. In this interface, extra column band broadening is minimized because the mixing of the matrix and the column effluent takes place only at the CFFAB probe tip. Since most CFFAB sources are used in combination with sectorfield mass analyzers, the probe tip is maintained at the MS source voltage, which also serves as the reference potential for the high voltage used for the CEC separation, and thus completes the electrical circuit of CEC. For instruments with high voltage sources (such as Kratos instruments), an insulator is used to isolate the probe tip from the probe shaft, whereas such insulation is not necessary for

Hyphenation

279

arrangements, where the probe tip is at ground potential (as in Finnigan MAT instruments). Since water is being continuously evaporated at the probe tip, some heating is necessary to prevent freezing of the liquid on the surface of the target or in the exit portion of the capillary. Therefore, the source is typically maintained at a temperature of 30-50~ Stable operation of the interface is achieved when the rate of liquid onto the surface is balanced by the rate of evaporation, resulting in a thin liquid film on the target surface. The major advantage of the CFFAB interface is the ability to detect [M+H] + or [M-H]-pseudomolecular ions of a wide variety of polar and charged molecules at high detection sensitivities and low limits of detection. A disadvantage of CFFAB is the relatively high chemical background recorded, especially at low m/z values (< 400). Moreover, in situations where the electrosmotic flow does not counterbalance the electrophoretic migration of the ions in the background electrolyte, the conductivity of the background electrolyte decreases due to the lack of an outlet electrolyte reservoir that replenishes those ions, eventually leading to a breakdown of current through the separation column. 8.2.2.2 CEC--API interfaces

Although the choice of interface for the coupling of CEC and MS is ultimately governed by the instrumentation available, the modern trend now clearly favors atmospheric pressure ionization (API) methods, including electrospray ionization (ESI) and atmospheric pressure chemical ionization (APCI). The API-MS interface is well suited for CEC--MS interfacing, since it produces ions directly from liquid solutions at atmospheric pressure. An API interface consists of six major parts, as depicted in Fig. 8.5: (1) the liquid introduction and spraying device, (2) the atmospheric pressure ion source region, where the ions are generated by means of APCI or ESI, (3) an ion sampling aperture, (4) an atmospheric pressure to vacuum interface, (5) a skimmer for entrance of the ions into high vacuum, and (6) an ion optical system to transport the ions to the mass analyzer [ 11 ]. The column effluent is nebulized into the atmospheric pressure region either pneumatically with heat assistance (in APCI), by means of a strong electric field (in ESI), or by a combination of an electric field and pneumatic nebulization (pneumatically assisted ESI, also referred to as ionspray). Additionally, ultrasonically assisted electrospray ionization has also been described [50]. Ions are produced from the evaporating droplets, either by gas-phase ion-molecule reactions initiated by a corona discharge plasma (in APCI) or by consecutive desolvation of ions already preformed in solution (both in APCI and ESI). The generated and partially solvated ions, together with solvent vapor and spray gas are transferred by an ion sampling device into a first pumping stage. Usually, the transfer is accompanied

References pp. 313-316

Chapter 8

280

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II

4

II ua ru oe

CEC

() () ()

10 s Pa

I

) )

analyzer

';'

10 2 Pa

10 -s - 10 .7 Pa

mechanical pump

turbopump

Fig. 8.5. Schematic of an API interface for CEC--MS. 1, introduction of column effluent from CEC and spraying device; 2, atmospheric pressure region; 3, ion sampling aperture; 4, atmospheric pressure to vacuum interface; 5, skimmer; 6, ion transfer optics (adapted from ref. [11]).

by heat transfer to the ion beam in order to assist desolvation of the ions either by means of a counter flow of heated drying gas (see Fig. 8.6) or by passing the ion beam through a heated metal capillary tube (see Fig. 8.7). The mixture of gas, solvent vapor, drying gas and ions is supersonically expanded into the low-pressure region. The core of the expansion is sampled by a skimmer into a second pumping stage, where an ion focusing and transfer device aids in the transport of the ions to the mass analyzer.

8.2.2.2.1 CEC--ESI-MS interface The three major steps in the production of gas-phase ions from electrolyte ions in solution by ESI include (1) production of charged droplets at the electrospray capillary tip, (2) shrinkage of the charged droplets by solvent evaporation and repeated droplet disintegrations, and (3) release of ions into the gas phase. The details of these processes have been investigated in detail [9,51] and a brief summary is illustrated in Fig. 8.8. The voltage applied to the capillary tip of the electrospray ion source results in a high electric field that penetrates the solution at the capillary tip. When the electrospray tip is the positive electrode, negative charge is removed by electrolysis generating an excess of positive ions in the solution. The positive ions drift downfield the solution, that is away from the tip, toward the meniscus of the solution. Because of repulsion between the positive ions, the surface begins to expand, a Taylor cone forms, and, if the applied field is sufficiently high, a fine jet emerges from the cone tip which breaks up into small, charged droplets. The charged droplets produced by the spray shrink owing to solvent evaporation, while the charge remains constant. The

Hyphenation

281

I

atmospheric atmospheric pressure to pressure region vacuum interface I II

electrical contacts for iII 100 V spray voltage (3-5 kV) --] I I I metal coating I ground+

to mass "7 /1 analyzer

CEC column ....................! ........:...........'.- '........'F_ - ~".,--.

dielectric capillary tube

eath gas syringe pump delivering sheath liquid

II II

x,y,z-positioning

skimm

drying gas vacuum pump

Fig. 8.6. Schematic of a CEC--ESI-MS interface with triaxial probe arrangement, allowing addition of sheath liquid and sheath gas, with dielectric vacuum transfer conduit, and drying gas.

atmospheric atmospheric pressure to pressure region vacuum interface I

40-100 MQ resistor electrical contact for spray voltage (3-5 kV)

I

II

- ground metal connect

I"

/ I to mass analyzer

CEC column x,y,z-positioning

pulled silica

skim~

'1'

vacuum pump

Fig. 8.7. Schematic of a CEC--nanospray ESI-MS interface with conducting vacuum transfer conduit and heated capillary.

Referencespp. 313-316

Chapter 8

282

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chargeddropletcontainingone solvatedmultiplycharged desolvatedmultiply multiply chargedanalytemolecule gas-phase ion chargedgas-phaseion Fig. 8.8. Major processes involved in the formation of gas-phase ions by ESI operated in the positive ion mode. Enrichment of the liquid surface by positive ion leads to the formation of a cone and jet emitting droplets with excess of positive ions (a). Charged droplets shrink by evaporation and split into smaller droplets (b) and finally gas-phase ions (c).

decrease of droplet radius at constant droplet charge leads to an increase in the electrostatic repulsion of the charges at the surface until the droplets surpass the Raleigh stability limit [9], after which the droplets undergo fragmentation, generally referred to as coulombic fission. Repeated evaporation and coulombic fission finally result in the formation of desolvated gas-phase ions, either by the formation of ex-

Hyphenation

283

tremely small droplets that contain only a single ion (charged residue mechanism [52]) or by the emission of ions from very small and highly charged droplets (ion evaporation mechanism [53]). One notable feature of ESI is its ability to produce multiply charged gas-phase ions of macromolecules by the adduction of several protons at basic functional groups or by the removal of several protons from acidic functional groups [54]. The majority of mass spectrometers are limited to analyzing ions with mass-to-charge rations of only a few thousands, but by increasing the number of charges associated with a molecule, compounds with a molecular mass of several million yield m/z values within the mass range of conventional mass analyzers [55,56]. Thus, a typical positive ion spectrum of a single protein consists of a series of peaks, each of which represents a multiply charged ion of the intact protein having a specific number of protons attached to the basic sites of the amino acid sequence. The m/z values for the ions have the general form [M+zH]/z, where z equals the number of protons attached. It follows that the molecular mass can be readily calculated from two measured, adjacent m/z values, given the additional information that the two multiply charged ions differ in charge by 1 [54]. Once M and z are determined for one pair of signals, all other m/z signals can be deconvoluted into one peak on a real mass scale, yielding the molecular mass typically with an accuracy of better than 0.01-0.1%. The submicroliter per minute flows through the capillary columns generally used in CEC are too low for establishing a stable electrospray under classical ESI conditions, which require flow rates of 1-10 ~tl/min. To tackle this discrepancy, addition of a make-up liquid to increase the flow of analyte solution into the ion source has been proposed. Moreover, the utilization of a conductive make-up liquid provides a means to establish an electrical contact between the exit of the separation column and an electrode, and thus completes the electrical circuit both for the CEC separation system and the electrospray ion source through the make-up liquid itself. Addition of the make-up liquid has been realized either by using a sheath liquid interface, which allows the simultaneous addition of sheath gas and the make-up liquid (the so-called "sheath liquid") at the tip of the electrospray needle by means of two concentric tubes [57,58], or with a liquid junction interface [59], where the make-up liquid is merged with the column effluent in a T-piece before the electrospray tip. Another approach for the interfacing of capillary liquid phase separation techniques with ESI-MS is the use of nano electrospray devices, which enable the formation of a stable electrospray at very thin capillary tips from nanoliter per minute flows of analyte solution without addition of make-up liquid or sheath gas [58,60,61 ]. In the arrangement of the most widely used sheath liquid interface illustrated in Fig. 8.6, the column is introduced into the atmospheric region of the ESI source through a coaxial, narrow metal tube, which delivers the sheath liquid to the column

Referencespp. 313-316

284

Chapter 8

exit. Additionally, a third concentric tube delivers a nitrogen gas flow to assist in spray formation and stabilization via nebulization. The sheath liquid, which essentially acts as the terminal electrolyte reservoir, is usually introduced at a flow rate of a few microliters per minute, whereas the electrosmotic flow through the capillary column contributes a few nanoliters per minute. In this way, the electrospray process is dominated by the sheath liquid. This arrangement is advantageous since a sheath liquid composition can be chosen with optimal electrospray characteristics to overcome possible problems associated with poor electrospray compatibility of the electrolyte used for the CEC separation. The sheath liquid is typically a solution of water containing a high percentage (50% or more) of an organic solvent (methanol, acetonitrile, isopropanol), that facilitates the droplet formation process and helps to maintain a stable spray by lowering the surface tension of the electrosprayed solution. Volatile acids, bases, or salts, such as formic acid, acetic acid, ammonia, or ammonium acetate, at concentrations of 0.1-1% are usually added to make the sheath liquid conductive. Since the potential required to initiate the electrospray process decreases with a reduction in the diameter of the tip from which the spray is generated [9], very sharp capillary tips allow the formation of a stable spray from very small flows of neat aqueous solutions before the onset of corona discharge without the necessity for sheath gas or sheath liquid. In the sheathless ESI-MS interface, the CEC column exit is carefully sharpened or connected to a short capillary tube which has been pulled to a fine point of a few micrometers outer diameter (Fig. 8.7). Electrical contact is possible upon coating of the tip with a thin, conductive layer of metal which is easily connected to the high voltage power supply of the ESI source. However, the metal coating present at the sharpened tips has only a limited lifetime. Alternatively, CEC column and spray tip can be joined through a metal union, that is used to establish electrical connection (Fig. 8.7). Since there is no outlet electrolyte reservoir present in this configuration, the sheathless interface works only properly when the electrosmotic flow is greater than the electrophoretic mobility of ionic species moving against the electrosmotic flow, otherwise depletion of these ions in the electrolyte will occur. In the sheath liquid approach the column effluent is diluted by the sheath liquid and hence it results in a lower sensitivity compared to the sheathless interface. Moreover, ions in the sheath liquid compete with the analytes for the transport of charge during the electrospray process, again resulting in a loss of sensitivity [62]. Finally, because the sheath liquid technique involves the input of additional solvents and other chemicals, the potential to add significantly to the background noise exists. However, the use of a sheathless CEC--ESI-MS interface implies the need for specialized capillary and electrospray hardware, and thus is not as popular as the sheath liquid interface.

Hyphenation

285

8.2.2.2.2 Implications of ESI on interface configuration As has already been discussed previously, CEC--ESI-MS interfacing is complicated by the need to complete electrical paths for both the CEC and ESI systems. The ESI source must apply a potential of several kilovolts between the CEC column exit and the entrance into the vacuum of the mass spectrometer in order to create the electrospray itself. The ideal arrangement to accomplish these requirements is to maintain the CEC column exit at ground potential and to apply high voltage to the counter electrode on the MS side of the interface. Achieving this goal places several demands on the MS entrance configuration. For a positive ion, the potential which first comes into contact with the ion beam at the point of supersonic gas expansion (skimmer in Fig. 8.6) and transfer into the vacuum system must be in the range of a few hundred positive volts. This condition is a result of basic ion optics and cannot be violated, regardless of the type of mass spectrometer [23]. When the conduit which samples the electrosprayed ions from the atmospheric region and transfers them into vacuum is a conductive metal tube as illustrated in Fig. 8.7, it clearly mandates that the entrance of this conduit, which serves simultaneously as the ESI counter electrode, must have a positive potential of a few hundred positive volts. In this case, the only way to create an electric field for ESI in the positive ion mode is to apply a high positive potential to the electrospray needle and thus the CEC column outlet. Another complication of this system arises due to necessity to transport the current flowing through the CEC capillary column from the ESI tip to the grounded counter electrode of the ion source. Since this current through the gas phase is transported by the charged droplets of the electrospray, a certain potential difference between the electrospray tip and the grounded electrode is required to effect a given current. Depending on the current that flows through the column, this potential may be higher than the optimal potential for the electrospray process. Moreover, if a relatively high current has to be carried by the electrospray, the relative portion of charge carried by analyte ions in relation to that carried by background electrolyte ions is very low, which results in poor sensitivity. To alleviate this problem, the electrospray high-voltage power supply and, hence, the outlet of the capillary column can be electrically connected to ground through a 40-100 Mr2 resistor (Fig. 8.7). This connection reduces the current from the capillary tip to the grounded counter electrode of the ion source and eliminates the occurrence of arcing. Moreover, it maintains the capability of regulating the ESI voltage independently of separation voltage, especially at CEC currents higher than 30 ~tA. If the electrode in the vacuum is isolated from the ESI counter electrode, then the voltages in the atmospheric region can be reversed essentially and the CEC column exit can be set to ground potential with the ESI counter electrode at a sufficient negative potential for the generation of positive ions. This can be achieved by replac-

Referencespp. 313-316

286

Chapter 8

ing the metal tube with a dielectric glass capillary which has a metal coating on both entrance and exit ends as illustrated in Fig. 8.6.

8.2.2.2.3 CEC--APCI-MS interface Although there are no principal restrictions to interface CEC to APCI-MS only very few information is available about the combination of these techniques [63]. Though, since APCI is very effective for the ionization of medium- and low-polarity compounds or when relatively non-polar solvents have to be used, its application as complementary ionization technique to ESI will most probably expand in the future. During APCI, the column effluent is nebulized pneumatically or electrically into a heated vaporizer tube, where the solvent evaporation is almost completed. In the gas-vapor mixture APCI is initiated by electrons that are emitted from a corona discharge needle. Analyte ionization is due to gas-phase ion-molecule reactions between analyte ions and reagent gas ions [64]. The reagent gas is generated from the solvent vapor in the high-pressure region of the ion source either by means of electrons or as a result of volatile electrolyte ions present in the mobile phase. The major positive reactand ions formed in the APCI source are proton hydrates, (H30[H20]n) +, whereas the superoxide anion (O2)-, its hydrates (O2[H20]n)-,

and clusters

(O2[O2]n)- are the major negative reactand ions [64,65]. These reactand ions may

react with analytes by charge transfer or proton transfer, yielding analyte ions that are extracted into the mass analyzer. An APCI interface for capillary electrophoresis that was capable of ionizing analytes from flows in the low microliter per minute range [66] was described. Whereas most commercial APCI interfaces are constructed to nebulize relatively high flows of analyte solution, is has been found difficult to establish stable nebulization using a pneumatic nebulizer. Therefore, a sheath-flow assisted electrospray-type nebulizer is used to obtain fine droplets in the design illustrated in Fig. 8.9. The sheath liquid delivered by a syringe pump is pure methanol. The fine spray is passed through a heated nebulizer made from a stainless steel block, which is heated to 300~

by

means of cartridge heaters. A corona needle is located between the vaporizer and ion sampling aperture, and a voltage of 3 kV is applied to the needle to induce corona discharge and ionization of analytes.

8.2.3 Mass analyzers Mass analyzers are devices which separate and detect ions according to their mass-to-charge (m/z) ratios. The choice of an appropriate mass analyzer that is used in combination with liquid phase separation techniques depends on a number of interrelated factors, including (1) the range of detectable m/z values (mass range), (2) ability to separate ions of closely similar m/z (resolution), (3) accuracy of m/z measurement

287

Hyphenation

electrical contact for corona voltage (3 kV) electrical contact for spray voltage (2-3 kV) ~] / CECcolumn

................................................. ' ','

syring:hPeumh Pliqelideringl/I| "

m

vaporizer I ~ ~ (300 ~ needle electrode,

to mass analyzer

I I

Fig. 8.9. Schematic of a CEC--APCI-MS interface with electrospray nebulization, heated vaporizer and corona discharge needle.

(mass accuracy), (4) scanning speed (usually quoted as time to scan one decade of m/z values), (5) dependence of signal on amount of analyte introduced into the mass analyzer (sensitivity and linear dynamic range), (6) minimum detectable amount of analyte (limit of detection), and (7) compatibility with liquid introduction interfaces. In addition to exact molecular mass determination, mass spectrometry can also provide significant information about how a molecule is put together. The clearest examples for structure determination come from the interpretation of electron impact spectra, in which a significant number of fragment ions are generated during the ionization process that can be related to specific structural features in the molecule [67]. However, mass spectra obtained by the soft ionization techniques used to obtain ions from liquid phase, such as FAB, ESI, and APCI, frequently do not contain significant fragment ions. Therefore, mass spectrometric techniques have been devised that allow fragmentation of ions after the ionization process. In tandem mass spectrometry (MS-MS), an ion is selected by the mass analyzer and forced to fragment by collisions with inert gas atoms or molecules, a process which is called collisionally induced dissociation (CID). The fragment ions are subsequently analyzed in a second mass analysis step, resulting a mass spectrum showing fragment ions of structural relevance. No one single mass analyzer is suitable for all applications and the choice of instrument is determined by the type of problem under investigation [68]. Double focusing magnetic/electrostatic sectorfield-, quadrupole-, quadrupole ion trap-, time-

References pp. 313-316

288

Chapter 8

of-flight-, and ion cyclotron resonance mass analyzers are the principal types of mass analyzers encountered in analytical laboratories. These mass analyzers differ from each other in the fundamental principles underlying mass-to-charge discrimination, resulting in different performance characteristics. While the sectorfield and ion cyclotron resonance mass analyzers offer unparalleled resolution, mass accuracy, and mass range, they are difficult to operate, expensive, bulky, and require relatively complex interfaces. Therefore, quadrupole, quadrupole ion trap and time-of-flight mass analyzers are the most commonly used analyzers for hyphenation to liquid phase separation techniques, because of their comparatively low costs, easy tuning and operation, and moderate space consumption. Since the majority of CECmMS applications are run on quadrupole-, quadrupole ion trap, or time-of-flight mass analyzers in combination with ESI interfaces, the basic principles of these three mass analyzers and their qualifications for interfacing with ESI are briefly discussed in this section. For a more elaborate discussion the reader is referred to an overview given by Brunnee [69]. 8.2.3.1 Linear quadrupole mass analyzers The linear quadrupole mass analyzer actually is a mass filter. It consists of four rods with circular or, ideally, hyperbolic cross section that are placed in a radial array. Opposite rods are charged by a positive or negative DC potential onto which an oscillating radiofrequency potential is superimposed. Ions are introduced into the quadrupole field by means of a low accelerating potential, typically 10-20 V. The ions traveling longitudinally between the four symmetrically arranged rods maintain their velocity along this axis while they are subjected to the influence of a total electric field made up of a quadrupolar alternative field superimposed on a constant field resulting from the application of the potentials upon the rods. They will experience attractive and deflective forces in the plane perpendicular to the rod length [70]. Therefore, the ions begin to oscillate as they traverse through the quadrupole filter. At a given combination of DC potential and radiofrequency potential, only ions of a narrow m/z range have stable trajectories through the quadrupoles, while all other ions cannot pass the mass filter, because the amplitudes of their oscillations increase until they are lost from the filter. By systematic variation of the applied potentials, ions of different m/z can consecutively pass the linear quadrupole filter towards the detector where their abundance is measured, which finally results in the recording of a mass spectrum. Another important operation of a linear quadrupole is the radiofrequencyonly mode, where the DC potential is zero. The quadrupole then acts as ion transmission device, which has the property of focusing ions of a broad m/z range toward a stable trajectory toward in the center of the quadrupole. Quadrupole instruments have become extremely popular for mass analysis owing

Hyphenation

289

I

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Fig. 8.10. Schematic illustration of a triple quadrupole mass analyzer with electrospray ion source.

mainly to their relatively low cost, small dimensions and weight, and ease of operation by nonspecialist operators. Low source voltages provide an added safety factor and the ability to withstand higher source pressures results in an ease of interfacing with liquid phase inlet systems and interfaces. The arrangement in a modem triple quadruple mass spectrometer with ESI ion source is depicted in Fig. 8.10. In this instrument, mass analysis is performed in the first and third quadrupoles, while the second quadruple can be used as collision cell in the radiofrequency-only mode. The collision cell is a box with small inlet and outlet holes which contains a quadrupole operated in the radiofrequency-only mode for ion focusing and which can be filled with collision gas pressurized to 0.1-1 Pa, such as argon or nitrogen. In modem triple quadrupole instruments hexapoles or octapoles are used instead of quadrupoles, because these provide higher ion transmission. An MS/MS experiment is performed with a triple quadrupole instrument by selecting ions of one specific m/z ratio in the first quadrupole mass analyzer, fragmenting them by collision with inert gas atoms in the collision cell, and finally analyzing the fragment ions in the second quadrupole mass analyzer. 8.2.3.2 Quadrupole ion trap mass analyzers The quadrupole ion trap is a three dimensional analogue of the linear quadrupole mass analyzer [71,72]. It consists of a cylindrical ring electrode and two end-cap electrodes. Both end-cap electrodes contain a whole for injecting and ejecting ions into and out of the ion trap (Fig. 8.11). A relatively high pressure of helium damping gas (about 0.1-0.4 Pa) is present in the ion trap in order to kinetically cool the trapped

References pp. 313-316

290

Chapter 8

ESl-source

sheath-gas sample sheath liquid

octapole lens

heated

capillary ~ ,5

L_.

II ~ I~

octapole lens

ion trap

/

I rotary pump

,cN~V{ end-cap electrodes

turbo pump

detector

He damping

gas

Fig. 8.11. Schematic illustration of a quadrupole ion trap mass analyzer with electrospray ion source.

ions to the center of the trap due to collisions, which substantially increases the sensitivity and mass resolution of the instrument. A three-dimensional quadrupole field is formed by the three electrodes when a suitable radiofrequency potential is applied to the ring electrode. This quadrupole field traps ions by continuously forcing them toward the center of the trap. When the amplitude of the ring electrode radiofrequency voltage is low, all ions above a minimum m/z are trapped. This radiofrequency voltage is referred to as the storage voltage. In an m/z selective stability scan, the ring electrode radiofrequency voltage is ramped causing ions of successively increasing m/z to adopt unstable trajectories and to exit from the trap through the wholes in the end cap electrodes, so that they can be detected by using a conversion dynode and an electron multiplier. The voltage at which an ion is ejected from the mass analyzer is defined as its resonance voltage. From the above it follows that with an ion trap the mass analysis process is a discontinuous, pulsed process. First, ions are injected from the external ion source through the ion optics into the ion trap by electrostatically "opening" the interoctapole lens (Fig. 8.11) for a short period of time in order to fill the ion trap with ions. Subsequently, in full scan mode, the ions of different m/z are consecutively ejected and detected by the detection system. In contrast to triple quadrupole instruments, where MS-MS experiments can be conducted in space in separate regions of the instrument, ion traps enable MS-MS sequentially in the same physical space, and thus, occur tandem in time. After the ions have been formed an trapped, a parent ion is selected by resonance ejection of all ions except those of the selected m/z ratio. This is done by applying a resonance ejection radiofrequency voltage to the end-cap electrodes which stimulates motion of the ions in the axial direction. The next step in the MS-MS sequence is to effect collisionally

Hyphenation

291

induced dissociation of the selected ion, which is achieved as result of energetic collisions with the helium damping gas. Resonance excitation is used to increase the kinetic energy of the ion, while keeping the amplitude low enough to avoid resonance ejection. The MS/MS spectrum is then recorded by sequentially ejecting the product ions. Due to the capability to perform multiple stages of mass spectrometry, separated in time, in a single mass analyzer, the number of sequential MS/MS operations in ion traps is virtually limited only by sensitivity constraints, and experiments involving 10 or more stages of fragmentation and mass analysis have been demonstrated.

8.2.3.3 Time-of-flight analyzers The time-of-flight (TOF) analyzer is conceptually the simplest of the mass analyzers in common use. It is based on the fact that ions of different m/z but the same kinetic energy leaving the ion source simultaneously as ion packet will take different times to drift toward the detector through a drift tube of defined length [73]. Time-offlight mass analysis is, therefore, used with a source that can provide short pulses of ions. These pulses typically have a frequency of 10-50 kHz and a lifetime of 25 ~ts. The ions produced in this way are then accelerated by an electric field pulse of 1000-10000 V. Subsequently, the accelerated ions travel through a field-free drift tube, about 0.5-3 m in length. Because all ions entering the drift tube have the same kinetic energy, their velocities must vary inversely with the square root of their m/z ratio, thus ions of small m/z arrive at the detector earlier than those of larger m/z. Since typical flight times are 1-100 ~ts, the use of a detector with very fast time resolution is required. A mass spectrum is obtained by measuring the time taken by ions of different m/z to travel through the drift tube. Modern time-of-flight instruments have many intrinsic advantages, especially for coupling with separation techniques. Because the short flight times, an entire mass spectrum can be obtained within a few tenths of a millisecond, thus allowing a very high data acquisition rate. Moreover, ion transmission is very high in time-of-flight mass analyzers (20-100%) which makes them very sensitive. The usable mass range of time-of-flight mass analyzers is very broad since there is no mass discrimination. In its simplest version, an ESI-time-of-flight mass spectometer comprises an ion source which continuously generates an ion beam from the analyte solution, an ion pulsing optics which deflects a portion of the beam orthogonally into the flight tube, a linear field-free drift region, and a detector, normally a multichannel array, which is capable of resolving the resulting tiny differences in flight time of the arriving ion packets (Fig. 8.12). Due to the distribution of translational directions and kinetic energies and the spatial spread of the ion pulses leaving the ion source, the mass resolution of a

References pp. 313-316

Chapter 8

292

detector

ions ~

of m/z1

9~

ions of m/z2(~ (~ ~

(~

field-free drift region

drying gas

sheath-gas ~

sample sheath liquid

s,0ro e , ~<

ion guiding optics

~,___] dielectric conduit '~ ~ 9

i

9 ion pulsing optics

/

Fig. 8.12. Schematic illustration of a time-of-flight mass analyzer with an orthogonal electrospray ion source.

conventional linear time-of-flight instrument is only moderate. To enhance the mass resolution of time-of-flight mass analyzers, an electrostatic reflector device can be added to the linear flight tube. This reflectron focuses the ion beam on the detector, compensating for the initial energy spread of the ions, because the faster ions penetrate more deeply into the reflector than the slower ions and thus have a somewhat longer path to the detector. Classical MS-MS experiments by selecting one ion species in a first mass analysis step, followed by fragmentation and another mass analysis are not feasible with simple time-of-flight mass spectrometers. Therefore, hybrid instruments,

such

as

linear

quadrupole-time-of-flight

or

quadrupole

ion

trap-time-of-flight combinations are available. Alternatively, the reflector can be used to perform MS/MS experiments [74]. Many metastable ions dissociate after acceleration, and fragments can be generated by collisions. In the linear mode, these arrive simultaneously with the precursor ions, since no field is present to induce separation. If the voltage of the reflectron is then changed stepwise, such ions can be separated prior to arrival at the detector, since their masses are different.

Hyphenation

293

8.2.3.4 Comparison of mass analyzers used for CEC--ESI-MS A practical comparison of mass analyzers is given in Table 8.1. Although the performance characteristics of commercial mass analyzers may vary considerably and prototype instruments have been constructed that show exceptional properties, the table tries to give some rough values that are representative for commercial routine mass spectrometers. TABLE 8.1 COMPARISON OF MASS ANALYZERS FOR CEC--ESI-MS

Mass analyzer/ Feature

Linear Quadrupole

Quadrupole ion trap

Linear time-of-fight

Reflectron time-of-flight

Upper m/z limit

4000

6000

> 10000

> 10000

Resolution

medium

high

high

very high

Scanning speed for 0.5-1 spectra per 1-4 spectra per 1000 m/z range second second

10-50spectra per second

10-50spectra per second

Mass accuracy

100 ppm

100 ppm

5-10 ppm

5-10 ppm

Sensitivity

medium

high

high

high

8.2.4 Column technology for C E C m M S

Despite considerable advances in packing technology, the present day status of manufacturing capillary columns for CEC is more of an art than a science [75]. Two major techniques for packing capillary columns have emerged over the past years. These are pressure filtration [76] and electrokinetic packing [77]. Because of their flexibility, relatively inert surface, optical transparency for on-column detection with nanoliter detection volumes, relatively low cost, and availability in a wide range of diameters, fused silica capillaries are the almost ideal tubing material to prepare microcolumns. The majority of columns for CEC is obtained by packing microparticular stationary phases into fused silica capillaries of appropriate dimensions. Presently, stationary phases based on porous silica gel are most frequently used as packing materials. An important part of microcolumn technology is the placement of small frits at the column inlet and outlet that retain the packing material inside the capillary [78]. Heat-assisted sintering of silica particles is the method of choice to prepare stable and small size frits for CEC capillary columns. With siliceous station-

References pp. 313-316

Chapter 8

294

,ri, ~

internal

(~~

external

' taper

9 9aeooeo 9 .~= ~"-.e,,o "~= o o o : o,,~., '. -. ,~w eoo,,o ~. "=..oe".,I ,o~o:O~ ~ ,, 9~ , , o" v.OOoO

taper

Fig. 8.13. Different configurations for the termination of capillary columns suitable for retaining the stationary phase particles.

ary phases, the flits can be made directly from the packing material by sintering a tiny segment of silica particles with the help of an electrically heated filament ring [79]. The correct choice of flit material is important because adsorption of analytes that elute from the column causes decreased sample recovery and loss of column performance [80]. External or internal tapers offer an alternative means to retain the stationary phase particles in the column tube. Tapered column ends of approximately 10 lam inner diameter are narrow enough to form a liquid-permeable plug that completely retains stationary phases of 3 ~tm particle size [81,82]. External tapers are prepared by drawing the end of a fused silica capillary in a hot flame whereas internal tapers are obtained by sealing the end of a fused silica capillary in a hot flame and subsequently grinding an opening of 10 ~tm with the help of a ceramic tile (Fig. 8.13). Drawing a tapered end reduces both the inner and outer diameter, which makes such capillary columns with metal coating suitable for direct mounting in the sheathless ESI-MS interface. Such capillary tips are particularly fragile, require a great deal of care with handling, and have only limited lifetime. Internal tapers are more robust than external tapers, and are most conveniently used in combination with the sheath liquid ESI-MS interface [82]. Despite many advantages, CEC columns packed with microparticulate sorbents do have some limitations such as the relatively large void volume between the packed particles and the slow diffusional mass transfer of solutes into the stagnant mobile phase present in the pores of the separation medium [83,84]. Alternative approaches to alleviate the problem of mass transfer and intraparticular void volume are the concepts of monolithic chromatographic beds and open-tubular columns. In mono-

Hyphenation

@

295

packedcolumn

monolithiccolumn

Fig. 8.14. Configuration of the chromatographic bed in (a) packed and (b) monolithic capillary columns.

lithic columns, the separation medium consists of a continuous rod of a rigid, macroporous polymer which has no interstitial volume but only internal porosity consisting of micropores and macropores [85], whereas in open-tubular columns the stationary phase is immobilized at the inner wall of very thin capillaries with inner diameters of less than 20 m [86]. Because of the absence of intraparticular volume, all of the mobile phase is forced to flow through the large pores of the separation medium [87] or the open tube. According to theory, mass transport is enhanced by such convection [88-90] and has a positive effect on chromatographic efficiency [91 ]. The structure of the column bed in packed and monolithic capillary column is compared in Fig. 8.14. The macroporous structure is generally achieved as a result of the phase separation which occurs during the polymerization of a monomer or monomer mixture containing appropriate amounts of both a crosslinking monomer and a porogenic solvent or a mixture of porogenic solvents [92-94]. A significant advantage of monolithic columns for CEC--MS is the possibility to immobilize the rod-shaped polymer covalently at the fused silica wall through anchoring groups which keeps the References pp. 313-316

Chapter 8

296

column packing in place without the need for frits [95]. Open-tubular columns do not need frits because of the absence of any particular or continuous chromatographic bed. In spite of these advantages, open-tubular columns are not widely used for CEC separations, mainly because of the difficulties associated with sample injection, detection, column preparation, and clogging. 8.3 STATIONARY PHASE-MOBILE PHASE SYSTEMS USED FOR C E C - MS 8.3.1 Stationary phases

Usually, CEC is performed in 20-200 ~m i.d. fused silica capillary columns packed with a suitable stationary phase. The requirements posed on the stationary phase are that it should have a large and readily accessible surface with favorable retention thermodynamics and kinetics [96]. In CEC the column packing that serves as a stationary phase plays a dual role. It has to have a charged surface with an appropriate zeta potential to generate and maintain electrosmotic flow as well as a chromatographic surface to bring about the retention of the sample components with the selectivity required for their separation. Furthermore, the support should have a structure which facilitates rapid mass transfer between stationary and mobile phase in order to minimize band spreading and thus make it possible to obtain high column efficiency [97]. It has been customary to classify the various techniques of liquid chromatography according to the stationary phase employed. By virtue of its high selectivity and resolving power, reversed-phase liquid chromatography is the most popular branch of liquid chromatography. Reversed-phase liquid chromatography separates analytes on the basis of differences in their hydrophobic properties. The stationary phase in reversed-phase liquid chromatography is a very hydrophobic, nonpolar surface, whereas the mobile phase is polar, usually a mixture of water and organic solvents. Because of the volatility of the applied eluents, reversed-phase liquid chromatography has also emerged as the most important separation technique for CEC--MS coupling. In reversed-phase CEC, separation of the analytes is achieved through partitioning between the mobile and stationary phase and, if they are charged, through differential electromigration under the influence of the electric field (see 8.2.1.2). The great majority of CECmMS applications is run in the reversed-phase mode using alkyl-bonded silica stationary phases [12,14,24,26,38,45,81,82,98-104]. The dual functionality concept is represented in these stationary phases by the alkyl chains, most frequently octadecyl chains, that constitute the top retentive layer, and residual silanol groups on the surface, that dissociate at pH values higher than 3-4 and

Hyphenation

297

hence generate a zeta potential. Linear electrosmotic flow velocities in the range of 1-3 mm/s are obtained with such octadecyl silica stationary phases at an electric field strength of 600 V/cm and with a 10 mmol/1 borate electrolyte, pH 8.0, containing 80% acetonitrile [20]. The concept of dual function has also been realized with mixed mode stationary phases, where the silica surface is derivatized both with alkyl groups and ion-exchange functionalities. Examples for such combinations are alkyl- and sulfonic acid groups [58,105-107], or alkyl- and dialkylamino groups [108] on the same support particle. With the latter combination, the direction of electosmotic flow is reversed from cathodic to anodic. Since the sulfonic acid functional groups are ionized over the entire pH range, sufficient cathodic electrosmotic flow can be obtained even with eluents having acidic pH. Moreover, compared to octadecyl silica stationary phases, higher column efficiencies have been observed with hexyl/sulfonic acid stationary phases, especially for basic compounds [107]. The need of column configurations and surface chemistries especially designed for CEC is now generally appreciated and novel approaches to improve the column technology for CEC--MS applications include the use of monolithic stationary phases [109,110], open-tubular capillary columns [86] and chip technology [111]. These configurations are currently under detailed investigation and the future will have to prove their applicability in routine analysis. 8.3.2 CEC eluents compatible with mass spectrometry

The most distinguishable feature of CEC is that it makes use of electrosmotic flow to drive the mobile phase through the capillary columns. Since in CEC the electrosmotic flow velocity strongly depends on the electric field strength as well as on the nature of both the stationary and mobile phase, the major disadvantage of electosmotic flow is the lack of reliable control over the flow velocity of the mobile phase. This is in strong contrast to HPLC, where the flow is generated by high precision metering pumps. Eluents for CEC comprise aqueous buffers or electrolytes mixed with organic modifiers such as acetonitrile, methanol, isopropanol, or tetrahydrofuran. The most widely used organic modifier is acetonitrile because of its high UV transparency. Ionogenic components are added to make the mobile phase conductive, to adjust or buffer the eluent to a desired pH, and sometimes to suppress ionic interactions between analytes and the charged surface of the stationary phase. Investigation of the effect of ionic strength on electrosmotic flow velocity reveals, that with increasing electrolyte concentration in the range of 5-100 mmol/1 the electosmotic flow velocity almost linearly decreases [112]. This is expected since the zeta potential decreases with increasing ionic strength resulting in a decrease in electrosmotic flow [76]. Another important consideration for the appropriate choice of a mobile phase for References pp. 313-316

298

Chapter 8

CEC--MS is the compatibility with the chosen ionization method. FAB ionization and APCI are relatively insensitive to the mobile phase composition. Yet, for FAB ionization, no other non volatile components than the matrix should be present, because solid deposits will efficiently inhibit the desorption process at the FAB target. The ESI process, however, is strongly determined by the composition of the electrosprayed solution. Generally, highest ESI-MS sensitivity will be obtained at the lowest CEC currents, where the rate of delivery of charged species through the electrospray is minimized and the relative portion of current transported by the analyte ions is maximized [9]. Some commercial ESI interface designs enable the use of mobile phases containing non-volatile additives, e. g. phosphate, borate [81,98,99], sodium chloride, because an orthogonal- or Z-spray geometry prevents the entrance of solid particles into the mass spectrometer. Still, deposition of solids rapidly results in fouling of the surfaces in the ion source, which usually constitutes a non optimal condition. Therefore, optimal eluents should contain exclusively volatile buffer additives, such as formic acid, trifluoroacetic acid [12,45], acetic acid, ammonium formate, ammonium acetate [26,38,82,100], ammonia, or volatile organic amines. The conductivity of the mobile phase should be kept at the lowest practical level by using low concentrations of volatile additives of small equivalent conductivity. In addition, additives that interact strongly with the analytes (e. g. denaturants) substantially degrade ESI-MS sensitivity, largely due to association in the gas phase. Surfactants as components of the mobile phase have to be avoided because they give rise to intense signals in ESI-MS. If the electrolyte contains alkali metal salts, metal ion adducts can be formed, such as (M+Na)+, (M-H+2Na)+, etc. Sometimes this kind of adduction is a desired effect for ionization of compounds of low proton affinity, such as carbohydrates. Nevertheless, if multiple adduction between metal ions and analytes such as nucleic acids or polypeptides is possible the ion current is divided over several species, reducing the overall sensitivity of MS detection. 8.4

OPTIMIZATION OF ELECTROCHROMATOGRAPHIC AND MASS SPECTROMETRIC CONDITIONS

Optimization of electrochromatographic conditions includes the appropriate choice of stationary phase functionalities, stationary phase particle size and pore diameter, column dimensions, solvent strength of the mobile phase, mobile phase additives, separation voltage, isocratic or gradient elution conditions. Considerations regarding stationary phase and mobile phase chemistry have already been discussed in the previous section. Small stationary phase particle sizes are desirable, because they afford columns with a high number of theoretical plates. Since most of the chromatographic surface is provided by the inner surface in the pores of the packing,

Hyphenation

299

A

V

t

r. t_ !__

I 2

r r.

o

,.

I 4

I 6

I 8

P

Q

0 ,4.=1

T I

0

I

2

I

4

AY

I

6

I

8

I

10

time [min]

Fig. 8.15. Effect of particle diameter and pore diameter on the separation of 12 PTH-amino acids. Column, 150 x 0.075 mm i.d. packed with 6 ~tm/300 A or (b) 3.5 ~tm/80 A Zorbax ODS; eluents, (A) 2 mmol/1 ammonium acetate, pH 7.0, (B) 2 mmol/1 ammonium acetate, pH 7.0, 90% acetonitrile; gradient elution with 30-90% B in 15 min; flow rate of mobile phase through inlet reservoir, 100 ~tl/min; applied voltage, 20 kV; detection, ESI-MS, 0.5 s/spectrum integration time; sheath liquid, 1 mmol/1 ammonium acetate, pH 7.0, 90% methanol, 3 ~tl/min; injection, electrokinetic, 2 kV, 2 s; sample, PTH-asparagine, PTH-glutamine, PTH-threonine, PTH-glycine, PTH-alanine; PTH-tyrosine, PTH-valine, PTH-proline; PTH-tryptophan, PTH-phenylalanine, PTH-isoleucine, PTH-leucine (in order of elution). (Reproduced from ref. [ 113] with permission of the author).

stationary phases with small pore diameters offer the highest specific surface area. The selection of an optimal pore size for a particular sorbent is made on the basis that the solute molecular diameter must be at least one tenth the size of the pore diameter of the packing material to avoid restricted diffusion of the solute and to allow the total surface of the sorbent material to be accessible. Therefore, 8-15 nm pore sizes are suitable for separating small molecules, whereas larger pore sizes of 15-100 nm or totally non-porous packing materials are needed for macromolecules. In CEC, the effect of particle diameter and pore diameter on the separation efficiency is similar to that in HPLC. Fig. 8.15 illustrates the gradient separation of PTH-amino acids in 150 • 0.075 mm i.d. columns packed with either 6 ~m/300 A or 3.5 ~tm/80 A Zorbax ODS particles at an applied voltage of 20 kV [82,113]. It is seen, that the first six PTHamino acids can not be resolved on the 6 ~tm Zorbax ODS stationary phase. In contrast, all 12 sample components were well resolved on the 3.5 ~tm stationary Referencespp. 313-316

Chapter 8

300

Q

./, e" I

i

I

I

L_

o e,,

.2 0

@ 0

1'0 time [rain]

Fig. 8.16. Effect of applied voltage on the separation of PTH amino acids. Column, 150 • 0.075 mm i.d. packed with 3.5 ~tm/80 A Zorbax ODS; eluents, (A) 2 mmol/1 ammonium acetate, pH 7.0, (I3) 2 mmol/1 ammonium acetate, pH 7.0, 90% acetonitrile; gradient elution with 30-90% B in 15 min; flow rate of mobile phase through inlet reservoir, 100 ~tl/min; applied voltage, (a) 15, (b) 20, (c) 25 kV; detection, ESI-MS, 0.5 s/spectrum integration time; sheath liquid, 0.2 mmol/1 ammonium acetate, pH 7.0, 90% methanol, 3 ~tl/min; injection, electrokinetic, 2 kV, 2 s; sample, PTH-asparagine, PTH-glutamine, PTH-threonine, PTH-glycine, PTH-alanine; PTH-tyrosine, PTH-valine, PTH-proline; PTH-tryptophan, PTH-phenylalanine, PTH-isoleucine, PTH-leucine (in order of elution). (Reproduced from ref. [82] with permission of Elsevier Sciences 13. V.).

phase. The length of the separation column for CEC--MS is usually dictated by the construction of the CEC unit and the interface dimensions. While most commercial systems require relatively long columns of 450-750 mm length, laboratory-made constructions allow the use of CEC columns as short as 150 mm [82]. The optimal column inner diameter depends on the conductivity of the mobile phase and the applied voltage. While large-diameter columns (100-360 pm i.d.) are advantageous for better detection sensitivity because more sample can be loaded onto such columns, the current through these columns may be too high for successful CEC--MS coupling. Therefore, column inner diameters of 20-75 ~m are most suitable for CEC--MS, especially with highly conductive eluents and high electric field strengths. The electrochromatograms presented in Fig. 8.16 demonstrate that applied voltage has a significant effect on the separation of PTH-amino acids [82]. At 20 kV (Fig. 8.16b), all sample components are separated to baseline, whereas at lower (Fig. 8.16a) and

301

Hyphenation

T

Y

| Q

T Y,A

@ I

I

2

I

4

time [rain] Fig. 8.17. Effect of gradient slope on the separation of PTH amino acids. Column, 150 • 0.075 mm i.d. packed with 3.5 ~tm/80 A Zorbax ODS; eluents, (A) 2 mmol/1 ammonium acetate, pH 7.0, (B) 2 mmol/1 ammonium acetate, pH 7.0, 90% acetonitrile; gradient elution with (a) 30-80% B and (b) 30-60% B in 5 min; flow rate of mobile phase through inlet reservoir, 100 ktl/min; applied voltage, 20 kV; detection, ESI-MS, 0.5 s/spectrum integration time; sheath liquid, 0.2 mmol/1 ammonium acetate, pH 7.0, 90% methanol, 3 ktl/min; injection, electrokinetic, 2 kV, 2 s; sample, PTH-asparagine, PTH-glutamine, PTH-threonine, PTH-glycine, PTH-alanine, PTH-tyrosine (in order of elution). (Reproduced from ref. [82] with permission of Elsevier Sciences B. V.).

higher (Fig. 8.16c) no complete resolution is obtained. As seen in Fig. 8.16a, PTHproline and PTH-tryptophan as well as PTH-isoleucine and PTH-leucine are not completely separated at 15 kV applied voltage. Similarly, at 25 kV the resolution between PTH-valine and PTH-proline as well as PTH-isoleucine and PTH-leucine decreased. Moreover, upon increasing the applied voltage from 15 to 25 kV the time required to complete the separation was reduced by about 3 min. The effect of changing the gradient slope on the resolution of PTH-amino acids is illustrated in Fig. 8.17 [82]. At an applied voltage of 20 kV, a 5-min linear gradient from 27-54% acetonitrile suffices to completely resolve the 6 PTH-amino acids within 5 min as shown in Fig. 8.17a. In order to decrease the overall analysis time a steeper slope from 27-72% acetonitrile in 5 min can be used. It is seen that upon increasing the gradient slope to 27-81% acetonitrile in 5 min all sample components elute faster, but the overall resolution deteriorates significantly (Fig. 8.17b). The results strongly indicate that the effect of gradient slope on the separation in reversedReferences pp. 313-316

302

Chapter 8

phase CEC is similar to that in reversed-phase HPLC. Several parameters of the ESI emitter, e. g. the inner and outer diameters of the spray tip and its position relative to the sampling orifice, have to be optimized to improve ionization efficiency and ion sampling [114]. It has been noted that the production of electrospray with small plumes enhances sampling efficiency. Therefore, the spray performance can be improved by adjusting the flow-rate and tip diameters, which determine the formation and size of the droplets from which the ions will desorb. ESI functions best with solutions having high concentration of organic solvent and low ionic strength. Through the addition of a coaxial flow of sheath liquid to the column effluent solvent conditions otherwise inappropriate for ESI-MS can be used for the separation and detection of the analytes. The idea of using coaxial flow was originally developed for the postcolumn derivatization of analytes [115,116]. Later, addition of a sheath liquid was proposed for the coupling of CE and ESI-MS [57]. Moreover, sheath liquids that contain volatile acids or bases offer the possibility of manipulating the charge state distribution of multiply charged ions [ 117,118]. The effect of sheath liquid composition on the separation and ESI-MS detection of PTHamino acids is illustrated in Fig. 8.18. With neat methanol as the sheath liquid, the migration times of the analytes are quite long, as seen in Fig. 8.18a. A possible explanation for this rests with the absence of electrolyte in the sheath liquid and the concomitant low conductivity of neat methanol. Upon including 0.2 mmol/1 ammonium acetate in the sheath liquid, the components elute in less than half of the time as before (Fig. 8.18b). Because of the faster migration, some of the resolution is sacrificed and the signal intensity is six times lower than that obtained with neat methanol as sheath, which is attributed to the charge competition between ions supplied by the sheath liquid and analyte ions. Weakly polar or nonpolar substances yield very poor ESI sensitivity, because protonation or deprotonation is not possible in the absence of acidic or basic functional groups. However, charged coordination compounds can be formed upon the addition of a central complexing ion, which allows the detection of the charged complex by ESI-MS with high sensitivity. This technique that has been termed "coordination ion spray mass spectrometry" (CIS) by Bayer et al. [119]. The complexing ion is added to the column effluent post-column through a triaxial sheath liquid interface. Fig. 8.19 shows a coordination ionspray mass spectrum with silver ions as complexing ions, which proves the interaction of silver with estradiol. The coordination capability of silver ions has also been demonstrated for unsaturated fatty acid methyl esters and vitamins [104], which proves that this approach is applicable to a wide range of medium to non polar analytes, and thus represents an ionization method complementary to conventional ESI. The realization of high efficiency CEC--MS separations, leading to peak widths

Hyphenation

303

17--

G

Q

.4.,.,

/I

|

r==-l

0-

I

t=. t_ :3

!

I

s

1

12

N,Q

t::: 3 - o

I

A

Y

.,==

o

T

@ i

0

2

I

4 time [min]

i

6

Fig. 8.18. Effect of sheath liquid composition on the separation of PTH amino acids. Column, 250 • 0.075 mm i.d. packed with 3.5 ~m/80 A Zorbax ODS; eluents, (A) 2 mmol/1 ammonium acetate, pH 7.0, (B) 2 mmol/1 ammonium acetate, pH 7.0, 90% acetonitrile; gradient elution with 30-80% B in 5 min; flow rate of mobile phase through inlet reservoir, 100 ~tl/min; applied voltage, 20 kV; detection, ESI-MS, 0.5 s/spectrum integration time; sheath liquid, (a) neat methanol, (b) 0.2 mmol/1 ammonium acetate, pH 7.0, 90% methanol, 3 ~l/min; injection, electrokinetic, 2 kV, 2 s; sample, PTH-asparagine, PTH-glutamine, PTHthreonine, PTH-glycine, PTH-tyrosine, PTH-alanine (in order of elution). (Reproduced from ref. [82] with permission of Elsevier Sciences B. V.).

of only a few seconds, presents challenges to most mass spectrometers. The majority of mass analyzers are based on the use of changing electric and/or magnetic fields to selectively detect ions across the m/z range. The result of this process is that 1-20 s are routinely required to complete a single scan. The effect of scan time on observed peak shape can be deduced from the electrochromatograms shown in Fig. 8.20 [82]. The base width of the PTH-asparagine peak as derived from the reconstructed ion chromatogram is about 6 s. At a scanning speed of 2s/spectrum only three data points are acquired under the peak, and the peak shape is distorted due to artificial band broadening (Fig. 8.20a). Time-of-flight mass analyzers have the potential to rapidly collect m/z data. Because the flight times are short and no electric or magnetic fields have to be varied with time, complete mass spectra can be acquired at rates of several

Referencespp. 313-316

Chapter 8

304

+

H3C

OH

<

i HO §

< r

+

+

o Z

<~

9

I

400

o ~

§,-g

z g

O ~-

m

<

+~ "I

§,--g 0

+~ <

~"

'-"

,

I

600

"

+ 'q.

+

13}

I

800

(m/z) Fig. 8.19. Coordination ion spray mass spectrum of estradiol extracted from the reversed-phase CEC--CIS-MS separation of estrogenic compounds. Column, 250 x 0.1 mm i.d. packed with 3 ~m GROM-SIL ODS-0 AB; eluent, 4 mmol/1 ammonium acetate, pH 9.0, 50% acetonitrile; applied pressure, 8 MPa; applied voltage, 15 kV; detection, ESI-MS, 0.85 s/spectrum; sheath liquid, 100 ~tg/ml aqueous silver nitrate, 3 ~l/min; mass spectrum extracted from a reconstructed ion chromatogram of the separation of estriol, estradiol, equiline, and estrone. (Reproduced from ref. [ 104] with permission of Wiley-VCh).

kilohertz. Because it is impractical and unnecessary to individually store so many records, the mass spectra are integrated over a time frame which is appropriate for the peak widths of the separation. Upon decreasing the integration time to 0.25 s/spectrum, 24 data points could be acquired for the PTH-asparagine peak, which allowed to accurately portray the characteristics of the actual separation. While the physics of time-of-flight mass analysis enables the rapid scanning of a mass spectrum within a fraction of a second, the scanning speed with linear quadrupole and quadrupole ion trap mass analyzers can only be increased at the cost of resolution and signal-to-noise ratio. Therefore, usually a compromise has to be found between optimal characterization of electrochromatographic peaks and quality of the recorded mass spectra [120,121].

305

Hyphenation

L_ L_ U

T

'-

yA

-

0

3.0

'

410 3a.0

I

4t.0

time [min] Fig. 8.20. Effect of integration time on reconstructed ion chromatograms for the separation of PTH amino acids. Column, 250 x 0.075 mm i.d. packed with 3.5 ~tm/80 A Zorbax ODS; eluents, (A) 2 mmol/1 ammonium acetate, pH 7.0, (B) 2 mmol/1 ammonium acetate, pH 7.0, 90% acetonitrile; gradient elution with 30-80% B in 5 min; flow rate of mobile phase through inlet reservoir, 100 ~tl/min; applied voltage, 30 kV; detection, ESI-MS, (a) 2, (b) 0.25 s/spectrum integration time; sheath liquid, 0.2 mmol/1 ammonium acetate, pH 7.0, 90% methanol, 3 ~tl/min; injection, electrokinetic, 2 kV, 2 s; sample, PTH-asparagine, PTH-glutamine, PTH-threonine, PTH-glycine, PTH-tyrosine, PTH-alanine (in order of elution). (Reproduced from ref. [82] with permission of Elsevier Sciences B. V.).

8.5 EXAMPLES OF A P P L I C A T I O N

CEC--MS has now been applied to the analysis of a wide range of species, from small ions to biomolecules. Since CEC--MS is a less familiar technique and instrumentation and column technology still needs to be improved, the first wave of reports are those investigating the general applicability of the technique to solve analytical problems. Yet, applications of CEC--MS to routine, qualitative and quantitative analysis are relatively few. Gradually, however, more real problems are being examined and the number of reports describing such real application is expected to increase rapidly. 8.5.1 Small molecules

An electrochromatographic separation system with both pressure- and electrodriven flow was coupled to a mass spectrometer using a CFFAB interface by Verheij et al., who termed the method "pseudo-electrochromatography" [24]. Capillary col-

umns of 200 ~tm inner diameter packed with a Nucleosil 100-5C18 stationary phase were used to separate morphine alkaloids, nucleotides, and antiviral drugs a flow rate of 2-4 ~tl/min and 5-10 kV applied voltage. The column effluent was mixed post-column with a make up liquid containing 15% glycerol and conducted to the CFFAB ion

References pp. 313-316

306

Chapter 8

277

209 testosterone

361 369

369

,,wfllj,I ...,.,,,,,,..,, ........ , ...... I ....... I...... I...... I

aldosterone

hydrocortisone

277

280

277

36:

'

I

320 m/z

'

I

360

,,,,I,, , ,I,,I, , , , ,q, ,,, ,iI]~tLl , tl,ltL. t

280

320

m/z

36O

]llh

,,,,,,,J,............ ,,,,,,,,,I,,,,I,,,,,,I,,,,,,,,

280 I

'

320 I

360 m/z '

I

Fig. 8.21. FAB mass spectra of steroids separated by CEC. Column, 350 x 0.05 mm i.d. packed with 3 pm Hypersil ODS; eluent, 4 mmol/1 sodium tetraborate, 80% acetonitrile; applied voltage, 21.5 kV; detection, CFFAB-MS, 230-500 amu; matrix, post column addition of 1% glycerol in 50% water-50% methanol, 5 pl/min; sample, testosterone ([M+H] + 209), hydrocortisone ([M+H] § 363), aldosterone ([M+H] § 361) (in order of elution). (Reproduced from ref. [25] with permission of John Wiley & Sons).

source of a double focusing sectorfield mass spectrometer. An electrochromatographic separation system without additional pressure driven flow was used for the separation of steroids. They could be separated in a 350 x 0.05 mm i.d. capillary column packed with Hypersil-ODS particles using 4 mmol/1 sodium tetraborate, 80% acetonitrile as mobile phase. The analytes were detected by CFFAB-MS after post-column addition of glycerol as matrix compound. In this separation system, the enhanced performance of electrosmotically driven chromatography compared to hydraulically driven chromatography was clearly demonstrated [25]. The FAB mass spectra of three steroids taken from the electrochromatographic peaks are shown in Fig. 8.21. Although a sodium salt, albeit at low concentration, was used as mobile phase additive, this was found not to cause any problems and significant sodium adduct species are not observed in the spectra. The analysis of compounds of pharmaceutical relevance is one of the most promising application areas for CEC--MS, because it offers high sensitivity, high selectivity and structural information [38,98,99,105]. Paterson et al. [105] utilized CEC--ESIMS for the analysis of potential drug candidates down to the 1 ng/ml level in solid-phase extracts of plasma samples. Relative standard deviations of retention time and peak area of 0.4-1.7% and 2.6-10.7%, respectively, were achieved, which proves, that the method is also applicable to quantitative analysis. The analysis of a pair of

307

Hyphenation

cefuroxime axetil

~.,

r-.

oUoy~.,

L_ t_

0

CH 3

533 [M+Na] §

e-, o

400

500

600

700

rn/z

'

0

I

'

I

20

'

I

'

I

40 time [mini

'

I

'

I

60

Fig. 8.22. Reconstructed ion chromatogram from the CEC--ESI-MS analysis of cefuroxime axetil diastereomers. Column, 900 x 0.05 mm i.d. packed with 3 pm Hypersil ODSA; eluent, 5 mmol/1 sodium tetraborate, pH 9.0, 80% acetonitrile; applied voltage, 30 kV; detection, ESI-MS, 400-700 amu; sheath liquid, 0.3% aqueous formic acid, 50% methanol, 10 pl/min; injection, electrokinetic, 30 kV, 5 s. (Reproduced from ref. [98] with permission of John Wiley & Sons).

diasteroisomers of cefuroxime axetil, a semisynthetic broad-spectrum cephalosporin antibiotic for oral administration, is illustrated in Fig. 8.22. The separation was performed in a 450 x 0.1 mm i.d. Spherisorb ODS-1 column using an eluent comprising 5 mmol/1 sodium tetraborate-80% acetonitrile. The mass spectrum of the eluting compounds showed an abundant [M+Na] + signal at m/z 533 (inset in Fig. 8.22), indicating that adduction with sodium ions is the principal ionization mechanism for this compound. Mixtures of benzodiazepines and thiazide diuretic drugs were separated by gradient elution CEC and identified using ESI-MS by Taylor and Teale [38]. They used 330-500 x 50-75 lam i.d. colums packed with Hypersil ODS and Apex ODS and gradients of 50-80% acetonitrile in 5 mmol/1 aqueous ammonium acetate to elute the sample components. Benzodiazepines were detected in the positive ion mode using 1% acetic acid as the sheath liquid, whereas the thiazide diuretics were detected in the negative ion mode with 80% isopropanol in water as sheath liquid. Steroids are a group of hormones with similar chemical structures that help to control growth, metabolism, development of sexual characteristics, and the ability to withstand the stress of illness and injury. Because of their legal use as medicaments or illegal use as growth promoters, analysis and positive identification of trace amounts

References pp. 313-316

308

Chapter 8

bufalin

M + H * = 387

cinobufagin M + H * = 443 I"

4=1

L_ L_

digitoxig enin

o tO

+H + = 375

,,==

O cinobufatalin

~

lo

~ d i g o x i g e n i n M + H + = 391

l's

'

2'o

'

25

'

a'o

'

time [min]

Fig. 8.23. CEC--ESI-MS analysis of steroids. Column, 450 x 0.1 mm i.d. packed with 3 ~m Sherisorb ODS-1; eluent, 4 mmol/1 sodium tetraborate-sodium hydroxide, pH 8.0, 70% acetonitrile; applied voltage, 21.5 kV; detection, ESI-MS, 370-470 amu; sheath liquid, 0.1% formic acid, 50% methanol, 6 ~tl/min; injection, electrokinetic, 5 kV, 10 s; sample, bufalin, cinobufagein, digitoxigenin, cinobufatalin, digoxigenin, gitoxigenin (in order of elution). (Reproduced from ref. [81 ] with permission of John Wiley & Sons.).

of steroids is frequently necessary. Fig. 8.23 illustrates the selected ion chromatograms of the protonated molecules from the reversed-phase electrochromatographic separation of six bufadienolide- and cardenolide steroids [81]. For ESI mass spectrometric detection, a coaxial sheath liquid interface which allowed the addition of 0.1% formic acid in methanol as sheath liquid was used. The same component mixture was also analyzed using a sheathless interface and the authors observed less band broadening with the latter interface, as exemplified by peak widths at half height for digoxigenin of 19 s with the sheathless interface versus 27 s with the coaxial sheath interface. Moreover, a significant difference in the obtained mass spectra was noticed with both interfaces. In the sheathless approach, sodium adducts were the most prominent species in the mass spectrum, whilst the protonated molecule is the most abundant species in the mass spectrum after addition of the acidic sheath liquid. This example clearly demonstrates, that the addition of sheath liquid is an appropriate means of manipulating the ionization process in ESI-MS. The separation and ESI-MS

Hyphenation

309

detection of steroids has also been accomplished with mixed mode reversedphase/strong cation-exchange stationary phases [58,103]. Lower limits of detection were found to be in the 1 fmol range with the coaxial sheath interface and were predicted to be in the 100 amol range with the sheathless nanospray interface.

8.5.2 Polypeptides and enzymatic digests The development of C E C n M S techniques for analyzing small amounts of complex polypeptide mixtures and digests for sequence determination is a rapidly growing area of research, especially in proteomics. Standard peptides have been used for CEC--MS method development [12]. In a series of papers, Lubman et al. demonstrated the applicability of open-tubular- [86] and pressure assisted gradient CEC [ 14,45,102,108] coupled to electrospray ion trap storage/reflectron time-of-flight mass spectrometry for the analysis of peptides and protein digests (Fig. 8.24). Using this hybrid mass analyzer, fast spectra acquisition rates of up to 10 spectra per second and lower limits of detection in the low fmol range were demonstrated [ 14]. A major problem with the reversed-phase chromatographic analysis of peptides on silica-based stationary phases is the interaction of positively charged, basic residues of the polypeptide chain with dissociated silanol groups of the stationary phase, resulting in band broadening due to ion-exchange interactions. Therefore, strong acids are added to the eluent in order to suppress dissociation of the silanol groups. In CEC, however, suppression of silanol dissociation is undesirable because electrosmotic flow will be suppressed, too. A constant anodic electrosmotic flow over pH 2-5 could be maintained with acidic eluents containing 0.01-0.05% trifluoroacetic acid or 4 mmol/1 acetic acid-ammonium acetate using a mixed mode reversed-phase/weak anion-exchange stationary phase [ 108]. The positively charged amino groups on the stationary phase repel the peptides and thus do not act as ion-exchange sites. The separation mechanism for charged peptides in reversed-phase CEC is complicated, since both reversed-phase partitioning and electrophoretic migration contribute to the separation. Because the extent of partitioning and the electrophoretic mobility are two orthogonal properties of a peptide, there are cases where electrophoretic migration might improve or deteriorate a reversed-phase separation. Therefore, it is usually necessary to optimize the applied voltage in CEC of peptides. In a solely electrodriven CEC system a change in applied voltage is accompanied by both a change in electrophoretic migration and mobile phase flow rate. Consequently, optimization of separation selectivity using the applied field may be difficult. In pressurized CEC, however, the electrophoretic migration rate and the mobile phase flow rate can be optimized independently [45,108]. The effect of voltage on the selectivity during the separation of tryptic peptides from cytochrome C by pressure assisted CEC is depicted in Fig. 8.24. Using a 60 x 0.16 mm i.d. capillary column References pp. 313-316

310

Chapter 8

2 3

2 1

3

2

4

34

1

o

'

i

'

8

'

l'Z

'

1'6'

z'o

time [rain]

Fig. 8.24. Separation of a tryptic cytochrome C digest by pressure assisted gradient CEC. Column, 60 x 0.18 mm i.d. packed with 3 l.tm Vydac C18; eluents, (A) 0.07% trifluoroacetic acid in water, (B) 0.07% trifluoroacetic acid in acetonitrile; gradient elution with 0-50% B in 20 min; applied pressure, 9 MPa in (a), 5 MPa in (b); 7 MPa in (c); applied voltage, 0 kV in (a), 1 kV in (b), 0.6 kV in (c); Detection, nanospray ESI-MS, m/z 200-1500, 10 spectra/s; sample, tryptic digest of 8 pmol bovine cytochrome C. (Reprinted with permission from ref. [45]. Copyright [1997] American Chemical Society).

containing a 3 ~m Vydac C 18 stationary phase, the digest was separated into 9 peaks in the HPLC mode with a back pressure of 9 MPa (Fig. 8.24a). The use of a short column in the HPLC mode made it difficult to resolve all compounds in the digest, as indicated by peak number 3, which contained two coeluting peptides. After application of 1 kV voltage to the column and reduction of the backpressure to 5 MPa in order to keep the mobile phase flow rate constant, all peptides eluted earlier and as sharper peaks, which can be explained by the contribution of electrophoretic migration and by increased separation efficiency due to the electrosmotic flow component (Fig. 8.24b). The two coeluting compounds in peak number 3 in Fig. 8.24a were separated to baseline in Fig. 8.24b (marked with number 3 and 4), whereas the first two peaks marked by 1 and 2 in Fig. 8.24a migrated into one peak. Optimization of both pressure and voltage, enabled the separation of all four peaks at a voltage of 600 V and a supplementary pressure of 7 MPa (Fig. 8.24c). This experiment shows, that the combination of the enhanced separation performance of pressurized gradient CEC with the molecular mass information from ESI-MS permits the analysis of complex protein digests in a short period of time.

Hyphenation

311

8.5.3 Nucleic acids

Humans are exposed to a wide array of chemical compounds that can interact with nucleic acids. Interactions can lead to the formation of a chemical bond between such compounds and the DNA, leading to damaged DNA and so-called DNA adducts. The structural elucidation of these DNA adducts is an important research topic in cancer etiology, while their detection and quantitation is substantial for cancer prevention. The CEC analysis of two common DNA adducts, acetylaminofluorene de0xyguanosine and the anti-benzo[g]chrysene 11,12-dihydrodiol 13,14-epoxide deoxyguanosine adduct, is shown in Fig. 8.25. An on-column focusing method, i. e. injection at low solvent strength, was used in order to introduce picomole amounts of DNA adducts present at low concentration. The samples were dissolved at concentrations of 1.10 -6 M in 5 mmol/1 aqueous ammonium acetate-10% acetonitrile and electrokinetically injected under 12 kV for durations of 4-7 min. Elution of the adducts in a capillary column packed with octadecylsilica particles took place with 5 mmol/1 aqueous ammonium acetate-60% acetonitrile. The extracted ion chromatograms of m/z 489.5 and

c" (D z._ z._

0

m/z 596.5

20

I

I

25

I

I

30

time [min]

Fig. 8.25. CEC--ESI-MS analysis of the adducts between acetylaminofluorene or the dihydodiol epoxide of benzo[g]chrysene with deoxyguanosine. Column, 280 x 0.075 mm i.d. packed with 5 ~tm/100 A Nucleosil-C18; eluent, 5 mmol/1 ammonium acetate, 60% acetonitrile; applied voltage, 125 kV; detection, ESI-MS, 370-470 amu; sheath liquid, 2 mmol/1 aqueous ammonium acetate, 50% acetonitrile, 3 ~tl/min; injection, electrokinetic, 12 kV, 7 min. (Reprinted with permission from ref. [ 100]. Copyright [ 1997] American Chemical Society.). References pp. 313-316

312

Chapter 8

373.7

o HOHzC~

MH* o. 489.7 - ribose I

e" L_ L_

OH

MH* 569.5

e,o

/ 300

i

400

500

600

(m/z)

!

700

300

I

400

i(/+Na)* 500

600

700

(m/z)

Fig. 8.26. Mass spectra of the adducts between (a) acetylaminofluorene or the (b) dihydodiol epoxide of benzo[g]chrysene with deoxyguanosine. (Reprinted with permission from ref. [ 100]. Copyright [1997] American Chemical Society).

596.5 and the full-scan mass spectra of the two components are illustrated in Fig. 8.25 and Fig. 8.26. Acetylaminofluorene deoxyguanosine and the anti-benzo[g]chrysene 11,12-dihydrodiol 13,14-epoxide deoxyguanosine adduct were readiliy identified with molecular masses of 488.5 and 595.5, respectively. Recently, the principal applicability of CEC to the separation of nucleotides transfer-RNAs has been demonstrated using an octadecyl-sulfonated mixed mode silica stationary phase. Although UV absorbance was used for detection of the analytes, it can be expected that such a separation system will be coupled to ESI-MS in the near future. Presently, our laboratory is working on the development of monolithic capillary columns for ion-pair reversed-phase CEC--ESI-MS of nucleic acids. 8.6 ACKNOWLEDGMENTS This work was supported by grants from the Austrian Science Fund (P-13442PHY, P-14133-PHY).

Hyphenation

313

8.7 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

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316

Chapter 8

104 C. Rentel, P. Gfrorer and E. Bayer, Electrophoresis, 20 (1999) 2329. 105 C.J. Paterson, R.J. Boughtflower, D. Higton and E. Palmer, Chromatographia, 46 (1997) 599. 106 S.J. Lane and A. Pipe, Rapid Commun. Mass Spectrom., 12 (1998) 667. 107 V. Spikmans, S.J. Lane, U.R. Tjaden and J. Vandergreef, Rapid Commun. Mass Spectrom., 13 (1999) 141. 108 P. Huang, X. Jin, Y. Chen, J.R. Srinivasan and D.M. Lubman, Anal. Chem., 71 (1999) 1786. 109 C. Ericson and S. Hjerten, Anal. Chem., 71 (1999) 1621. 110 I. Gusev, X. Huang and C. Horvfith, J. Chromatogr. A, 855 (1999) 273. 111 C. Eicson, J. Holm, T. Ericson and S. Hjerten, Analy. Chem., 72 (2000) 81. 112 M.M. Dittmann and G.P. Rozing, J. Microcol. Sep., 9 (1997) 399. 113 Choudhary, G. Fundamentals of Capillary Electrochromatography and Its Coupling

with Mass Spectrometry, Dissertation I998, Yale University, New Haven, CT, USA. 114 115 116 117 118 119 120 121

J. Abian, A.J. Oosterkamp and E. Gelpi, J. Mass spectrom., 34 (1999) 244. L.W. Hershberger, J.B. Callis and G.D. Christian, Anal. Chem., 51 (1979) 1444. M. Kohler and J.A. Leary, Anal. Chem., 67 (1995) 3501. J. Cai and J.D. Henion, J. Chromatogr., 703 (1995) 667. C.G. Huber and A. Krajete, J. Chromatogr. A, 870 (2000) 413. E. Bayer, P. Gfrorer and C. Rentel, Angew. Chem., 38 (1999) 992. C.G. Huber and A. Premstaller, J. Chromatogr. A, 849 (1999) 161. C.G. Huber, A. Premstaller and G. Kleindienst, J. Chromatogr. A, 849 (1999) 175.

Chapter 9

Pressure Supported CEC: a High-Efficiency Technique for Enantiomer Separation Dorothee W I S T U B A * and Volker S C H U R I G

Institute of Organic Chemistry, University of Tiibingen, Auf der Morgenstelle 18, 72076 Tiibingen, Germany

CONTENTS

9.1 9.2

9.3

9.4 9.5

9.6

9.7 9.8 9.9

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Techniques of pressurizing . . . . . . . . . . . . . . . . . . . . . . . . . 9.2.1 Pressurization of the capillary from both sides . . . . . . . . . . 9.2.2 Pressurization of the capillary from one side . . . . . . . . . . . Theory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3.1 Efficiency and flow profile . . . . . . . . . . . . . . . . . . . . 9.3.2 Selectivity and retention . . . . . . . . . . . . . . . . . . . . . . Instrumentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.5.1 Chiral stationary phases . . . . . . . . . . . . . . . . . . . . . . 9.5.1.1 Cyclodextrin and cellulose derivatives . . . . . . . . . 9.5.1.2 'Brush-type' chiral stationary phase . . . . . . . . . . 9.5.1.3 Macrocyclic antibiotics . . . . . . . . . . . . . . . . . 9.5.1.4 Quinine-based anion-exchange type CSPs . . . . . . . 9.5.1.5 Chiral ligand-exchange CSPs . . . . . . . . . . . . . . 9.5.1.6 Polyacrylamide derivatives . . . . . . . . . . . . . . . 9.5.1.7 Molecularly imprinted polymers (MIPs) . . . . . . . . 9.5.2 Chiral buffer additives . . . . . . . . . . . . . . . . . . . . . . . Advantages and disadvantages of pressurized CEC . . . . . . . . . . . . 9.6.1 Comparison of pressurized CEC and normal CEC . . . . . . . . 9.6.2 Comparison of pressurized CEC and LC . . . . . . . . . . . . . Conclusion and future trends . . . . . . . . . . . . . . . . . . . . . . . . Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

318 319 319 319 321 321 323 325 326 326 327 330 330 331 333 333 333 335 335 335 336 336 337 337

318

Chapter 9

9.10 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

337

9.1 INTRODUCTION Capillary electrochromatography (CEC) combines the selectivity of chromatographic methods with the high efficiency of electromigration methods. Typical LC or GC stationary phases can be used as separation media, while the pressure driven flow in chromatography is replaced by the electroosmotic flow (EOF) and additionally by the electrophoretic mobility in the case of charged solutes. Thus, due to of its high efficiency, CEC represents an attractive alternative to chromatography in the field of chiral analysis. Enantiomer separation by CEC can be performed by three different methods: (i) wall-coated open tubular electrochromatography (o-CEC), in which the internal capillary wall is coated with the chiral stationary phase (CSP), (ii) packed capillary electrochromatography (p-CEC), in which capillaries are filled with typical chiral HPLC packing material and (iii) capillary electrochromatography with rods or monoliths (rod-CEC), in which capillaries with monolithic chiral stationary phases prepared by in situ polymerization methods within the capillaries are employed. Recent reviews summarized these methods ((i)-(iii)) and reported on a number of applications [ 1-4]. A drawback of p-CEC is the formation of air bubbles. This effect can lead to baseline noise or even a breakdown of the current. The reason for air bubble formation is primarily the differences in the EOF between the frit separating the packed and the unpacked part of the capillary [5-7]. Variations of the flow occur in the regions of frits which hold the packing bed in position. Another assumption is that the Joule heating promotes air bubble formation [8,9]. The particles of the packed bed act as nucleation sites for bubble formation. Once formed, a small bubble is immobilized by the particles and the heat generation in this region increases as a result of increased electrical resistance. A self-acceleration of the bubble takes place and results in a rapid dry out of a part of the packing bed. To remove formed air bubbles or to wet dry areas it is necessary to flush the capillary with the mobile phase at high pressure using an HPLC pump. Often a long flushing time is a necessity. The formation of bubbles can be prevented by pressurizing the flow system. Other means of suppressing of bubble formation are degassing of the eluent, working at low temperatures, the use of buffers which generate low currents (low-concentrated buffers or low-conductive buffers, such as zwitterionic buffers) and the use of capillaries with small inner diameters and highly permeable frits. In principle, separation of enantiomers by CEC can be performed with or without

Pressure Supported CEC

319

pressure support. While relatively few research groups practice p-CEC without pressure support, o-CEC and rod-CEC are commonly carried out without pressure support. 9.2 TECHNIQUES OF PRESSURIZING

Two successful approaches of pressurization are known: (i) Both the inlet and the outlet buffer vial are constantly pressurized [ 10-13]. (ii) Pressure can be applied either to the inlet [9,14-18] or the outlet end of the capillary [5,6]. This technique is called pressure supported or pressure assisted electrochromatography (other synonyms are: pressurized CEC, pressurized flow CEC), pressure electrochromatography (PEC), pseudoelectrochromatography (pEC) or electro-HPLC [9,14,15,19-21 ]). 9.2.1 Pressurization of the capillary from both sides

The pressurization of the flow system is carried out on both the inlet and the outlet side of the buffer reservoir with the same pressure as was first suggested by Knox and Grant [8]. Thus the analysis takes place at a higher pressure than the normal pressure. This reduces the risk of bubble formation in both the packed part and the unpacked part (where the detection window is situated) of the capillary. In this variant of pressurization, the mobile phase is only electro-driven. The advantage of this method is the high efficiency of the capillary electrophoresis caused by the flat flow profile. The majority of CEC separations that have been published were performed by this method. Commercially available instruments allow the pressurization of both ends of capillaries up to about 12 bar. 9.2.2 Pressurization of the capillary from one side

As first described by Tsuda [9,14], the flow system can be pressurized by coupling an HPLC pump to the inlet vial. The EOF is supplemented by a pressure-driven flow, so that this method is a hybrid form of pure CEC and capillary LC. The flat profile of the electro-driven system is overlaid with the parabolic flow profile of the pressuredriven method. Thus a decrease in efficiency of CEC proportional to the extent of pressure support can be observed. The advantages of this method are: (i) the possibility of switching between the CEC and the LC mode [18,19, 21-24], (ii) the feasibility of easily changing the mobile phase by flushing the capillary [18], (iii) the ease of interfacing to ESI-MS [15,16,25-30], (iv) shorter analysis times [17,18], (v) the possibility of generating a solvent gradient by means of the HPLC pump, [21-23], (vi) the minimizing of bubble formation, (vii) the feasibility to operate at low pH values with reasonable mobile flow velocities, and (viii) the variability of the applied pressure References pp. 337-339

320

Chapter 9 2,3.4

L 5

1 E t," 'T

N v o

.,

C I

3

,12 0

<

5

9

0

5

---

9

4

2 1

......

10

9 ,,

I--

15

_

i|

ill v

20

9

25

Jl,II

(mln)

Fig. 9.1. Separation of carboxylic acids and hydrophobic compounds with (a) LC and (b) pressure supported CEC ((1) folic acid, (2)p-hydroxybenzoic acid, (3) acetylsalicylic acid, (4) nicotinic acid, (5) thiourea and (6) nicotinamide). Capillary: 15 cm x 100 ~tm I.D. packed with Nucleosil 100 3-C18 (3 p,m). Conditions: disodium tetraborate (20 mM, pH 8.5)methanol, 25:75 (v/v); 63 bar at the inlet vial (a,b);-6 kV (b). Reproduced from [17], with permission. support up to 400 bar. For charged analytes, the combination of electro- and pressuredriven flow can lead to changes in selectivity compared to the purely pressure or electro-driven system [17,19]. For example, Eimer et al. [17] demonstrated the separation of carboxylic acids and hydrophilic compounds with both methods, LC and pressure supported CEC. As shown in Fig. 9.1, a quite different elution pattern was observed in the LC and in the pressure supported CEC mode. Furthermore, p-hydroxybenzoic acid (2), acetylsalicylic acid (3) and nicotinic acid (4) were not resolved with the pressure-driven method, while a baseline separation of all three compounds was realized by pressure supported CEC. The possibility of switching between the LC and the CEC mode enables six modes of operations in one instrumental set-up: isocratic and gradient CEC, isocratic and gradient Ia-LC, and isocratic and gradient pressure supported CEC (LC/CEC mixed

321

Pressure Supported CEC

rra~ t a~

2 1 13

4

1 -____ I[.~lJl / l, . . . . . . 1

,1 ot. .2o

[

/

80

. 110 b a t + 10 kV

LC m o d e 110 bar ~20 t ~L_ ....

~.,~ . . . . . ~s . . . . . .

?~

.... 40"

- - ~ ; s . . . . ~s" - - ~ ' r ~ o

" s.o .....

z2;s-

~ n

Fig. 9.2. Comparison of LC and pressure supported CEC at different voltages. Compounds: (1) thiourea, (2) dimethylphthalate, (3) diethylphthalate, (4) biphenyl and (5) o-terphenyl. Column: CEC Hypersil C18 3 ~tm, 25 (40) cm • 100 ~tm I.D. Conditions: Tris.HC1 (25 mM, pH 8)-acetonitrile-H20, 20:70:10 (v/v/v); 20~ 100 bar (at the inlet vial). Reproduced from [31 ], with permission.

mode) [31]. Fig. 9.2 demonstrate the influence of applied voltage on the separation of five compounds at constant pressure. With LC (110 bar), the retention time is about twofold longer than with pressure supported CEC (110 bar, 20 kV), while the efficiency is maintained (peak 5 has a plate number of 25 000 in all separations) [31 ]. An example for a separation by pressurized gradient electrochromatography was demonstrated by Behnke et al. [32]. Tryptic peptides of cyctochrome c were separated using a combination of inlet side pressurization with an HPLC pump and resistor capillary at the outlet side for avoiding air bubbles. As shown in Fig. 9.3, a significant improvement compared to the LC separation can be observed. Unresolved peaks in the LC run were now separated. An alternative technique to circumvent bubble formation is the pressurization of the outlet vial. Rebscher and Pyell reported that a counterpressure of 15 bar did not significantly influence the efficiency, but minimized the tendency to bubble formation [5,6].

9.3 THEORY 9.3.1 Efficiency and flow profile In packed capillaries, differences in the diameters of the channels between the silica particles and between the silica particles and the capillary wall lead to different References pp. 337-339

322

Chapter 9

0kV/m

E i-o v O t-

(b)

50 kV/m

O (/3 ..Q

[

i

i

5

!

"i"

"1

10

i

i

time (min)

Fig. 9.3. Tryptic maps of cyctochrome c by (a) nano-LC (250 bar) and (b) pressurized electrochromatography (12 kV, 250 bar (at the inlet vial)). Column: 6 cm (20 cm overall length) x 100 ~tm I.D. packed with ODS (1.5 ~m; 100 A); eluent: 0.1% v/v trifluoracetic acid in water (pH 2) containing (A) 0 % and (B) 50 % acetonitrile; gradient, 0-100 % B in 10 rain. Reproduced from [32], with permission.

flow velocity and thus to a parabolic flow profile in the case of a pressure-driven mobile phase (see Fig. 9.4). By way of, the electro-driven flow is largely independent of the diameter of the flow channels (except for particles smaller than 0.5 ~m where the electrical double layer overlaps within the extremely small channels) and a characteristic flat plug-like profile can be observed (see Fig. 9.4) [33]. The different flow profiles in LC and CEC are the reason for the greater band broadening and thus the lower efficiency of the pressure-driven method. The van Deemter equation describes the relationship between the height equivalent to one theoretical plate (H) and the linear flow velocity (u):

H=A+

B bl

+Cu

(9.1)

(A is the Eddy diffusion; B is the longitudinal diffusion; C is the mass transfer; u is the mobile-phase linear velocity) In LC, the Eddy diffusion arising from the differences in flow velocity in the packing bed channels is much larger than in CEC, with the result that the H of LC columns are always higher than those in CEC columns. In pressure supported CEC (as

323

Pressure Supported CEC

] Pressure drive m ~ part|cl~.~

"o'~ ~-/channel~~cw flow

velocityprofile

velocity

Electroosmotic drive 1~article '~ , r Ip ~ !> velocityprofile ~ /channel~~

-o~-n,

flowvelocity

,,

L_

Fig. 9.4. Flow velocity profiles vs. channel diameter for pressure driven flow (top) and electroosmotically driven flow (bottom). Reproduced from [33], with permission.

described in: 9.2.2 Pressurization of the capillary from one side), a combination of the parabolic and flat flow profile leads to an increase in H compared with pure CEC. With increasing pressure support the flow profile changes from flat to parabolic. Gfr6rer [34] demonstrated that the influence of 30 bar pressure support on the plate height is negligible both for slightly retained and for strongly retained analytes. The reduced plate height h increased from 1.50 to 1.57 for a slightly retained analyte (methylbenzoate, k = 0.4) and from 1.71 to 1.80 for a stronger retained analyte (butylbenzoate, k = 1.1) [34]. At higher pressure support (> 100 bar) the reduced plate height approaches to that observed in pure HLPC. Fig. 9.5 shows the van Deemter curves of butylbenzoat in the HPLC (CHPLC), pure CEC and pressure supported CEC mode.

9.3.2 Selectivityand retention While for uncharged analytes the selectivity in CEC and LC is identical, a different selectivity can be expected for charged analytes. Therefore, Tsuda et al. [9,14,17,35,36] introduced an electrochromatographic retention factor for charged analytes. In pressure supported CEC the retention time is dependent upon the electrophoretic mobility Uep, the electroendoosmotic velocity Ueo, the velocity of the pressurized flow Up and the chromatographic retardation. The velocity of the mobile phase Um is:

References pp. 337-339

324

Chapter 9

9

15-

CEC

9

CEC + 30 bar

9

CEC+60bar

9

CEC + 80 bar

9 CEC+10Obar

*

CHPLC

12.

§

~,R

6

9

9

Iiiiii. . . .

.. ,<

,1,,,!!! 0,0 0,2 0,4 0;6

,,,,,

1,2 1,4 1,6 1,8 2,0 2,2 2,4

u [mm/s]

Fig. 9.5. van Deemter curves of butylbenzoate in the HPLC, pressure supported CEC and the pure CEC mode. Reproduced from [34], with permission.

(9.2)

Urn = Ueo + Up

For unretained analyts the total linear velocity

bltot = blep -t- bleo + Up

Utot

is:

(9.3)

For retained analytes with a chromatographic retention factor k the linear flow velocity

Ux

is:

u~ = Uto, R -

Utot

l+k

(9.4)

325

Pressure Supported CEC

where R =

(l+k)

is the retardation factor or instead of eq (9.4) one can write:

(9.5)

Um

Ux=~ 1 +k*

where k* is the electrochromatographic retention factor. From (9.4) and (9.5) k* was derived:

k, =Um (1 + k)

(9.6)

- U,o,

Utot

The elution time of a retained analyte is derived from eq (9.4) where Ux = L/t.

t=--

L

(9.7)

R Utot

where L is the column length, and R is the retardation factor

(1 + k)"

9.4 I N S T R U M E N T A T I O N Electrophoretic systems for CEC which allow the pressurization of the separation capillary from both sides (see 9.2.1) up to 12 bar are commercially available. Hewlett Packard (HP) (now Agilent, Waldbronn, Germany) provides an option for pressurization up to 12 bar and Beckmann/Coulter (Fullerton; CA, USA) up to 8 bar. Also some home-made instruments are described in the literature. Electrophoretic systems with the capability of pressurizing one end of the capillary are generally realized by coupling an HPLC pump to the buffer inlet or outlet vial, respectively. These instruments are usually home-made [5,6,9,14,17] but two are also commercially available (Grom (Herrenberg, Germany); Prolab (Rheinach, Switzer-

References pp. 337-339

Chapter 9

326 Injector . . . . .

Filter'W--I. . . . .

[ . Pump

Column for analysis '

"I~

] B

[ "il

7

Resistant tube l

~) I

[__ J

i

i

i

J High voltage power supply J

.

Fig. 9.6. Instrumentation for pressure supported CEC. Reproduced from [35], with permission.

land)). Fig. 9.6 shows a home-made electrophoresis apparatus with a coupled HPLC pump, an injector with a split system and a UV-detector. The use of a gradient HPLC pump makes gradient CEC possible [21]. With such a system, analysis can be performed in the CE, CEC, pressure supported CEC, LC, gradient CEC or gradient LC mode. A recently published review described the instrumentation for CEC [37] 9.5 APPLICATIONS Enantiomer separation by p-CEC or rod-CEC can be performed in two ways: (i) the chiral selector is bound to the separation bed (silica particle or monolith) or (ii) the chiral selector is introduced as a buffer additive into the mobile phase of a capillary filled with an achiral separation medium.

9.5.1 Chiral stationary phases The first generation of research groups working in the field of enantiomer separation by CEC did not pressurize the flow system [38-41]. The result were long elution times associated with broad tailing peaks. Now, most of the recently published papers deal with electrochromatographic enantiomer separation under elevated pressure by pressurizing both ends of the capillary [48-52, 54,55,57,58,62]. In some cases pressurization of the inlet vial alone was described [42-45,59,60].

327

Pressure Supported CEC 9.5.1.1 Cyclodextrin and cellulose derivatives

Enantiomer separation of various compounds such as barbituric acids, benzoin, MTH-proline, glutethimide, et-methyl-et-phenyl-succinimide, 7-phenyl-7-butyrolactone, methyl-mandelate, 1-(2-naphthyl)ethanol, mecoprop methyl, diclofop methyl and fenoxaprop methyl by pressure supported CEC on a permethyl-13-cyclodextrin modified stationary phase was described by Wistuba and Schurig [42-44]. Three different separation beds were used: (i) permethyl-13-cyclodextrin was covalently attached via a thioether to silica (Chira-Dex-silica) [42], permethyl-[3-cyclodextrin was linked to a dimethylpolysiloxane and thermally immobilized (ii) on silica (ChirasilDex-silica) [43] or (iii) on a silica monolith (Chirasil-Dex-monolith) [44], respectively. An electrophoretic system coupled with an HPLC pump for pressurizing the inlet vial (10-15 bar) was used for all analysis. Beside the ability to circumvent bubble formation, this technique offers also the possibility of switching between the LC and

OH 3

0

0

20kV, 10bar

W

0

I

.N_

20 [min]

10

10bar only [

I

I

I

I

0

40

80

120

160

[min]

Fig. 9.7. Influence of pressure support on the enantiomer separation of mephobarbital on 100 ~m I.D. x 23.5 cm (overall length, 40 cm) capillary packed with Chira-Dex-silica. Conditions: phosphate buffer (5 mM, pH 5.0)-methanol (4:1, v/v); UV detection, 230 nm; upper chromatogram: 20 kV, 10 bar (at the inlet vial); lower chromatogram, 10 bar. Reproduced from [42], with permission. References pp. 337-339

Chapter 9

328

CH 3 I O,.. N. .,O ~ ~CH2CH3 p-CEC

p-LC

I

N, = 16 870 N2 = 19 415

N1= 5606

T" 0

10

20

[min]

0

10

20

[min]

Fig. 9.8. Comparison of pressure supported CEC (only the inlet vial is pressurized) and HPLC: enantiomer separation of mephobarbital. Conditions: 100 lam I.D. x 23.5 cm capillary (overall length, 40 cm) packed with Chira-Dex-silica; phosphate buffer (5 mM, pH 7.0)-methanol (4:1, v/v); UV detection, 230 nm; CEC, 20 kV; 10 bar; HPLC, 140 bar. Reproduced from [42], with permission.

CEC modes in a single instrumental set-up. Thus LC and CEC measurements can conveniently be performed with the same column. The influence of pressure support on the enantiomer separation of mephobarbital was investigated by comparing p-LC at 10 bar with p-CEC at 10 bar and 20 kV (see Fig. 9.7). With comparable chiral separation factors and resolutions, the elution time observed in the LC mode was longer than that in the CEC mode by a factor of approximately 10 for Chira-Dex-silica [42], 3 for Chirasil-Dex-silica [43] and 4 to 5 for Chirasil-Dex-monolith [44]. The elution time is thus clearly dominated by the substantial contribution of the EOF.

3 29

Pressure Supported CEC

O~

CH3 I N~ ~.0

~"~CH2CH3 HINO'~~ 20 kV no pressure support

20 kV 6 bar

20 kV 12 bar

() 10 :~0 min Fig. 9.9. Enantiomer separation of mephobarbital on a Chirasil-Dex-monolith by pressure supported CEC (only the inlet vial is pressurized) and pure CEC. First peak, dimethylformamide; second and third peak, enantiomers of mephobarbital. Top: without pressure support. Center: 6 bar pressure support. Bottom: 12 bar pressure support. Conditions: 20 cm (overall length 35 cm) x 100 ~m I.D. capillary; 20 kV; MES (20 mM, pH 6.0)-methanol (1:1, v/v). Reproduced from [44], with permission.

Comparing the two methods p-LC and p-CEC, the electrochromatographic method always shows higher theoretical plate numbers and resolutions at comparable elution times [39-41] (see Fig. 9.8). The Chirasil-Dex-monolith [44] consists of a single piece of a chiral modified porous solid, thus frits are no longer required and the risk of air bubble formation decreases. Pressurization is not insisted. As shown in Fig. 9.9 the enantiomer separation of mephobarbital on a Chirasil-Dex-monolith can be performed Referencespp. 337-339

Chapter 9

330

without pressure support. Yet, the pressure support leads in this case to a more stabile baseline and a reduction of the elution time of the first eluted enantiomer from 26.3 min (without pressure support) to 17.4 rain (12 bar pressure support). The theoretical plate number varied from 88 400 (without pressure support), over 65 000 (6 bar pressure support) to 50 600 (12 bar pressure support). Methanol or acetonitrile were used as organic modifiers, with the best enantioselectivity being observed with methanol. Cellulose tris(3,5-dimethylphenylcarbamate) or cellulose-tris(4-methylbenzoate) coated on silica has been used as a chiral stationary phase for enantiomer separation of lorazepam, c~-hydroxyethyl-naphthalene, benzoin, glutethimide and trans-stilbene oxide, ecomazole, 2,2'-diamino-6,6'-dimethylbiphenyl, glutethimide, piprozolin, etozolin, Tr6ger's base, indapamide, aminoglutethimide and metonidate by pressurized CEC whereat the pressure is applied at the inlet and the outlet buffer vial [46,61 ] and of indapamine, whereat only the inlet vial is pressurized [45]. Enantiomer separation of thalidomide and its hydroxylated metabolites was performed on a mixed phase consisting of cellulose-tris(3,5-dimethylphenylcarbamate) and amylose-tris(3,5-dimethylphenylcarbamte) under non-aqueous conditions [47].

9.5.1.2 'Brush-type' chiral stationary phases Wolf et al. found excellent resolution for the enantiomer separation by p-CEC of more than 30 neutral compounds on (S)-naproxen-derived and (3R,4S)-Whelk-O selectors immobilized on silica [48,49] (see Fig. 9.10). Generally, short analysis time combined with high selectivity and high efficiency were observed, and resolution Rs of up to 48 being achieved. Acetonitrile and methanol were used as organic modifiers to the zwitterionic buffer and the enantiomer separation was performed by pressurizing both the inlet and outlet vial during the electrochromatographic run. Peters et al. reported on rod-CEC on a chiral monolith [50] which was prepared by copolymerization of the chiral monomer 2-hydroxyethyl methacrylate (N-L-valine3,5-dimethylanilide)

carbamate with

ethylene dimethylacrylate, 2-acrylamido-

2-methyl-1-propanesulfonic acid and butyl or glycidyl methacrylate in the presence of a porogenic solvent. The electrochromatographic enantiomer separation of N-(3,5dinitrobenzoyl)leucine diallylamide was feasible at 25 kV; the inlet and outlet buffer vials were both pressurized.

9.5.1.3 Macrocyclic antibiotics The macrocyclic antibiotics vancomycin [51,52] and teicoplanin [53] were successfully employed as chiral selectors for enantiomer separation in p-CEC. Warfarin,

Pressure Supported CEC

331

0

! 1

,~11 0

2

4

6

8

Fig. 9.10. Enantiomer separation on a 29.5 cm (overall length, 38 cm) capillary packed with (3R,4S)-Whelk-O-stationary phase by CEC under slight overpressure (10 bar). Conditions: MES (25 mM, pH 6.0)-acetonitrile (1:3.5, v/v); 25 kV; UV detection, 230 nm. Reproduced from [48], with permission. hexobarbital [51], thalidomide (see Fig. 9.11)and a number of basic compounds including []-adrenergetic blocking agents [52] were separated on a vancomycin chiral stationary phase with resolutions of up to 2.7. Using a teicoplanin stationary phase, the enantiomer separation of tryptophan and dinitrobenzoyl leucine was feasible. Short elution times (less than 6 min) and resolutions of up to 2.39 were found. [53]. Analysis were performed with an electrophoresis system which allows the pressurization of both ends of the capillary up to 12 bar.

9. 5.1.4 Quinine-based anion-exchange type CSPs L~immerhofer and Lindner reported on the enantiomer separation of derivatized amino acids and profens on a weak-anion-exchange(WAX)-type stationary phase based on chiral quinine carbamate selectors by p-CEC [54,55]. The separations were performed either under aqueous or [54] non-aqueous conditions [55]. The efficiency obtained in the p-CEC mode was about two to three time higher than with LC using an acetonitrile/buffer flow system [54]. Very high resolutions and efficiencies were found for non-aqueous p-CEC. For example, the enantiomer separation of Fmocleucine was achieved in less than 10 rain with a resolution Rs of 6.9 at about 100 000

References pp. 337-339

332

mAU

Chapter 9

120100-

80604020O= 0

5

10

Min Fig. 9.11. Enantiomer separation of thalidomide on a vancomycin chiral stationary phase by CEC under a slight overpressure (10 bar). Conditions: 75 gm x 35.5 cm capillary; mobile phase, methanol-acetonitrile-glacial acetic acid-triethylamine (80:20:2:0.2, v/v/v/v); 20 kV, UV detection, 220 nm. Reproduced from [52], with permission.

mAU 17.5 15

(R) I

N (R) = 18 219 N (S) = 16 085 u = 0.16 mmls

I

I

(s) I I I I

12.5 10

-~~

7.5

~

5 2.5 0

-2.5 o ..........

~i0

-

-

20

. . . .

:~om~.

Fig. 9.12. Enantiomer separation of Fmoc-Leu on a quinine carbamate-based WAX-type CSPs by CEC under a slight overpressure (8 bar). Conditions: capillary, 100 ~tm I.D. • 25 cm (overall length, 33.5 cm); acetonitrile-methanol (80:20, v/v); 200 mM acetic acid, 10 mM triethylamine;-25 kV; UVdetection, 254 nm. Reproduced from [58], with permission.

Pressure Supported CEC

333

theoretical plates per meter [55] (see Fig. 9.12). All electrochromatographic runs were performed by pressurizing the inlet and the outlet vial of the electrophoresis system with 8 bar [54,55]. 9.5.1.5 Chiral ligand-exchange CSPs

Schmid et al. [60] demonstrated the enantiomer separation of underivatized amino acids on a monolithic chiral ligand-exchange phase by rod-CEC. The chiral stationary phase was prepared in situ in the capillary by polymerization of methacrylic acid, piperazine diacrylamide, vinylsulfonic acid and N-(2-hydroxy-3-alloxypropyl)-L-4hydroxyproline. The monolithic separation bed was covalently linked to the intemal capillary wall and thus no frits were required. Fig. 9.13 shows the enantiomer separation of phenylalanine by (A) pure CEC (30 kV), (B) nano-LC (12 bar) and (C) pressure supported CEC (30 kV, 12 bar at the inlet vial). The shortest elution time was clearly obtained by pressure supported CEC, while the highest resolution was found in the pure CEC mode (CEC: Rs = 2.11; nano-LC: Rs = 0.98; pressure supported CEC: Rs = 1.60). 9. 5.1.6 Polyacrylamide derivatives

Capillaries packed with poly-N-acryloyl-L-phenylalanine ethyl ester (Chiraspher) modified silica were used for electrochromatographic enantiomer separation of bendroflumethiazide. To suppress bubble formation, the inlet buffer vial was pressurized to 12 bar and the outlet buffer vial to 4 bar [42]. Acetonitrile or methanol were used as organic modifier whereby acetonitrile was superior to methanol. Non-aqueous p-CEC was performed on helical poly(diphenyl-2-pyridylmethylmethacrylate) coated on wide-pore aminopropyl silica [56]. With this chiral stationary phase, the enantiomer separation of Tr6gers base, benzoin acetate, methylbenzoin and trans-stilbene oxide was achieved by pressure-supported CEC, applying a higher pressure to the inlet than to the outlet buffer vial. 9.5.1.7 Molecularly imprinted polymers (MIPs)

MIPs used as chiral stationary phases in o-CEC, p-CEC as well as in rod-CEC have shown high selectivity but relatively low efficiency. Most of the reported enantiomer separations on these phases were performed without pressurization of the flow system. Only Schweitz et al. described on the enantiomer separation of propranolol and metoprolol (print molecule: R-propranolol or S-metoprolol) [57] and ropivacaine, mepivacaine and bupivacaine (print molecule: S-ropivacaine) [58] by

References pp. 337-339

Chapter 9

334

< E E rO3 O4

18.49

A

40-

k'1=10.56 k'2=12.11

30-

20.98

20-

0(=1.14 Rs=2.11

10-

0 0

1'0

310

2'0

0

time(min)

40-~ 20.96 k'1=10.03 k'2=10.75

< 30 E ,,..,, E E r r u} E on

22.32

20

0( =1.08 Rs=0.98

10

I

10

0

I

20

3=0

4=0

3=0

;0

time(min) 40-

< E E ro3 eu

C

5.87

k'1=5.52 k'2=6.28

30-

6.55

20-

0(=1.14 Rs=1.60

E o u~

I0-

0

0

1~0

210 time(min)

Fig. 9.13. Enantiomer separation of phenylalanine by (A) pure CEC, (B) nano-LC and (C) pressure supported CEC (pressurization of the inlet vial). Conditions: sodium dihydrogenphosphate (50 mM, pH 4.6), Cu(II) (0.1 mM); 26 cm x 75 lam I.D. capillary; (A) 30 kV; (B) 12 bar; (C) 30 kV and 12 bar. Reproduced from [60], with permission.

Pressure Supported C E C

33 5

rod-CEC under an overpressure of 7 bar. 9.5.2 Chiral buffer additives

In CE a successful method for enantiomer separation is the addition of a chiral selector to the mobile phase. This practice can be transferred to p-CEC by using a packing bed which consists of bare silica or ODS (octadecylsilica) and a mobile phase containing a chiral additive. At the first time, Leli6vre et al. added hydroxypropyl-[3cyclodextrin to the mobile phase using an ODS packed capillary. The enantiomer separation of chlorthalidone with a resolution Rs of 1.4 was feasible [41 ]. Deng et al. [59] used an ODS-packed column and [3-cyclodextrin as a mobile phase additive. A theoretical model for the enantiomer separation of salsolinol was developed and compared with the experimental data. For pressure supported CEC, very high pressure (about 100 bar) was applied to the inlet vial so that the mobile phase was mainly driven by the applied pressure. Lfimmerhofer and Lindner [62] reported on the enantiomer separation of derivatized amino acids on an ODS-packed capillary with a chiral quinine-derived selector as buffer additive in two different modes: (i) in an electrophoretically dominated mode at high electrolyte concentration and (ii) in an electroosmotically dominated mode at a low electrolyte concentration. Enantiomer separation in the electrophoretically dominated mode (i) leads to high efficieny (about two to three times higher than in LC) but to a moderate enantioselectivity (about the same as in LC). In the electroosmotically dominated mode (ii) a higher enantioselectivity but a lower efficiency (even inferior to LC) occurs. The separations can also been performed in a non-aqueous buffered mobile phase. Pressurization (8-10 bar) of the flow system on both ends of the separation capillary was applied. 9.6 ADVANTAGES AND DISADVANTAGES OF PRESSURIZED CEC 9.6.1 Comparison of CEC and pressurized CEC

Pressurized CEC has many advantages, but also suffers from some disadvantages. An important advantages of pressurization of both ends of the capillary with the same pressure is the suppression of air bubble formation. This electrochromatographic method, carried out under low overpressure, leads to a characteristic high efficiency, a stable baseline and a constant current. Advantages of pressure supported CEC with only the inlet buffer vial pressurized over normal CEC are the shorter analysis time, a more stable baseline, no bubble formation, a higher variability in selecting the optimum separation conditions, the

References pp. 337-339

336

Chapter 9

possibility of overcompensating for the electrophoretic mobility of charged analytes which is sometimes opposite to the direction of the EOF. Furthermore, the use of gradient CEC is possible and CEC-MS coupling is easily feasible. A disadvantage is a lower efficiency caused by the additional pressure-driven flow. Most instruments used are home-made and, with two exceptions, not commercially available. Automatization has not yet been achieved. Normal CEC has the advantage of high efficiency at the expense of long elution times. Moreover, baseline instability, bubble formation and fluctuation of the current is commonly observed.

9.6.2 Comparison of pressurized CEC and LC The efficiencies obtained with pressure supported CEC whereat pressurization takes place at one end of the capillary are always higher than those observed with LC. Another advantage is the shorter elution time resulting from the combination of pressure-driven and electro-driven flow. An advantage of the LC method is the availability of a diversity of commercially available instruments which are simple and robust. Enantiomer separation on a preparative scale is easier accomplished with LC than with CEC. 9.7 CONCLUSION AND FUTURE TRENDS The method of pressure supported CEC (pressurization of the inlet vial) is a hybrid method of normal CEC and LC, and the chromatographic parameters lie between those of LC and CEC. Depending on the extent of pressure support the above mentioned method tends either towards pure CEC or LC. Usually pressures of only up to 15 bar are applied and under such conditions efficiency is scarcely affected while the elution time is clearly reduced. If the CEC analysis is performed with slight overpressure by equal pressurization of both ends of the capillary the pressure-driven flow is absent. This technique serves only to suppress bubble formation. In the future the development of multifunctional systems allowing the switching between CE, CEC, pressure supported CEC and LC modes can be expected and tailor made conditions for each separation can be selected. The breakthrough of pressure supported CEC for enantiomer separation awaits adequate electrophoresis systems to become commercially available.

Pressure Supported CEC

337

9.8 A B B R E V I A T I O N S

CE CEC o-CEC p-CEC rod-CEC EOF PEC pEC ESI-MS CSP ODS WAX

capillary electrophoresis capillary electrochromatography capillary electrochromatography using an open-tubular capillary capillary electrochromatography using a packed capillary capillary electrochromatography using a monolithic (rod) capillary electroosmotic flow pressure electrochromatography pseudoelectrochromatography electrospray ionization mass spectrometry chiral stationary phase octadecylsilica weak anion exchange

9.9 A C K N O W L E D G E M E N T

The authors gratefully acknowledge Graeme J. Nicholson for helpful device in preparing the manuscript. 9.10 R E F E R E N C E S

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

V. Schurig, D. Wistuba, Electrophoresis, 20 (1999) 2321. A. Dermaux, P. Sandra, Electrophoresis, 20 (1999) 3027. D. Wistuba, V. Schurig, J. Chromatogr. A, 875 (2000) 255. G. Gt~bitz, M.G. Schmid, Enantiomer, 5 (2000) 5. H. Rebscher, U. Pyell, Chromatographia, 38 (1994) 737. H. Rebscher, U. Pyell, Chromatographia, 42 (1996) 171. M.T. Dulay, C. Yan, D.J. Rakestraw, R.N. Zare, J. Chromatogr. A, 725 (1996) 361. J.H. Knox, I.H. Grant, Chromatographia, 32 (1991) 317. T. Tsuda, Anal. Chem., 59 (1987) 521. N.W. Smith, M.C. Evans, Chromatographia, 38 (1994) 649. N.W. Smith, M.C. Evans, Chromatographia, 41 (1995) 197. R.J. Boughtflower, T. Underwood, C.J. Paterson, Chromatographia, 40 (1995) 329. R.J. Boughtflower, T. Underwood, J. Maddin, Chromatographia, 41 (1995) 398. T. Tsuda, Anal. Chem., 60 (1988) 1677. E.R. Verheij, U.R. Tjaden, W.M.A. Niessen, J van der Greef, J. Chromatogr., 554 (1991) 339. M. Hugener, A.P. Tinke, W.M.A. Niessen, U.R. Tjaden, J. van der Greef, J. Chromatogr., 647 (1993) 375. T. Eimer, K.K. Unger, T. Tsuda, Fresenius J. Anal. Chem., 352 (1995) 649. B. Behnke, E. Grom, E. Bayer, J. Chromatogr. A, 716 (1995) 207. T. Eimer, K.K. Unger, Trends Anal. Chem., 15 (1996) 463. D.A. Stead, R.G. Reid, R.B. Taylor, J. Chromatogr. 798 (1998) 259. B. Behnke, E. Bayer, J. Chromatogr. A, 680 (1994) 93.

33 8

Chapter 9

22 A. Apffel, H. Yin, W.S. Hancock, D. McManigill, J. Frenz, S.-L. Wu, J. Chromatogr. A, 832 (1999) 149. 23 J.N. Alexander, J.B. Poli, K.E. Markides, Anal. Chem., 71 (1999) 2398. 24 Y. Zhang, W. Shi, L. Zhang, H. Zou, J. Chromatogr. A, 802 (1998) 59. 25 D.B. Gorden, G.A. Lord, Rapid Commun. Mass Spectrom., 8 (1994) 544. 26 K. Schmeer, B. Behnke, E. Bayer, Anal. Chem., 67 (1995) 3656. 27 S.E.G. Dekkers, U.R. Tjaden, J. van der Greef, J. Chromatogr. A, 721 (1995) 201. 28 M. Taylor, P. Teale, J. Chromatogr. A, 768 (1997) 89. 29 J.-T. Wu, P. Huang, M.X. Li, D.M. Lubman, Anal. Chem., 69 (1997) 2908. 30 P. Huang, J.-T. Wu, D.M. Lubman, Anal. Chem., 70 (1998) 3003. 31 M.M. Dittmann, G.P. Rozing, G. Ross, T. Adam, K.K. Unger, J. Cap. Elec., 4 (1997) 201. 32 B. Behnke, J.W. Metzger, Electrophoresis, 20 (1999) 80. 33 M. Dittmann, G. Rozing, J. Chromatogr. A, 744 (1996) 63. 34 P. Gfr6rer, Dissertation, Tt~bingen,1999. 35 S. Kitagawa, T. Tsuda, J. Microcolumn Sep., 6 (1994) 91. 36 S. Kitagawa, A. Tsuji, H. Watanabe, M. Nakashima, T. Tsuda, J. Microcolumn Sep., (1997) 347. 37 F. Steiner, B. Scherer, J. Chromatogr. A, 887 (2000) 55. 38 S. Li, K.D. Lloyd, Anal. Chem., 65 (1993) 3684. 39 D.K. Lloyd, S. Li, P. Ryan, J. Chromatogr. A, 694 (1995) 285. 40 D.K. Lloyd, S. Lee, J. Chromatogr. A, 666 (1994) 321. 41 F. Leli6vre, C. Yang, R.N. Zare, P. Gareil, J. Chromatogr. A, 723 (1996) 145. 42 D. Wistuba, H. Czesla, M. Roeder, V. Schurig, J. Chromatogr. A, 815 (1998) 183. 43 D. Wistuba, V. Schurig, Electrophoresis, 20 (1999) 2779. 44 D. Wistuba, V. Schurig, Electrophoresis, 21 (2000) 3152. 45 K. Krause, M. Girod, B. Chankvetadze, G. Blaschke, J. Chromatogr. A, 837 (1999) 51. 46 S. Mayer, X. Briand, E. Francotte, J. Chromatogr. A, 875 (2000) 331. 47 M. Meyering, B. Chankvetadze, G. Blaschke, J. Chromatogr. A, 876 (2000) 157. 48 C. Wolf, P.L. Spence, W.H. Pirkle, E.M. Derrico, D.M. Cavender, G.P. Rozing, J. Chromatogr. A, 782 (1997) 175. 49 C. Wolf, P.L. Spence, W.H. Pirkle, M. Deniz, D.M. Cavender, E.M. Derrico, Electrophoresis, 21 (2000) 917. 50 E.C. Peters, K. Lewandowski, M. Petro, F. Svec, J.M.J. Fr6chet, Anal. Commun., 35 (1998) 83. 51 A. Dermaux, F. Lynen, P. Sandra, J. High Resol. Chromatogr., 21 (1998) 575. 52 H. Wikstr6m, L.A. Svensson, A. Torstensson, P.K. Owens, J. Chromatogr. A, 869 (2000) 395. 53 A.S. Carter-Finch, N.W. Smith, J. Chromatogr. A, 848 (1999) 375. 54 M. L~immerhofer, W. Lindner, J. Chromatogr. A, 829 (1998) 115. 55 E. Tobler, M. L~immerhofer, W. Lindner, J. Chromatogr. A, 875 (2000) 341. 56 K. Krause, B. Chankvetadze, Y. Okamoto, G. Blaschke, Electrophoresis, 20 (1999) 2772. 57 L. Schweitz, L.I. Andersson, S. Nilsson, Anal. Chem., 69 (1997) 1179. 58 L. Schweitz, L.I. Andersson, S. Nilsson, J. Chromatogr. A, 792 (1997) 401.

Pressure Supported CEC

33 9

59 Y. Deng, J. Zhang, T. Tsuda, P.H. Yu, A.A. Boulton, R.M. Cassidy, Anal. Chem., 70 (1998) 4586. 60 M.G. Schmid, N. Grobuschek, C. Tuscher, G. Gt~bitz, A. V6gv~ri, E. Machtejevas, A. Maru~ka, S. Hjerten, Electrophoresis, 21 (2000) 3141. 61 M. Girod, B. Chankvetadze, G. Blaschke, J. Chromatogr. A, 887 (2000) 439. 62 M. L~immerhofer, W. Lindner, J. Chromatogr. A, 839 (1999) 167.

This Page Intentionally Left Blank

Chapter 10

Applications Zden6k DEYL and Ivan M I K S [ K

Institute of Physiology, Academy of Sciences of the Czech Republic, Videhskti 1083, 142 20 Prague 4, Czech Republic

CONTENTS

10.1 10.2 10.3 10.4 10.5 10.6 10.7 10.8 10.9 10.10 10.11 10.12 10.13 10.14 10.15 10.16

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preconcentration procedures . . . . . . . . . . . . . . . . . . . . . . . Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ketones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Carbohydrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fatty acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Triglycerides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Steroids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Amino acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nucleic acids constituents . . . . . . . . . . . . . . . . . . . . . . . . . Drugs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Antibiotics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pesticides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Available applications (summarizing Table) . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

341 342 343 347 348 350 352 355 358 359 362 363 368 369 369 413

10.1 I N T R O D U C T I O N

Perusal of the reviews on capillary electrochromatography [1] and the recently published book on this subject by Krull et al. [2], as well as the special volume of J. Chromatogr. edited by Horv~ith [3], indicates clearly the applicability of this separation technique to a broad range of analytes, which differ considerably in their chemical nature and physical properties. It can be applied equally to low- and high-

342

Chapter 10

molecular-mass entities, or to analytes differing in polarity, or charge, or both. Particularly within the last four to five years the technique has matured in the sense that not only separations of artificial mixtures have been achieved, but the applicability to real samples (which is the ultimate goal) has been proven beyond doubt. Since the introduction of micellar (microemulsion) electrokinetic chromatography by Terabe and his co-workers [4] and the rather limited attempts at non-aqueous electrokinetic separations [5], capillary electrochromatography represents a real breakthrough in this area, because different partition mechanisms can be exploited on-line, offering better resolutions and shorter running times. In general, the different chromatographic and electromigration techniques tend to form a unified system that, by exploiting different partition mechanisms and separation modes, gives the analyst a broad spectrum of tools which, if properly applied, can bring a new quality into separation science. As capillary electrochromatography is the youngest of these techniques, little is known about its chemometric aspects. Because the applications of capillary electrochromatography are increasing rapidly, it is very difficult to give a detailed description of the different variations used, even for a definite set of compounds coming for analysis. Therefore, we have summarized the currently available applications in a Table 10.1, which provides the basic information and references to the original literature. However, the individual categories of compounds are given a brief commentary to highlight the most frequent approaches. Separations of model mixtures and of naturally occurring samples are included in the individual sections of the summarizing Table, as the transfer of separation conditions and modes can be made easily from one to the other. 10.2 PRECONCENTRATION PROCEDURES Removal of interfering matrix contaminants and/or improving the detection limits represent frequent problems in the analysis of natural, and particularly biological, samples. This potential has not yet been exploited fully in capillary electrochromatography. A typical example that demonstrates a combination of biorecognition-based separation with a subsequent capillary electrophoresis step is presented [ 180]. The technique has been named immunoaffinity electrochromatography (IACEC) [180]. In principle, the solute diluted in a large volume is applied to a capillary column packed with a support containing anti-solute IgG. The solute is selectively extracted by the immunoaffinity column, and the bound analyte is eluted and separated further (if desired) by capillary electrophoresis. In a particular case, fluorescein-labelled biotin was used as the solute to be accumulated and anti-biotin -IgG support was the immunoaffinity sorbent: the running buffer used was 5 mM Tris pH 8 at 10 kV (the currents resulting from this arrange-

Applications

343

ment are rather low, at 5-10 ~tA, creating an endo-osmotic flow of 0.5-1 mm/s in a 30-36 cm long packed bed. Samples were injected electrokinetically at 10 kV (the injection times were appropriately large, at 10-600 s). After the sample was applied, the immunoaffinity column was flushed with the running-buffer for 10-20 min to remove the matrix components and contaminants. Desorption of the bound analyte was done by a 20-30 s injection of the desorbing buffer (4 mM tetraborate, pH 9.2-propanol, 80:20 v/v). Next, the applied voltage was held at 0 V for a period of 40-270 s before the running buffer was applied at 10 kV. Desorbing injections were continued until two consecutive injections resulted in insignificant elution of the accumulated solute (FITC labelled biotin). Detection was done by laser-induced fluorescence and the immunoaffinity sorbent was based on diol-bonded silica. 10.3 HYDROCARBONS

Most frequently, the capillary electrochromatography system is equipped with a UV detector, so it is not surprising that - - to the best of our knowledge - - no paper dealing with the separation of aliphatic hydrocarbons has been published so far. On the other hand, quite a few reports deal with the separation of polyaromatic species. Nearly always the packing used is ODS and most of the separations are run isocratically. A typical application of CEC for the separation of polyaromatic hydrocarbons was published by Xin and Lee [9]. In this, a 35/43 cm x 50 ~tm i.d. fused silica capillary packed with 3 ~tm Spherisorb ODS-1 particles or, alternatively, 41/53 cm • 75 ~tm i.d. capillaries packed with 5 mm Spherisorb ODS-1, were used at 30 kV with UV detection (at 230 nm). The sample was introduced electrokinetically. Acetonitrile-Tris buffer (50 mM, pH 8.1) (80:20 v/v) was used as mobile phase. The results are seen in Fig. 10.1. Another technique reported for polyaromatic hydrocarbons separation uses ODSfilled (with frits) or entrapped (without frits) columns with 80% acetonitrile-20% 0.1 M acetate buffer (v/v) at pH 3 (30 kV per 17-25 cm - long capillary, i.d. 100 ~tm) [ 10]. The results are seen in Fig. 10.2. The technique of using entrapped sorbents can also be used for materials other than ODS (typically, imprinted polymers for chiral separations). Another application [12] uses a packed capillary segment combined with a bare silica section that serves to speed up the endo-osmotic flow, while the ODS-packed part of the capillary was used as the separation part of the capillary column. The detector's window was placed in-between the two segments. Monobasic sodium phosphate, 1.25 mM, pH 6.0, containing 75% v/v acetonitrile was used as background electrolyte at 625 V/cm, and 20 cm of the ODS-packed bed was used with the result References pp. 413-419

Chapter 10

344

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Fig. 10.1. Separation of polycyclic aromatic hydrocarbons (PAHs) on columns packed with Spherisorb ODS particles. Conditions: (A) 35(43) cm x 50 gm i.d. fused silica capillary column packed with 3 gm Spherisorb ODS-1 particles; (B) 41(53) cm x 75 gm i.d. fused-silica capillary column packed with 5 [am Spherisorb ODS-1 particles; 30 kV applied voltage; 5 kV, 5 s electrokinetic injection; acetonitrile-50 mM Tris buffer, pH 8.1 (80:20 v/v). Peak identifications: 1, benzene; 2, naphthalene; 3, acenaphthylene; 4, fluorene; 5, acenaphthene; 6, phenanthrene; 7, anthracene; 8, fluoranthene; 9, pyrene; 10, benz[a]anthracene; 11, chrysene; 12, benzo[b]fluoranthene; 13, benzo[k]fluoranthene; 14, benzo [a]pyrene; 15, dibenz[a, h]anthracene; 16, indeno[1,2,3-cd]pyrene; 17, benzo[ghi]perylene. Reproduced with permission from Xin and Lee [9].

seen in Fig. 10.3. The important point of these experiments is not so much in the acceleration of the analysis but the observation that, in capillary-electrochromatography-columns containing retaining frits, the flow of the mobile phase is not only based on electro-osmosis but is also contaminated by Poiseuille flow. Polyaromatic hydrocarbons and their nitro derivatives can be separated by capillary electrochromatography, as described in ref. [181]. Besides C18 phases, C8

345

Applications 6O

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20

Fig. 10.2. Separation of a mixture of PAHs on reversed-phase capillaries: (a) without and (b) with silicate entrapment. Conditions: 75 ~tm i.d. fused-silica capillary packed with 5 ~tm Nucleosil ODS particles; column effective lengths 25 cm for the non-entrapped column and 17 cm for the entrapped column. Both electrochromatograms were obtained under the same conditions: mobile phase, acetonitrile-0.1 M acetate buffer, pH 3.0, 80:10 (v/v); applied voltage 30 kV; UV detection at 254 nm; 20~ pressure 9 bar applied to both vials; electrokinetic injection, 10 kV for 10s. Reproduced with permission from Chirica and Remcho [ 10].

sorbents have also been used for the separation of alkylbenzenes [182]. The same purpose was served by monolithic beds [21,22], prepared by Hjerten and his co-workers, in which dextran sulfate, or sulfonic acids, formed a part of the bed that was supposed to create endo-osmotic flow while C18 and C4 moieties formed the hydrophobic partition site. Experiments were done with the presence of SDS both in supraand sub-micellar concentration, showing that the addition of SDS to a buffer-acetonitrile gradient results in a considerable shortening of the run time. Other monolithic systems were described by Peters et al. [23,24,183]. Another system based on the octadecylsilane moiety is octadecylated cellulose, described by Maruska and Pyell ([34,184]) as giving good results. C 18 Reversed phases are not necessarily limited to polycyclic and halogenated (or nitrated) hydrocarbons, but can be used for a number of oxygen-, nitrogen- and sulphur- containing heterocyclic compounds in a similar way to that used in reversedphase chromatography. In this context, ref. 185 describes the separation of compounds of this category. A plethora of similar separations can be found in the Section devoted

References pp. 413-419

Chapter 10

346

a, Nay

u = 0.8 -- I 0 1 , 0 0 0

mm/s plates/m

E

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!l

Cw

~f

u - 1.1 mm/s Nav= 130,000 plates/m

io

2~o

Min

Fig. 10.3. Electrochromatograms exhibiting the effect of the length of the bare-silica segment on the separation of benzene and alkylbenzenes. Capillary, 100 ~m i.d. with a 20 cm ODS segment and a variable-length bare-silica segment of; (a), 0 cm; (b), 6 cm and (c), 28 cm. Mobile phase, 1.25 mM sodium dihydrogenphosphate, pH 6.0, containing 75% (v/v) acetonitrile. Applied field strength, 625 V/cm. Detection, UV at 254 rim. Solutes: 1, benzene; 2, toluene; 3, ethylbenzene; 4, propylbenzene; 5, butylbenzene; 6, pentylbenzene. Reproduced with permission from Yang and E1 Rassi [12].

to drugs (see Section 10.12) or in the analysis of environmental pollutants (pesticides, insecticides, and softeners [ 19]). Two categories of capillary electromigration separations deserve further attention. The first group is separations that make use of monolithic packings and the other uses the so-called open-tubular-column separations. For hydrocarbon separations, fluorinated

columns

(exploiting

tridecafluoro-l,l,2,2-tetrahydro-octyl-l-triethoxysilane

(F 13-TEOS) and C8 ligands coupled to the inner side of the capillary wall have been used [ 186,187].

347

Applications

10.4 KETONES

Ketones (acetone, acetophenone and butyrophenone) can be separated successfully in 50 mM Tris-HCl buffer, pH 7.3, using capillaries coated with hydrophobic and charged groups containing polymers (Fig. 10.4) [11 ]. In this case, the partition occurs between the moving mobile phase (background electrolyte) and the surface-modified capillary. The apolar regions of the surface coating functioned as a reversed stationary phase while the polar domains (open tubular capillary electrochromatography mode) of the coating kept the endo-osmotic flow sufficiently large. This principle was also applied for the separation of five Parabens shown in Fig. 10.5, and polynuclear aromatic hydrocarbons, as shown in Fig. 10.6.

2

I

0.5mAU

I

0

,,,

I

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

1

I

2

,

4

,

,

.

,a

6

....

_ !

8

Time / min

Fig. 10.4. Separation of three types of ketones in 50 mM Tris-HC1 buffer, pH 7.3 as the mobile phase with the poly(TBAAm-co-AMPS)-coated column. Conditions: column, 750 mm x 25 ~tm i.d. (600 mm effective length); mobile phase, 50 mM Yris-HC1 buffer, pH 7.3; field strength, 400 V/cm; injection, 12 kV for 1 s at the side of the anode; detection wavelength, 254 nm. Peak identification: 1, acetone; 2, acetophenone; 3, butyrophenone. Reproduced with permission from Sawada and Jinno [11 ].

References pp. 413-419

Chapter 10

348

/2

I

0.5 m A U

I 0

5

,

I

,

4

i 8

Time

/ min

Fig. 10.5. Separation of parabens with the poly(TBAAm-co-AMPS)-coated column. Conditions: column, 750 mm x 25 pm i.d. (600 mm effective length); mobile phase, 20% acetonitrile (v/v) in 50 mM Tris-HC1 buffer; field strength, 400 V/cm; injection, 12 kV for 3 s at the side of the anode; detection wavelength, 254 nm. Peak identification: 1, methylparaben; 2, ethylparaben; 3, propylparaben; 4, butylparaben; 5, amylparaben. Reproduced with permission from Sawada and Jinno [11 ].

10.5 CARBOHYDRATES

Much work has been published about the separation of carbohydrates (and peptides) [36,188]. The separations originating from Novotny's group [36] almost invariably use polyacrylamide-polyethylene glycol monolithic copolymer packings with added acrylic- or vinylsulfonic acid to create the endo-osmotic flow. In addition, these phases also possessed hydrophobic ligands (C4, C6 or C12). The phases were characterized by the %T/%C ratio, describing the ratio of the linear- and cross-linked monomers, respectively. In order to make the carbohydrates detectable the analytes were derivatized by reductive amination with 2-aminobenzamide, and detected after separation by the Cd laser at 325 nm. This technique was used for malto-oligosaccharides (glucose Glc 1 to maltohexose Glc 6) with a 32 cm (25 cm packed length) x 100 ~tm capillary at 950 V/cm, 20 ~tA current. Altematively, absorption at 200 nm can be used for detection [36]. Another possibility published in the same paper [36] introduces the possibility of separating oligosaccharides (Glc 1 - G l c 4) in a C4-enriched monolithic packing (again, a polyacrylamide-polyethylene glycol packing) (C4 ligand, 15%; vinylsulfonic acid, 15%). The capillary used was 50 cm (40 cm packed length) •

Applications

349

21j3

I

I

,

0

2

,,,

4 Time/ min

I

,

I

6

,

I

8

Fig. 10.6. Separation of PAHs with the poly(TBAAm-co-AMPS)-coated column. Conditions: column, 600 mm x 25 ~tm i.d. (450 mm effective length); mobile phase, 30% acetonitrile (v/v) in 50 mM Tris-HCl buffer; field strength, 400 V/cm; injection, 12 kV for 5 s at the side of the anode; detection wavelength, 254 nm. Peak identification: 1, naphthalene; 2, fluorene; 3, phenanthrene; 4, pyrene; 5, benz[a]anthracene. Reproduced with permission from Sawada and Jinno [ 11].

100 ~tm i.d., with mobile phase containing 0.1% acetic acid and 5% (v/v) acetonitrile. The results of the separation are shown in Figs. 10.7 and 10.8. Apart from the monolithic polyacrylamide-polyethylene glycol-based packings, another possibility for separating oligosaccharides uses ODS silica on which 75% of the original silanol groups are not modified by the C18 moiety [39,189,190]. The analytes were derivatized to yield p-nitrophenylglycosides prior to separation. The success of the separation is based on a balance between the proportion of the hydrophobic (ODS) moiety present (where the partition occurs) and the unmodified silanol groups which ensure adequate endo-osmotic flow. This approach is also capable of separating anomers (in the presence of a borate buffer and a small proportion of an organic modifier). A typical example, for p-nitrophenyl-c~-D-glucopyranosides and malto-oligosaccharides is in Fig. 10.9, where the results obtained with NaH2PO4water-acetonitrile mixtures of different concentrations are shown. Objections have been raised about the reproducibility of capillary separations (both in capillary electrophoresis and capillary electrochromatography) but we should emphasize that in the latter case the values of retention were 0.21-0.55% RSD, with plate-numbers of (72-152)• 103 per metre [ 189].

References pp. 413-419

Chapter 10

350

7.1 ilc3

Glel e2

GIc5

G! 4

1.6

1'0 Time (rain)

1'1

1'2

28.5

8

9

i0

11 12 13 1'4 Time (min)

15

1'6

Fig. 10.7. (A), Isocratic electrochromatography of malto-oligosaccharides (glucose (Glcl)-maltohexaose (Glc6)) in a capillary filled with a macroporous polyacrylamide/ poly(ethylene glycol) matrix, derivatized with a C4 ligand (15%) and containing vinylsulfonic acid (10%). 2-Aminobenzamide was used to tag the oligosaccharides for the laser induced fluorescence detection (LIF). Analysis (B) is the same as (A) including the peak of derivatization agent, which appears at 14-16 min. Conditions: capillary, 320 mm (250 mm effective length) x 100 ~tm i.d.; mobile phase, 10 mM Tris/15 mM boric acid, pH 8.2; field strength, 900 V/cm; injection 0.5 kV/1 s; detection, LIF (helium-cadmium laser, excitation at 325 nm, emission at > 495 nm). Reproduced with permission from Palm and Novotny [36]. Copyright 1997 American Chemical Society.

10.6 FATTY ACIDS

Fatty acids have so far been analysed by CEC either as the free acids or as phenacyl- or methyl esters. Aqueous acetonitrile (50 mM) at pH 6 (9:1 v/v) was shown to be the optimal mobile phase [ 191 ]. It is generally known that in reversedphase chromatography free fatty acids and fatty acid methyl esters separate according to the partition number, which is defined as the carbon number minus twice the number of double bonds. A double bond reduces the retention time by the equivalent

Applications

351

25

Glc7 Glc$ I Glc6

o,~ II 1'6

1'8

2'0 2'2 2'4 2~6 2'8 Time (min)

i

30

Fig. 10.8. Isocratic electrochromatography of the oligosaccharide ladder in a capillary filled with a macroporous polyacrylamide/poly(ethylene glycol) matrix, derivatized with a C4 ligand (15%) and containing vinylsulfonic acid (10%). Conditions: capillary, 500 mm (400 mm effective length) x 100 lam i.d.; mobile phase, acetic acid 1:1000 containing 5% (v/v) acetonitrile; field strength, 600 V/cm; injection 5 s (100 V/cm); detection, LIF (helium-cadmium laser, excitation at 325 nm, emission at > 495 nm). Reproduced with permission from Palm and Novotny [36]. Copyright 1997 American Chemical Society.

of two carbons (pairs corresponding to this rule are called "critical pairs"). With acetonitrile-50 mmol/1 MES [2-(N-morpholino)ethanesulfonic acid], pH 6 (9:1), and a run-voltage of 30 kV per capillary (30 or 40 cm long, 100 lam i.d.), good results can also be obtained. Fatty acids and fatty acid phenacyl esters originating from vegetable oils and margarines were subjected to capillary electrochromatography by Dermaux et al. [51 ]. Using the Hewlett Packard system for capillary electrophoresis pressurized from both sides at 10 bar, and fused silica capillary columns 35/25 and 50/40 cm long (100 lam i.d.) (the latter number specifies the active length of the packed bed), slurry-packed with 3 lam Hypersil ODS, the results shown in Fig. 10.10 were obtained. This Figure shows not only the comparison of results obtained using CEC with two different lengths of the packed bed, but also a comparison with micro reversed-phase chromatography (position A). Fatty acid phenacyl esters were run in acetonitrile-50 mM MES pH 6 (90:10), at 30 kV and 20~

the injection was electrokinetic. A comparison

of the separations obtained with derivatized and underivatized fatty acids is given in Fig. 10.11. As one would expect, it is also possible to separate at least some positional and geometrical isomers (Fig. 10.12).

References pp. 413-419

352

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0.006 0.004

<

0.002 0

0.005 I

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0

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,

.

10

. . . . . . . . .

.

30

20

.

.

.

.

40

J 50

Miu

Fig. 1 0 . 9 . Electrochromatograms of p-nitrophenyl-c~-D-glucopyranosides and malto-oligosaccharides. Column, 200 (270) mm x 100 pm i.d. packed with ODS; mobile phase; (a), 5 mM NaH2PO4, pH 6.0-water-acetonitrile 45:45:10 (v/v/v); (b), 5 mM NaH2PO4, pH 6.0-water-acetonitrile 42.5:42.5:15 (v/v/v); (c), 5 mM NaH2PO4, pH 6.0-water-acetonitrile 40:40:10 (v/v/v). Voltage, 20 kV; detection, UV at 254 nm; 18~ Peak identification: 1, p-nitrophenyl-ot-D-glucopyranoside; 2, p-nitrophenyl-c~-D-maltoside; 3, p-nitrophenyl-cz-D-maltotrioside; 4, p-nitrophenyl-oc-D-maltotetraoside; 5, p-nitrophenylGc-D-maltopentaoside; Reproduced with permission from Yang and E1 Rassi [39].

10.7 TRIGLYCERIDES

Separation of triglycerides is amenable to capillary electrochromatography using a 3 ~tm Hypersil ODS-packed column with acetonitrile-isopropanol-n-hexane (57:38:5 v/v/v) as mobile phase. With a 20- or 40 cm x 100 ~m i.d. column the result seen in Fig. 10.13 can be obtained at 30 kV and 20~

[52]. The retention parameters were

quite acceptable, as demonstrated by RSD of the individual analytes present. The same approach is recommendable for analysing a formulation consisting of some

Applications

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3

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.

.

.

.

, 15

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

rain

,

(c) 4

1'0

3~)

4~3

min

Fig. 10.10. Separation of the fatty acid phenacyl esters (FAPEs) by (A), micro RPLC; (B), CEC on a 25 c m - and (C), on 40 cm column. (A); Column, 300 mm x 320 [am i.d., FSOT, RoSil C18 HL 3 [am. Conditions: mobile phase, methanol-water 95:5 (v/v); injection volume, 60 nl; detection, UV at 242 nm. (B), Column, 250 mm x 100 [am i.d., FSOT, Hypersil C18 3 [am. Conditions: mobile phase, acetonitrile-50 mM MES, pH 6, 90:10 (v/v); injection, 10 kV for 3s; detection, UV at 242 nm; temperature, 20~ voltage, 30 kV. (C), Conditions as described in (B). Peaks: 1, 18:3; 2, 18:2cc; 3, 16:0; 4, 9c-18:1; 5, 18:0. Reproduced with permission from Dermaux et al. [51 ].

testosterone esters and a lipid mixture, using a 25 cm long column. Because the testosterone esters exhibit lower lipophilicity, they elute ahead of the triglycerides.

References pp. 413-419

354

Chapter i 0

(A)

(B)

2

3 I 1

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~

'

~ .....

4o'

-

min

'

9

10

T.

-

v

20

~

-

~

30

'

,

min

Fig. 10.11. Separation of: (A), the FAPEs and (B), the free fatty acids (FFAs) of soyabean oil by CEC on a 40 cm column. (A), Conditions as described in Fig. 10.10C; detection, however, at 200 nm. Peaks: 1, 18:3; 2, 18:2; 3, 18:1; 4, 16:0; 5, 18:0. Reproduced with permission from Dermaux et al. [51].

1

_1

zb. . . . . . 3 o - mid

Fig. 10.12. Separation of some positional and geometrical 18:1 isomers by CEC on a 40 cm column. Conditions as described in Fig. 10.10C. Peaks: 1, 9c-18'1" 2, 9t-18"1" 3, 6c-18:1" 4, 6t-18"1. Reproduced with permission from Dermaux et al. [51 ].

Applications

355

(a)

(b)

6 8

6

8

12

lO

11 ~131~

0

20

40

60

1

min 0

20

40

60

80

min

Fig. 10.13. Triglyceride analysis of corn oil by, (a), micro-LC and (b), CEC. (a), Column, 2 x 250 x 1 mm i.d. Biosil C18 HL 5 ~tm; mobile phase, acetonitrile-isopropanol-n-hexane 57:38:5 (v/v/v) at 55 ~tl/min; temperature, 20~ detection, evaporative light scattering detection (ELSD) (Sedex 45) at 50~ 1.8 bar nitrogen. (b) Column, 400 (500) mm x 100 ~tm i.d. packed with 3 ~tm Hypersil C18; mobile phase, acetonitrile-isopropanol-n-hexane 57:38:5 (v/v/v) + 50 mM ammonium acetate; voltage, 30 kV; temperature, 20~ detection, UV at 200 nm; injection, 10 kV for 3 s. Peak identification: 1, LLLn; 2, LLL; 3, OLnL; 4, OLL; 5, OLnO; 6, PLL; 7, POLn; 8, OLO; 9, SLL; 10, PLO; 11, PLP; 12, OOO; 13, SLO; 14, POO; 15, PLS; 16, POP. Reproduced with permission from Dermaux and Sandra [204].

10.8 STEROIDS

There are basically two types of steroid- containing samples that come for analysis, namely endogenous steroids and steroid drugs. In this Section we shall mainly describe the capillary electrochromatographic behaviour of the first category: the other is deferred to the Section on drugs. In the paper by Stead, Read and Taylor [62] a comparison of capillary electrochromatography with two different HPLC separations is presented (Fig. 10.14). For electrokinetic chromatography a 3 ~tm ODS Hypersil-packed capillary 20 (35 cm) • 100 [am i.d., with acetonitrile-methanol-20 mM Tris HC1, pH 8.0 (37.5:37.5:5.25) at 15 kV, was used. Also, in another paper [72], isocratic elution was used for the separation of hydrocortisone, testosterone, c~-methyl-testosterone, and progesterone. Owing to the physico-chemical properties of steroids, it is perhaps not surprising that ODS-packed columns are widely used [65,43,69,68,71,72]. Typical separations of test mixtures of endogenous steroids were done with 1.5-1.8 ~m porous or non-porous silica in a 23.8 (32 cm) • 100 ~tm capillary with acetonitrile-water (6:4, v/v) containing 5 mM SDS and made 1.6 mM with respect to sodium tetraborate (pH 9.25, 20 kV, 10 bar overpressure on both ends, 25~

References pp. 413-419

For real samples (typically equine

356

Chapter 10

(A)

(c)

(B)

thiourea

I

2

mAU

[,~

128 mAU

n~U

TA

TA 17P

17P

TA 9 17P 20P

I

0

I

I

I

l

10

I

I

0

I

I

time

1

I

10

I

i

I

0

10

N

i

I i

20

30

P

t

(rain)

Fig. 10.14. Separation of progesterone and its metabolites obtained by: (A), CEC, conditions: capillary, 350 (200) mm x 100 ~tm i.d. packed with 3 ~tm ODS Hypersil; mobile phase, acetonitrile-methanol-20 mM Tris-HC1, pH 8 (37.5:37.5:25 v/v/v); injection 5 s at 15 kV; (13), HPLC, conditions: column 20 cm x 4.6 mm i.d. packed with 3 ~tm ODS Hypersil; mobile phase, acetonitrile-methanol-20 mM Tris-HC1, pH 8 (37.5:37.5:25 v/v/v); flow-rate, 0.6 ml/min; injection 20 pl; (C) HPLC using mobile phase acetonitrile-methanol-20 mM Tris-HC1, pH 8 (30:20:50 v/v/v); flow-rate, 2.1 ml/min. Reproduced with permission from Stead et al. [62].

urine) pre-separation and gradient elution appear necessary [61 ]. The sorbent used can be, e.g., Apex ODS-3 with water (A), and 5 mM ammonium acetate in acetonitrile (B), from 1% to 80% B over 10 min. In another separation by the same authors, the gradient went from 9% to 80% B over 5 min [61 ]. Huber and his coworkers [68] also used gradient elution from a 9.6/17.6 • 50 pm i.d. Zorbax ODS (6 pm) column. Starting with acetonitrile-10 mMborate (65:35), pH 8 (A), at the same pH, with acetonitrile-10 mMborate (B), at a ratio 85:15, changing over 5 min from 0 to 100% B, then with 3 min at 100% B, yielded a baseline separation of formamide (internal standard), corticosterone, androsten-3,17-dione, androstan-3,17-dione and pregnan-3,20-dione (Fig. 10.15). An interesting comparison of separations that can be obtained by capillary electrochromatography, capillary electrophoresis, and high-performance chromatography has been published [192]. Sample pretreatment is needed in all cases (including capillary electrochromatography) to achieve reasonable lifetimes of the columns used, and the samples must be preconcentrated to overcome the high limit of detection. This is usually done by injecting the sample in a non-eluting solvent.

Applications

357

..9

6

"O.ooo..o ........2 3"............

=...= 0E LLI ............... 1.09 '~' ............ 85

,/- ......

@

1.43 ~'

l

/

/

F

4 // // ,/

//,/" 5

65 i

0

I

2

I

4

I

minutes

6

I

8

Fig. 10.15. Gradient elution CEC separation of steroid hormones. Conditions: capillary, 9.6 (17.6) cm x 50 ~m i.d. packed with 6 ~m Zorbax ODS; mobile phase, A, acetonitrile-10 mM borate, pH 8 (65:35 v/v); B, acetonitrile-10 mM borate, pH 8 (85:15 v/v), 0-100% B in 5 min, 100% B for 3 min at 0.1 ml/min; 14 kV; 25~ detection, UV at 205 nm; injection, 1 kV/0.5 s. Peak identification: 1, formamide; 2, corticosterone; 3, testosterone; 4, androsten-3,17-dione; 5, androstan-3,17-dione; 6, pregnan-3,20-dione. Reproduced with permission from Huber et al. [68]. Copyright 1997 American Chemical Society.

Mayer et al. [60] have used steroid and steroid glycoside mixtures for proving that tapered capillaries can be used to retain the packed bed in the appropriate position. For steroid separation 3 ~tm Hypersil ODS was used with two types of capillaries differing in their dimensions (depending on whether a frit and a taper, or a double taper were used) with acetonitrile-20 mM sodium acetate (70:30, v/v), pH 5.0 (voltage 20-25 kV at 30~

as mobile phase. Under very similar conditions (26.0/24.5 cm

capillary, 100 ~tm i.d., acetonitrile-25 mM Tris 1:1 v/v at pH 8.0, containing 0.3% SDS at 15 kV) separation of digitalis glycosides can be carried out. Separations of complex steroid mixtures were achieved recently by Que et al. [76] using both isocratic and gradient elution. Mass spectrometric detection gave femtomole detection limits while laser-induced fluorescence of dansylated ketosteroids ranged in attomole levels (Fig. 10.16). Monolithic column packings were used with a 35 cm (25 cm packed bed) x 100 lam i.d. capillary packed with a polymer prepared from 5% T (total monomer concentration), 60% C (total crosslinker concentration), 3% polyethylene glycol, 10% vinylsulfonic acid and 15% lauryl acrylate. Details of the monolithic column preparation can be found in refs. 36,76, and 193. Similar monolithic columns can be used for the separation of bile acids [ 194].

References pp. 413-419

35 8

Chapter 10

5 6 8

1 3

5

10

15

7b

20

25

Time (min)

Fig. 10.16. Gradient electrochromatogram of derivatized neutral steroids. Macroporous monolithic column 350 (250) mm x 100 gm i.d.; gradient mobile phase, acetonitrile-water-240 mM ammonium formate buffer, pH 3 (30:60:5-65:30:5 v/v/v); field strength, 600 V/cm; injection, 100 V/cm for 10s; detection, laser-induced fluorescence (excitation at 325 nm, emission at >495 nm). Reproduced with permission from Que et al. [76].

10.9 AMINO ACIDS Separation of amino acids in capillary electrochromatography can be achieved either on monolithic columns or on C 18 modified packings. With monolithic columns, cross-linked polyacrylamide containing 2-methyl-l-propanesulfonic acid (to create the endo-osmotic flow) is used [195,196]: the separated analytes are dansylated and the order of eluted analytes reflects their molecular size. PTH (phenylthiohydantoin-) derivatives can be separated on, e.g., Zorbax ODS (3.5 lam packing) either in isocratic or gradient mode [68]. However, the selectivity of the isocratic elution mode is not sufficient for complete separation of the protein constituting amino acids. Huber et al. [68] introduced an acetonitrile-phosphate buffer gradient (A, 5 mM phosphate-acetonitrile, 7:3; B, 5 mM phosphate-acetonitrile, 40:60, from 0 to 100% B over 20 min). In this system the authors [68] were able to separate twelve PTH amino acids (including phe, ile, leu and val, which are usually difficult to separate) (Fig. 10.17). Using the isocratic mode offered the possibility of separating a mixture of only six PTH amino acids. Very recently, pressurized gradient capillary electrochromatography was used to separate seventeen 2,4-dinitrophenyl amino acid derivatives using a C18 reversedphase column (3 lam, 130 mm • 75 lam i.d.) using 50 mM sodium acetate, pH 6.4, as

359

Applications

Q. i... I--

1.2 ~' E E i-..i

<

LL

o

!11

I.-

0.9 "~

I-u~

<

....

~,

I

I

I

i

0

5

10

15

-52.5

minutes

Fig. 10.17. Capillary electrochromatography of PTH-amino acids with gradient elution. Column, 207 (127) mm x 50 pm i.d. packed with 3.5 pm Zorbax ODS particles, 80 A pores. Starting eluent (A), 5 mM phosphate, pH 7.55, 30% acetonitrile; gradient former (B), 5 mM phosphate, pH 7.55, 60% acetonitrile; flow-rate (through inlet reservoir), 0.1 ml/min; gradient, 0-100% B in 20 min; voltage 10 kV; current, 1 pA; temperature, 25~ UV detection at 210 nm; electrokinetic injection, 0.5 s, 1 kV. Peaks in order of elution: formamide; PTH-asparagine; PTH-glutamine; PTH-threonine; PTH-glycine; PTH-alanine; PTH-tyrosine; PTH-valine; PTH-proline; PTH-tryptophan; PTH-phenylalanine; PTH-isoleucine; PTH-leucine. The concentration of the PTH-amino acids dissolved in the mobile phase was 30-60 gg/ml. Reprinted with permission from Huber et al. [68]. Copyright 1997 American Chemical Society. the eluting buffer in the presence of 1% N,N-dimethylformamide as ion-pairing reagent [197] (Fig. 10.18). 10.10 PEPTIDES

Peptide mapping represents one of the fundamental tasks in protein analysis. The fact that most of the applications of CEC are restricted to isocratic elution has considerably limited the practical applicability of this technique in the past, particularly in the analysis of biopolymers. Two approaches have been developed to generate continuous gradients, namely, the merging of two endo-osmotic flows [78] or the application of two separate HPLC pumps. The latter presents practically no complication with regard to instrumentation as the same equipment can be used for HPLC, References pp. 413-419

360

Chapter 10

4 16

7

I~

14 15

3 51

8

10

9

12

7

1

3

I

I

0

120

19

~ I

~

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10

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17

B

,.,__,.._j'~_...

I

I

360

480

I

I

el30 720 "urre(s)

I

I

t

840

930

1080

Fig. 10.18. The separation of eighteen-amino-acids samples by pCEC. Column, PC-C18, 3 gm, 130 mm x 75 ~tm i.d.; mobile phase; (A), 50 mM sodium acetate-l% N,N-dimethylformamide, pH 6.4; (B), 50% acetonitrile in water; linear gradient, 60-5% A in 6 min and kept at 5%; 3,000V positive voltage across the column and 1,000 psi pressure were added on column during the separation; flow-rate, 50 ~tl/min; 20~ detection, 360 nm; 20 nl injection. (A), 2 ~tg/ml of derived eighteen-amino-acids sample solution; (B), 2 ~tg/ml of derived eighteen-amino-acids standard solution. Reproduced with permission from Ru et al. [ 197].

CEC, and hybrid pressurized electrochromatography [68,66,99,104,106,119,198]. In the case applied for peptide mapping the instrumental steps were as described below. The HPLC system was operated at 0.3 ml/min and was connected through a T-piece with the packed capillary column. To ensure a proper flow-rate in the capillary one branch of the T-piece was connected to a flow restrictor and the packed capillary was connected through a stainless steel union to a resistor capillary. The T-piece was connected to a high-voltage power supply (+ end) while the metal union at the capillary column outlet, the restrictor at the other T-piece branch, and the two filters (one between the HPLC pump delivery line and the T-splitting piece, the other at the flow restrictor branch of the T-piece) were earthed (elecrically grounded). The filters pre-

Applications

361

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0

~--

TIME (min.) Fig. 10.19. Electrochromatogram of protein separation on a diol. Conditions: pH 4.14, capillary 450 mm x 50 ~tm i.d., 250 mm packed length. Peak identification: 1, cytochrome c; 2, lysozyme; 3, myoglobin; 4, ribonuclease A. Reproduced with permission from Pesek et al. [102]. vented clogging of the resistor- and separation capillaries. The column used was 150 ~tm i.d. and 20 cm long; 6 cm were packed with 1.5 ~tm ODS-silica spheres of 100 A pore-size; 0.1% TFA in water, pH 2, (A), and 50% acetonitrile, (B), were used as mobile phase (0-100% B) over a period of 10 min. The split was 1:300 in a split injection of 10 and 20 ~1 of a protein tryptic digest. There is a general problem with separating complex peptide mixtures by electrochromatography namely, the high concentration of trifluoroacetic acid in the eluent. This generates low pH values, high currents (and current fluctuations), strong electrical fields, and, consequently, bubble formation. Therefore either a considerable proportion (up to 50%) of the organic modifier, or a low concentration of trifluoroacetic acid (0.001%), have been used. However, in the latter case the peptide peaks tail badly and the resolution is considerably reduced [ 104]. Finally, the common procedure is to pressurize the capillary from both ends with nitrogen gas or, as reported by Behnke and Metzger [97], a restrictor capillary mounted to the column outlet can serve the same purpose (this has to have an i.d about 25 ~tm, and be 40 cm long). Diol-modified capillaries have been used successfully for the separation of peptides and proteins, as shown in Fig. 10.19. References pp. 413-419

Chapter 10

362 10.11 NUCLEIC ACIDS CONSTITUENTS

A thorough investigation of nucleosides was recently published by Helboe and Hansen [ 116]. Although no natural samples were reported, the separation of adenosine, cytidine, guanosine, inosine, thymidine and uridine was investigated and optimized with respect to buffer concentration, the effect of temperature and voltage, and acetonitrile concentration: validation data were also reported. The short-term- and the long-term repeatability of retention times were good, but, the area repeatability was poor (yielding RSD < 13% in the short term, and RSD < 23% in the long-term range). However, the authors claim that this can be improved considerably by using relative areas for the calculation (with thymidine as the intemal standard). The separation was performed with a packed bed capillary 25 cm long (CEC Hypersil C18 3 gm, Hewlett Packard) with an additional length of 8.5 cm of polyimide-coated capillary tubing. At least a 2 h conditioning period before a set of runs was necessary: during this period a voltage of 20 kV was applied. Both the inlet and the outlet were pressurized at 10 bar and the outlet- and inlet vials were replaced after 2 hours of run time. A background electrolyte (BGE) containing 3 mM TEA, 5 mM acetic acid (pH 5.0) (92 vol. %) containing 8 vol. % acetonitrile was found to be best. The optimization with respect to voltage and temperature is seen from Fig. 10.20.

'

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Fig. 10.20. Optimisation of temperature and voltage in separation of nucleosides. Column: 250 mm x 100 ~tm i.d. packed with CEC Hypersil C18, 3 lam, Conditions: mobile phase, (5 mM acetic acid, 3 mM TEA, pH 5)-acetonitrile, 92:8 (v/v); injection, 10 kV for 3 s. (A), 25~ and 20 kV; (B), 25~ and 25 kV; (C), 20~ 25 kV. Peak identification (in order of elution): 1, cytidine; 7, thiourea; 2, uridine; 3, inosine; 4, guanosine; 5, thymidine; 6, adenosine. Reproduced with permission from Helboe and Hansen [ 116].

Applications

0.025 L

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Time (min)

Fig. 10.21. Electrochromatogram of a mixture of purine and pyrimidine bases and their constituents. Conditions: capillary, 270 (205) mm x 100 ~tm i.d., whole capillary packed with 10 ~tm ODSS; mobile phase, 4.8 mM sodium acetate, pH 4.50, containing 40% (v/v) acetonitrile; voltage, +20 kV; injection, electrokinetic for 2 s at 1 kV. Peaks in order of elution: 1, Ura; 2, U; 3, Thy; 4, Cyt; 5, C; 6, I; 7, Ade; 8, Gua; 9, A; 10, G. Reproduced with permission from Zhang and E1 Rassi [ 117].

Instead of regular ODS systems, octadecyl-sulfonated silicas (ODSS) designed for capillary electrochromatography at strong electro-osmotic flow can be used (see Zhang and E1 Rassi [ 117]). This ODSS stationary phase is composed of a hydrophilic, negatively charged layer to which an additional (top) layer containing octadecyl ligands is attached covalently. While the sublayer ensures the endo-osmotic flow (owing to its ionizable groups) it is the upper layer which represents the area where

the reversed-phase partition occur (owing to its hydrophobic character). A typical electrochromatogram is seen in Fig. 10.21 where the separation of ten nucleosides and purine/pyrimidine bases is shown (using a 20.5 cm packed-bed capillary, 27 cm to the

detector with 100 ~tm i.d., containing 10 lam ODSS particles and operated with 4.8 mM sodium acetate, pH 4.5, containing 40% v/v acetonitrile, at 20 kV, with electrokinetic injection. 10.12 D R U G S

As in other separation techniques applied in the analysis of drugs there is no general rule that can advise the newcomer to the field on how to separate a particular drug (or drug candidate) from its metabolites or contaminants. Capillary electrochromatography does not offer such a solution, although this would be very desirable.

References pp. 413-419

364

Chapter 10

The capillary electrochromatographic separation of compounds (drugs) varying over a wide range of acidity or basicity has been explored recently by Lurie and Conver [136]. The test solutes were drugs of abuse, and the whole study was aimed towards forensic and toxicological analysis. Basically, the apparatus used was moreor-less conventional, using a CEC Hypersil C8 column (3 lam particle size) with acetonitrile-phosphate buffer pH 2.5 (containing hexylamine) as mobile phase. As demonstrated, fifteen basic, acidic and neutral drugs can be resolved by applying a stepped gradient. It is likely that the separation mechanism involved is of a multimode nature. Strongly and moderately basic drugs appear before the void volume (tO) of the column, which is ascribed to a combination of capillary zone electrophoresis taking part in the separation process. However, as with solutes that migrate after tO, the final separation reflects the simultaneous action of CEC and CZE phenomena. The problem that has been addressed by the above authors is of separating strongly basic, acidic and neutral compounds in a single run. In general, strongly and moderately basic drugs (e.g., tricyclic antidepressants) can be separated by CEC using a strong cation-exchanger, but multiple peaks and some focusing effects preclude this technique from practical applicability [65,199]. Bare silica with buffered acetonitrilewater phases can be used for the same purpose [200]. Also, in this case the separation mechanism involved is presumed to be a combination of different separation modes, namely cation exchange and normal phase chromatography. Strongly and weakly acidic drugs (typically cannabinoids) can be separated by using a C8 column with acetonitrile-phosphate at pH 8 [177] which can, with some limitations, also be used for weakly basic and neutral analytes. The problem is that under CEC conditions particularly, moderately basic organic solutes usually exhibit poor chromatographic performance, yielding multiple peaks (adsorption of heroin onto the unbound silanol groups may explain this phenomenon). To minimize tailing and non-specific sorption, addition of hexylamine has been recommended by several authors [136,201,202]. The results are impressive: quite unexpectedly, even when using hexylamine as mobile-phase modifier, the system still exhibits appreciable endo-osmotic flow (the retention time of a neutral marker was increased by only 2.5 min). Even strong bases such as amphetamine, methamphetamine, procaine, cocaine, heroin, quinine and noscapine (the pKa of which is below 7) can be separated in this system. All these solutes migrate before tO, indicating that electrophoretic phenomena play a fundamental role in this CEC mode. It is to be emphasized that the concentration of hexylamine should not exceed 2 ~tl/ml, as at higher concentrations lower endo-osmotic flow, higher current, and bubble formation degrade the separation. Having prevented the multiple peak formation and peak-tailing, Lurie and Conver also investigated the role of the acetonitrile concentration in the mobile phase. At

Applications

3 65

lower acetonitrile concentrations, lower separation voltages naturally have to be used as otherwise the current obtained is too high. All solutes moved before to (i.e., ahead of the neutral marker). Reducing the acetonitrile concentration from 75% to 30% not only slows down the separation and increases the resolution of basic compounds, but it also alters the selectivity of the system. It is noticeable that the retention time of the neutral marker is considerably increased at lower acetonitrile concentrations. This supports further the idea that several phenomena are involved in this type of separation. First, as the concentration of acetonitrile is increased, the relative hydronium ion concentration is lowered and the apparent pH is increased: concomitantly the pKa of the bases involved is lowered with increasing acetonitrile concentration. The plate counts varied between (8.2 and 13.0)•

depending on the nature of the analyte

separated. The final resolution reflects again a multimode mechanism in which hydrophobic and silanophilic effects as well as CZE phenomena are involved. It is proposed that hydrophobic interactions involve the interaction of the C8 moiety with the sorbent, while silanophilic phenomena include the interactions of the mobile phase with the free silanol groups (the tailing observed in the absence of hexylamine is ascribed to the latter category of interactions). To emphasize the role of column packing, i.e., to compare the chromatographic and CZE phenomena involved, it may be interesting to compare these modes. In the latter, the separation of identical solutes is made by CZE in the same background electrolyte as used for the CEC runs. The most distinct difference is the selectivity change, particularly at high (75%) acetonitrile concentration. It is to be emphasized that while the selectivity changes of quinine can be explained by the fact that the apparent pH of the mobile phase is near the first pKa of quinine, the selectivity alterations with other types of drugs, such as amphetamine, methamphetamine, heroin, and noscapin cannot be based on pKa considerations, and this is the area where hydrophobic and silanophilic interactions play their role. Of course, when comparing CZE and CEC separations the major differences in selectivity that occur can be explained on the basis of the much larger partition surface area available to analytes in a packed capillary. Also, at a given acetonitrile concentration the endo-osmotic flow in CZE is about 2-3 times higher than in capillary electrochromatography. Also, the peak counts with CZE are 7-20 times higher than with CEC, but the overall resolution is always better with CEC. This is attributed to a synergistic action of the chromatographic and electromigration phenomena. Strong bases (amphetamines, methamphetamine, procaine), bases of medium strength (cocaine, heroin, noscapine, quinine), weak bases (diazepam, methaqualone), strong acids (A-9-tetrahydrocannabinolic acid), moderately strong acids (phenobarbital) and weak acids (A-9-tetrahydrocannabinol) can be separated in a single run along with neutral solutes (testosterone, testosterone propionate). However, in order to

References pp. 413-419

366

Chapter 10

achieve this type of resolution an acetonitrile gradient has to be introduced. The separation combines an increased resolution of the strong bases and those of medium strength at lower acetonitrile concentrations, with good resolution (and higher speed of analysis) at a higher acetonitrile concentration. Strong and moderately strong bases elute before to, and all other solutes elute after to, which indicates that chromatographic partition plays the dominant role in the final result. At the end, it is useful to compare the results obtained with MEKC and CEC. The most obvious difference is that in MEKC (50 mM SDS, with buffers which are otherwise comparable to CEC, except for the pH value) moderately strong and strong bases elute after to (followed by, e.g., phenobarbital and methaqualone). Also, the elution order is nearly completely different for the two separation modes applied. Although both these categories of separation involve some sort of chromatographic partitioning along with electromigration phenomena, the result reflects the differences in pH needed for successful separation. Partitioning into the micelle which, although physically similar to reversed-phase packing, is still not the same, and ion-pairing between the micelle and the analytes all contribute to differences in selectivity between these two techniques. Naturally, the set of drugs investigated by Lurie, and Conver and Ford [136] can also be separated by reversed-phase chromatography. As would be expected, considerable differences in selectivity were revealed, particularly with strong and moderately strong bases. Moreover, in CEC these solutes elute before tO' while in HPLC they elute after to. Weak bases, weak-, and moderately weak acids, as well as neutral solutes exhibit similar selectivity in CEC and HPLC. Additional examples of drug separation can be seen from Figures 10.22, 10.23 [56] and 10.24 [176]. Drugs possessing a steroid structure are particularly easy to separate by CEC (see also the section on Steroids). Euerby et al. [203] published the separation of tipredane and five related compounds. A conventional capillary packed with 3 mm Spherisorb ODS-1 can be used for this purpose using acetonitrile-Tris pH 7.8 buffer (8:2) (50 mmol/1). Under these conditions it is also possible to separate the C-17 diastereoisomer of the active compound without the addition of a chiral modifier (b-cyclodextrin is needed to achieve a comparable result in other separation modes) (Fig. 10.25). The elution order of individual compounds was exactly the same as with reversed phase chromatography, and it was concluded that with unionized species HPLC methods should be directly transferable to the CEC mode. In order fully to achieve the potential of CEC separations the application of gradients in an automated sequence appears recommendable. The general approach starts with using the initial mobile phase followed by switching off the voltage and replacing both the inlet- and outlet vials for another couple containing the final (limiting) mobile phase. Then the voltage is switched on again and the analysis is continued.

Applications

3 67

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200

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116 1+2

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12

min.

Fig. 10.22. Comparison of separation of drug mix by CEC, HPLC and CZE. Column, Spherisorb ODS-1, 3 gm, 250 (335) mm x 100 lam i.d.; mobile phase, acetonitrile-25 mM phosphate, 0.2% hexylamine, pH 2.5 (80:20 v/v); voltage (CEC), 25 kV; pressure (HPLC), 200 bar; CZE, uncoated capillary 250 (335) mm x 75 lam i.d. Peak identification: 1, procaine; 2, timolol; 3, ambroxolol; 4, metoclopramide; 5, thiourea; 6, naproxene; 7, antipyrine. Reproduced with permission from Dittman et al. [56].

mAU

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71 80%ACN sol ~[iSI 7

5

10

15

20

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Fig. 10.23. Influence of percentage organic modifier and pH of the aqueous buffer on retention and selectivity. Column, Spherisorb ODS-1, 3 ~m, 250 (335) mm x 100 ~tm i.d.; mobile phase, acetonitrile-25 mM phosphate, 0.2% hexylamine; voltage, 25 kV. (A) pH 2.5, (B) pH 3.8. Peak identification as in Fig. 10.22. Reproduced with permission from Dittman et al. [56].

After the last peak is visualized by the detector, the voltage is switched off again and the original vials containing the start buffer are placed in the inlet and outlet positions. This approach has been applied to a set of six diuretics of different lipophilicities. The first four peaks were brought in front of the detector's window in a period of time

References pp. 413-419

368

Chapter 10

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Time / rain Fig. 10.24. Separation of benzodiazepines by CEC. Column, 200 mm (effective length) x 100 pm i.d. packed with cholesteryl-bonded silica. Mobile phases: (A), acetonitrile-5 mM Tris-HC1 buffer pH 7.7 (35:75 v/v); (B), acetonitrile-5 mM Tris-HC1 buffer pH 73 (35:65 v/v). Applied voltage 300 V/cm. Reproduced with permission from Jinno et al. [ 176].

similar to that in the isocratic run, and it was proposed that during this period the second buffer did not yet penetrate the capillary column while the remaining two solutes apparently experienced the gradient effects and gave very sharp peaks.

10.13 ANTIBIOTICS Antibiotics represent a chemically quite heterogenous category of compounds, for which the C 18 phases appear sorbents of the first choice. A group of these compounds (tetracyclines) was successfully separated by open tubular capillary electrochromatography using C18 modified capillaries. The results reported to be obtained in this system were better than those obtained with either monolithic columns or open diol columns [123] (Fig. 10.26).

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Fig. 10.25. Separation of tipredane (compound 14) from its five related substances (compounds 11-13, 15, 16) by CEC. Column: 250 mm x 50 ~tm i.d. packed with 3 ~tm Spherisorb ODS-1; mobile phase, acetonitrile-50 mM Tris buffer, pH 7.8, 80:20 (v/v); applied voltage, 15 kV; temperature, 15~ electrokinetic injection, 5 kV/15s. From Euerby et al. [63], @Journal of Microcolumn Separations, 1997. Reproduced with permission of John Wiley & Sons, Inc.

10.14 PESTICIDES So far, capillary electrochromatography has found only a few applications to pesticides. Separation of cinosulfuron and its by-products can serve as a good example. The separation was done with a C-8 non-endcapped 3 ~tm packing in a capillary whose dimensions were 33.5 cm (25.0 cm to the detector) x 100 ~tm internal diameter. A typical separation is presented in Fig. 10.27 [ 128]. 10.15 AVAILABLE A P P L I C A T I O N S (summarizing Table) It is evident that the different applications of electrochromatographic procedures are growing every day. Because some of the procedures represent only small variations of published procedures already applied to another category of compounds, we have decided to supply the basic information available for the various categories of compounds discussed briefly in the preceding sections. Although we have attempted to make this Table as complete as possible, we cannot guarantee that all application papers are included, for a number of reasons. Nevertheless we believe that this rather extensive Table can serve as a source of information for readers and can help newReferences pp. 413-419

3 70

Chapter 10

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Fig. 10.26. Separation of some likely components found in commercial tetracycline. (A), Bare capillary with buffer pH 3.0 (30 mM citric acid and 24.5 mM 13-alanine) 500 mm x 50 pm i.d.; 30 kV, electrokinetic injection for 5 s at 10 kV; (B), bare capillary, same as for A but with buffer-methanol (60:40 v/v); (C), Hydride (etched) capillary 250 mm x 50 pm i.d., other conditions same as for B; (D), C18 etched capillary, conditions same as for C. Peak identification: 2, tetracycline; 3, chlorotetracycline; a, 4-epitetracycline; b, anhydrotetracycline; e, 4-epianhydrotetracycline. Reproduced with permission from Pesek and Matyska [123].

comers to appreciate the categories (sets) of compounds for which capillary electrochromatography can be exploited.

Applications

3 71

N OCH3

UV-absorbance at 209 nm [mAU]

Cinosulfuron ~y- SO2-NH-CO-NH-~N 0CH2CH20CH3

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Fig. 10.27. CEC separation of Cinosulfuron and by-products. Column: 335 (250) mm x 100 pm i.d. packed with Synchropak non-endcapped C8, 3 pm, Conditions: mobile phase, acetonitrile-20 mM sodium dihydrogenphosphate, pH 4.0 (60:40 v/v); separation voltage, 15 kV; injection, 5 kV, 3s; applied pressure, 8 bar on both sides; sample, 2.4 mM cinosulfuron dissolved in mobile phase. Reproduced with permission from Rapp et al. [ 128].

References pp. 413-419

TABLE 10.1

taO "--1 bO

APPLICATIONS (compositions of mobile phases are described using volume proportions, i.e., v/v)

Compound

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

Acetonitrile-water (40:60) pH 9.1

500 mm x 50 ~tm i.d.

420 mm to the detector, 6 25~

Hydrocarbons and fullerenes Alkylbenzenes (toluene, ethyl- 40% Ethylacrylate, 50% benzene, propylbenzene, methacrylic acid, 10% lauryl butylbenzene) methacrylate (custom-made polymer), 4 wt.% polymer concentration Polyaromatic hydrocarbons (benzene, toluene, naphthalene, acenaphthylene, fluorene, anthracene, 1,2-benzanthracene, phenol, acetone)

ODS Hypersil, 5 gm

Gradient acetonitrile-water (from 60:40 to 90:10)

500 mm x 75 ~tm i.d.

200 mm packed length, flow-injection analysis

7

Polyaromatic hydrocarbons

40% Ethylacrylate, 50% methacrylic acid, 10% lauryl methacrylate (custom made polymer), 3.99-4.01 wt.% polymer concentration

Acetonitrile-buffer (40:60); pH 9.2 (10 mM sodium borate) or pH 11.5 (10 mM phosphate)

300 mm x 50 ~tm i.d.

250 mm to the detector

6

Polyaromatic hydrocarbons and parabenes (ethyl-hexyl parabene, naphthalene, fluorene, phenanthrene, fluoranthene, pentylbenzene)

Excil C 18

Acetonitrile-Tris HC1 buffer pH 8.0 (80:20)

380 mm x 100 p,m i.d.

85 mm packed length

8

"x

"~ .~ ~, & ~

Polyaromatic hydrocarbons Micra ODS-1, 1.5 gm or (benzene, naphthalene, Micra ODS-2, 3 gm acenaphthylene, fluorene, acenaphthene, phenanthrene, anthracene, fluoranthene, pyrene, benz[a]anthracene, chrysene, benzo[b]fluoranthene, benzo[k]fluoranthene, benzo[a]pyrene, dibenz[a,h]anthracene, indeno [ 1,2,3-cd]pyrene, benzo[ghi]perylene)

Acetonitrile-10 mM 440 mm x 50 ~m i.d. ammonium acetate, pH 6.5 (70:30) or acetonitrile-50 mM Tris buffer, pH 8.1 (80:20)

Polyaromatic hydrocarbons Spherisorb ODS-1, 3 gm or (benzene, naphthalene, Spherisorb ODS 2, 3 gm acenaphthylene, fluorene, acenaphthene, phenanthrene, anthracene, fluoranthene, pyrene, benz[a]anthracene, chrysene, benzo[b]fluoranthene, benzo[k]fluoranthene, benzo[a]pyrene, dibenz[a, h] anthracene, indeno [ 1,2,3-cd]pyrene,

Acetonitrile-50 mM Tris buffer, pH 8.1 (80:20)

430 mm x 50 ~tm i.d. 350 or 410 mm packed or 530 mm x 75 p.m i.d. length

Acetonitrile--0.1 M acetate buffer, pH 3.0 (80"20)

250 or 170 mm (non-entrapped or entrapped), 75 gm i.d.

340 mm packed length, 9 voltage programming tested

r,,~~

r,,~~

t~

9

benzo[ghi]perylene) Polyaromatic hydrocarbons Nucleosil ODS, 5 gm (naphthalene, fluorene, phenanthrene, anthracene), acetone

250 and 170 mm are 10 effective column lengths

ta~

ta9 4~

TABLE 10.1 (continued)

Compound

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

Polyaromatic hydrocarbons (naphthalene, fluorene, phenanthrene, pyrene, benz[a]anthracene

Linear polymer coated capillary [poly(N-tert.-butyl acrylamide-co-2-acrylamido2-methyl- 1-propanesulfonic acid]

Acetonitrile-50 mM Tris buffer, pH 7.3 (30:70)

600 mm x 25 ~tm i.d.

450 mm effective column length

11

Polyaromatic hydrocarbons (benzene, toluene, ethylbenzene, propylbenzene, butylbenzene, pentylbenzene)

Separation segment: ODS, 5 pm (200 mm), accelerating segment: bare silica (0-280 mm)

Acetonitrile-1.25 mM monobasic sodium phosphate pH 6.0 (75:25)

100 lam i.d., for length Two types of packing see stationary phase

12

Polyaromatic hydrocarbons (sixteen US EPA PAH)

SynChrom ODS (3 pm, 90%) Acetonitrile-4 mMborate + Spherisorb SW (80:20) (1 lam, 10%)

330 mm x 75 gm i.d.

13

Polyaromatic hydrocarbons (sixteen US EPA PAH)

Nonporous ODS (90%) + 1 ~tm silica gel (10%)

Acetonitrile-2 mMTris, pH 9 200 (300) mm x 100 (65:35) pm i.d.

14

Fullerenes C60, C70

Vydac C18 (3 gm)

Acetonitrile-tetrahydrofuran (1"1)

400 mm x 5 gm i.d.

15

Alkylbenzenes

Custom made styrenedivinylbenzene polymer 3 and 6 lam

Acetonitrile-water (80"20)

300 mm x 100 pm i.d.

16 c~

~ ~ .~ ~" tao ~.

17

Polyaromatic hydrocarbons Nucleosi14000-7 C18, (naphthalene, fluorene, anthra- Nucleosil 1000-7 C 18, cene, pyrene), acetone Nucleosil 500-7 C 18

Acetonitrile-water (80:20) containing 10 mM borate pH 8.3

720 mm x 100 pm i.d.

Polyaromatic hydrocarbons Modified C18 open tubular (naphthalene, acenaphthylene, column acenaphthene, fluorene, phenanthrene, anthracene, fluoranthene, pyrene)

Methanol-1 mM phosphate buffer, pH 7.0 (65:35)

400 mm x 9.6 pm i.d.

Aromatic compounds (benzene, 1,2,3-trimethylbenzene, nitrobenzene, etc.)

Spherisorb ODS, 3 pm

Isocratic elution: acetonitrile- 435 mm x 75 gm i.d. 4 mMTris, pH 9.2 (60:40); gradient elution: acetonitrile4 mMTris, pH 9.2 (60:40)

158 mm packed length

19

Aromatic compounds (alkyl benzenes)

Spherisorb ODS-2, 3 pm

Acetonitrile-4 mMTris, pH 9.2 (80:20)

270 mm x 75 pm i.d.

66 mm packed length, high speed CEC

20

Aromatic compounds (alkyl benzenes)

Non-porous ODS, 2 pm

Acetonitrile-4 mMTris, pH 9.2 (60:40)

270 mm x 75 gm i.d.

63 mm packed length, high speed CEC

20

Aromatic hydrocarbons (naphthalene, 2-methylnaphthalene, fluorene, phenanthrene, anthracene)

2-Hydroxymethacrylatepiperazine copolymer with C 18 ligands and immobilized dextran sulfate

62% Acetonitrile in 5 mM borate, pH 8.7

550 mm x 25 pm i.d.

525 mm active length

21

Aromatic hydrocarbons (2-methylnaphthalene, fluorene, phenanthrene, anthracene)

2-Hydroxymethacrylatepiperazine copolymer with C 18 ligands

50% Acetonitrile in 4 mM sodium phosphate, pH 7.4 (60%), acetonitrile-buffer (60:40) or gradient from 60:40 to 70:30 acetonitrilebuffer

200 mm • 75 9m i.d.

160 mm active length; comparison of isocratic and gradient elution

22

r.,~~

300 mm to the detector, 18 optimization of mobile phase, open tubular electrochromatography

r~

taO

L~

TABLE 10.1 (continued)

Compound

Stationary phase

0.3 wt.% 2-acrylamido-2Benzene derivatives (benzyl methyl- 1-propan sulfonic alcohol, benzaldehyde, benacid, pore size 465 nm zene, toluene, ethylbenzene, propylbenzene, amylbenzene), thiourea Polystyrene standards (987.103 rel. mol. mass) and toluene

Butyl methacrylate-ethylene dimethacrylate copolymer containing 0.3 wt.% 2-acrylamido-2-methyl- 1-propanesulfonic acid

Note

Ref.

Mobile phase

Capillary dimensions

Acetonitrile-4 mM phosphate buffer, pH 7 (80:20)

250 mm x 100 ~tm i.d. (250 mm active length)

23

Tetrahydrofuran containing 2 vol.% of water

300 m m x 100 ~tm i.d. (300 mm active length)

24

CEC Hypersil C 18, 3 gm Polyaromatic hydrocarbons and parabenes (thiourea, ethyl-hexylparabene, naphthalene, fluorene, phenanthrene, anthracene, fluoranthene)

450 mm x 100 gm i.d. Acetonitrile-water-25 mM Tris.HC1, pH 8.0 (gradient from 20 to 80% of acetonitrile)

250 mm to the detector 25

Hypersil C 18, 3 ~tm Aromatic hydrocarbons and other compounds (thiourea, benzyl alcohol, methyl benzoate, toluene, benzophenone, naphthalene, 1,4-dichlorobenzene, phenothiazine, biphenyl, 1,3,5-trichlorobenzene, 1,2,4,5-tetrachlorobenzene)

Acetonitrile-5 mM phosphate buffer, pH 7 (50:50)

Packed length 250 mm, 26 separations at 60 ~ and 20~ compared (inclusive temperature gradient)

335 mm x 100 ~tm i.d.

Aromatic hydrocarbons (naphthalene, phenanthrene, pyrene) 4~

Tetraethoxysilane-noctyltriethoxysilane hybrid gels

Aromatic hydrocarbons, Sol-gel bonded 3 gm benzyl alcohol, benzaldehyde, ODS/SCX, 80 A pores benzophenone, biphenyl, benzene, toluene, ethyl- and butylbenzene

Acetonitrile-water (1" 1)

470 mm x 50 ~tm i.d.

400 mm to the detector

27

Acetonitrile-water (80:20) containing 1.5 mM phosphate buffer, pH 3.0

340 mm x 75 lam i.d.

250 mm active length

28

Acetonitrile-25 mM morpholinoethanesulfonic acid (95:5), pH 6.2

300 mm x 75 gm i.d.

1O0 mm packed length

29

Polynuclear aromatic hydrocarbons (naphthalene, phenanthrene, pyrene), thiourea

Particle-loaded (3 gm, C18) monolithic sol-gel column

Polyaromatic hydrocarbons (naphthalene, fluoranthene), thiourea

BDS-ODS Hypersil, ODS Hypersil, Spherisorb ODS-2, Spherisorb ODS- 1, CEC Hypersil C 18

Polyaromatics (ethyl-hexylparabenes, naphthalene, fluorene, phenanthrene, anthracene, fluoranthene), thiourea

Hypersil ODS, 3 gm

Acetonitrile-25 mM Tris.HC1 buffer, pH 8.0 (80:20)

335 mm x 100 (75) gm i.d.

Polyaromatics (naphthalene, acenaphthene, phenanthrene, anthracene, fluoranthene, pyrene)

Capillary had porous silica layer (0.70 ~tm) with chemically bonded C 18

Acetonitrile-1 mM phosphate buffer, pH 7.0 (50:50)

400 mm x 9.60 gm i.d. 300 mm to the detector, 32 open tubular CEC

Comparison of different 30 C 18 stationary phases

250 mm effective 31 length, influence of the packing immobilization

TABLE 10.1 (continued)

Ref.

Stationary phase

Mobile phase

Capillary dimensions

Note

Phenols in tobacco smoke (hydroxyquinone, resorcinol, catechol, phenol, isomeric cresols)

Hypersil C 18, 3 ~tm

Acetonitrile-10 mM phosphate buffer pH 9.0 (70:30 or 60:40)

310 mm x 50 gm i.d.

250 mm to the detector, 33 25~

m-Cresol and pyridine

C18 Granocel-14Sh, 7 gm

Acetonitrile-2 mM sodium tetraborate (50:50)

283 mm x 150 gm i.d.

243 mm packed length

34

2-Phenylmethyl- 1-naphthol

Hypersil C 18, 3 gm

Acetonitrile-20 mM citrate, pH 4-8 (70-80/30-20)

350 mm x 100 gm i.d.

250 mm packed length

35

2-Phenylmethyl- 1-naphthol

Hypersil C 18, 3 ~tm

Acetonitrile-50 mM Tris-HC1, 350 mm x 100 gm i.d. pH 8 (80/20)

250 mm packed length

35

Compound

Phenols

Oxo compounds Acetophenone, propiophenone, butyrophenone, 2',5'-dihydroxyacetophenone, 2',5'-dihydroxypropiophenone

Custom-made macroporous 10 mM Tris + 15 mM boric polyacrylamide-poly(ethylene acid (pH 8.2) with 205 glycol) monolith (acrylate acetonitrile copolymer of acrylic acid, butyl acrylate, methylenebisacrylamide with 3% (w/v) poly(ethylene glycol))

250 mm • 100 ~tm i.d.

205 mm effective length 36

40% Ethyl acrylate, 50% methacrylic acid, 10% lauryl methacrylate (custom made polymer), 4% wt. polymer concentration

Acetonitrile-water (40:60) pH 500 mm x 50 gm i.d. 9.1

Acetone (along with polyaromatic hydrocarbons)

Nucleosil ODS, 5 ~tm

Acetonitrile-0.1 M acetate buffer pH 3.0 (80:20)

250 or 170 mm (nonentrapped or entrapped), i.d. not specified

10 250 and 170 mm are effective column lengths

Acetone, acetophenone, butyrophenone

Linear polymer coated capillary [poly(N-tert.-butyl acrylamide-co-2-acrylamido2-methyl- 1-propanesulfonic acid]

Acetonitrile-50 mM Tris buffer, pH 7.3

750 mm x 25 ~tm i.d.

600 mm effective column length

11

Formaldehyde, acetaldehyde, C18 bonded silica particles, acetone, propionaldehyde, cro- Unimicro 3 ~tm tonaldehyde, methacrolein, 2-butanone, butyraldehyde, benzaldehyde, valeraldehyde, p-tolualdehyde, hexaldehyde

60% Acetonitrile-4% tetrahydrofuran made 5 mM to Tris-HC1 (pH 8.0)or 60% acetonitrile-1 mM borate (pH 9.0) or 60% acetonitrile-4% tetrahydrofuran made 1 mM to borate (pH 9.0)

270 mm x 75 ~tm i.d.

Separated as DNP derivatives, 200 mm packed length

37

Thiourea, benzyl alcohol, Sol-gel bonded 3 ~tm benzaldehyde, benzophenone, ODS/SCX, 80 A pores biphenyl

Acetonitrile-water (80:20) containing 1.5 mMphosphate buffer, pH 3.0

340 mm x 75 ~tm i.d.

250 mm active length

28

Alkylphenones

4~

Hydroquinone and its ethers

420 mm to the detector, 6 25~ t,,~~ r~

LiChrospher 100 RP 18, 5 ~tm Acetonitrile (50-70%)-20 mM 300 mm x 100 gm i.d. ammonium acetate

215 mm effective length 38

taO

TABLE 10.1 (continued)

Compound

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

320-500 gm i.d.

250-400 mm packed length

36

270 mm x 1O0 gm i.d.

200 mm length

39

200 mm length

39

40

Carbohydrates Malto-oligosaccharides

10 mM Tris + 15 mM boric Custom made macroporous polyacrylamide-poly(ethylene acid (pH 8.2) in acetonitrile0.1% acetic acid (5:95) glycol) matrix

p-Nitrophenyl-c~-D-glucopyra- Zorbax PSM 150, 5 ~tm, nosides and p-nitrophenylconverted to ODS ot-D-malto-oligosaccharides

5 mMNa2HPO4 (pH 6.0)water-acetonitrile (40-45:40-45:20-10)

p-Nitrophenyl-(x-D-glucosides, p-nitrophenyl glucopyranosides, separation of anomers

Zorbax PSM 150, 5 gm, converted to ODS

270 mm x 100 ~tm i.d. 5 mMNa2HPO4 (pH 6.0)water-acetonitrile (42.5-37.5:42.5-37.5:15-25) or 30 mM boric acid (pH 7.0)water-acetonitrile (45-42.5:45-42.5:15-10)

Glucuronides, aromatic

PRP- 1, 10-20 gm

Acetonitrile-2 mM ammonium acetate (pH 7.0) (5:95)

200 mm x 220 jam i.d.

Polyether ketone (PEEK) capillary

Aldopentoses (as 1-phenyl-3methyl-5-pyrazolone derivatives)

Nucleosil silica, in-column 3-aminopropylated or Devosil NH2, 3 ~tm

(25 mM HEPES-NaOH, pH 6.0)-acetonitrile (2" 1)

345 mm x 100 gm i.d.

250 mm effective length 41

,~ ~

Monosaccharides of glycopro- DaisoGel silica, in-column teins (as 1-phenyl-3-methylderivatized by 5-pyrazolone derivatives) octadecyltrimethoxysilane or dimethyloctadecyltrimethoxysilylpropylammonium chloride, 3 gm Glycosphingolipids

Porous ODSS, 5 ~tm or non-porous ODSS, 2 ~tm

2.5 mM Phosphate buffer with 345 mm x 100 gm i.d. 5 mM SDS (pH 7.0)acetonitrile (6:4) or 10 mM Tris.HC1 (pH 7.0)-acetonitrile (55:45)

250 mm effective length 41

270 mm x 100 gm i.d. Acetonitrile-methanolaqueous sodium borate, pH 9.40 (30:50:20), tetrahydrofuran-aqueous ammonium phosphate, pH 7.00 (80:20), tetrahydrofuran-methanol with 2 mM ammonium acetate (80:20) or tetrahydrofuranaqueous with sodium borate, pH 9.00 (80:20)

205 mm effective length 42

~,.~~ t~

r~

Carboxylic acids, simple esters and related compounds Folic acid, p-hydroxybenzoic acid, acetylsalicylic acid, nicotinic acid, thiourea, nicotinamide

Nucleosil 100 3-C 18, 3 gm

Methanol-20 mM disodium tetraborate, pH 8.5 (75:25)

150 mm (effecive length) x 100 gm i.d.

40

Prostaglandins and impurities

Spherisorb ODS-1, 3 ~tm

Acetonitrile-2 mMNa2HPO4, 300 mm x 50 gm i.d. pH 7.3 (75:25)

43

Prostaglandins and impurities

Zorbax SBC8, 3 ~tm

Acetonitrile-10 mM Na2HPO4, pH 9.9 (70:30)

400 mm x 50 gm i.d.

43

ta~

OO

TABLE 10.1 (continued)

taO OO bO

Compound

Stationary phase

2-Phenylpropionic acid, racemic mixture

Mobile phase

Capillary dimensions

Note

Ref.

trans-3-(3-pyridyl)acrylic acid 50 mMNaH2PO4, pH 4.65 coating molecular imprinting, 3 ~tm

435 mm x 100 ~tm i.d.

350 mm packed length, 44 chiral separation

Phthalate esters

Spherisorb ODS-1, 3 gm

Acetonitrile-50 m M Tris, pH 8.0 (80:20)

350 mm x 100 gm i.d.

250 mm packed length

45

Malonic acid, sulfate

TSK IC-Anion-SW, 5 gm

Methanol-3 m M phthalic acid, pH 6.8 (adjusted by hexamethylenediamine) (10:90)

289 mm x 50 ~tm i.d.

230 mm packed length

46

Triglycerides

CEC Hypersil C 18, 3 gm

50 mM Ammonium acetate in 350-500 mm x 100 acetonitrile-isopropanol- ngm i.d. hexane (57:38:5)

250-400 mm packed length

47, 48, 49

Alkyl benzoates

40% Ethylacrylate, 50% methacrylic acid, 10% lauryl methacrylate (custom made polymer), 0-3.75 wt. % polymer concentration

Acetonitrile-water (40:60) pH 11.3

500 mm x 50 ~tm i.d.

420 mm to the detector, 6 25~

Phthalate esters

Exsil C18, 1.5 ~tm

Acetonitrile-100 m M Tris, pH 8.6 (90"10)

260 mm x 75 gm i.d.

195 mm packed length

50

4~

Fatty acids and their phenacyl esters

Hypersil ODS, 3 ~tm

Triglycerides in corn oil

Acetonitrile-50 m M MES [2- 350 or 500 mm x 100 (N-morpholino)ethanesulfonic ~tm i.d. acid], pH 6 (90:10)

250 or 400 mm packed length

51

Hypersil ODS, 3 gm

Acetonitrile-isopropanoln-hexane (57:38:5) 50 m M ammonium acetate

400 mm x 100 gm i.d.

20~

52

Chlorinated alkyl phenoxypropanoates, etc.

Silica gel (Nucleosil, 5 ~tm) coated with Chirasil-Dex

Methanol-20 mM MES buffer, pH 6 (1:1, 2:3 or 7:3)

400 mm x 100 gm i.d.

250 mm effective length, pressuresupported chiral separation

53

Triglycerides in pharmaceutical formulation including testosterone phenyl propionate, testosterone propionate, testosterone isocaproate, testosterone decanoate

Hypersil ODS, 3 gm

Acetonitrile-isopropanoln-hexane (57:38:5) 50 m M ammonium acetate

250 mm x 100 ~tm i.d.

Retinyl esters (C 16, C 17 FA)

Hypersil C18, 3 gm

Lithium acetate in dimethyl formamide (2.5 mM)methanol (99:1)

310-360 mm x 180 gm i.d.

250-300 mm packed length

54

Fatty acids and derivatives in food products

CEC Hypersil C18, 3 ~tm

Acetonitrile-50 mM MES, pH 6.0 (90:10)

350-500 mm x 100 ~tm i.d.

250-400 mm packed length

48, 49

t~ |

4~

~,,~~

52

TABLE 10.1 (continued) 4~

Compound

Stationary phase

Fatty acid methyl esters (palmitoleic, oleic, eicosenoic and erucic acids)

GROM-SIL ODS-0 AB, 3 ~tm 40 mM Ammonium acetate (pH 9) in water-acetonitrile (50:50)

Thiourea, dimethyl phthalate, diethylphthalate, biphenyl, o-terphenyl

Spherisorb ODS-1, 3 gm

Benzoic acid derivatives RP (Spherisorb ODS-1) and (nitro- and bromo-, positional SAX (Spherisorb SAX) isomers of bromobenzoic acid) packed capillaries, 5 ~tm

Thiourea, methyl p-hydroxybenzoate, ethyl p-hydroxybenzoate, n-propyl p-hydroxybenzoate, n-butyl p-hydroxybenzoate, propranolol, biphenyl, dibucaine

Mobile phase

Acetonitrile-25 mM phosphate, 0.2% hexylamine, pH 2.5 (4:1)

Capillary dimensions

Note

250 mm packed length Coupling with x 100 lam i.d. coordination ion spray mass spectrometry; capillary electrochromatography and pressurized capillary electrochromatography 335 mm x 100 pm i.d.

310 mm x 50 ~tm i.d. 60% Acetonitrile in 2 mM phosphate buffer (pH 2.3) for ODS, 50% acetonitrile in 10 mM phosphate buffer (pH 2.2) for ion-exchange CEC

C18 phases compared (5 lam) Acetonitrile-50 mMTris.HC1, 305 mm x 100 pm i.d. pH 8 (60:40)

Ref. 55

250 mm effective length 56

100 mm packed length

57

200 mm effective length 58

r~

Thiourea, dimethyl phthalate, diethyl phthalate, biphenyl, o-terphenyl

ODS-1, 3 pm

Benzoates (methyl- to pentyl-) Grom-Sil ODS-0 ABI, 3 ~tm 4~ ta~

31

Acetonitrile-25 mM Tris.HC1, 250 or 200 mm pH 8 (80:20) effective length x 100 gm i.d.

r,.,~

Acetonitrile-5 mM sodium tetraborate, pH 9.0 (80:20)

370 mm x 22-50 gm i.d.

250 mm effective length 59

r~

Steroids

Aldosterone, dexamethasone, 13-estradiol, testosterone

Hypersil C 18, 3 pm or NPS ODS-2, 1.5 pm

Acetonitrile-20 m M sodium acetate, pH 5.0 (70:30) or acetonitrile-25 m M Tris (70:30)

320 mm x 100 gm or 335 mm x 100 gm or 260 mm x 100 pm i.d.

85-250 mm effective length; comparison of frits, double tapers and fritless arrangement

60

Adrenosterone, hydrocortisone, dexamethasone, fluocortolone

Apex ODS, 3 pm

Acetonitrile-5 mM ammonium acetate in water, gradient from 91:9 to 20:80

240 mm x 50 pm i.d.

160 mm packed length

61

Testosterone, 17-~- and 20-ot-hydroxyprogesterone, androstenedione, progesterone, norethindrone

Hypersil C 18, 3 pm

Acetonitrile-methanol-20 mM Tris-HC1, pH 8.0 (37.5:37.5:25)

350 mm x 100 gm i.d.

250 mm packed length

62

Tipredane and related compounds

Spherisorb ODS-1, 3 pm

Acetonitrile-50 mM Tris, pH 7.8 (80:20)

250 mm x 50 lam i.d.

Fluticasone and impurities

Hypersil C 18, 3 ~m; Spherisorb ODS-1, 3 ~tm

Acetonitrile-5 mM borate, 400 (or 600) mm x 50 pH 9.0 (80:20) or acetonitrile- pm i.d. 2-10 mMNa2HPO4, pH 8.3-9.9 (75-70:25-30) or acetonitrile-100 mM Tris, pH 9.3 (90:10)

63, 64 43, 65

OO

TABLE 10.1 (continued)

Compound

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

Triamcinolone, hydrocortisone, prednisolone, cortisone, methylprednisolone, betamethasone, dexamethasone, adrenosterone, fluocortolone, triamcinolone acetonide

Hypersil C 18, 3 ~tm

Acetonitrile-5 mM ammonium acetate, gradient from 17:83 to 38"62

420 mm x 50 pm i.d.

300 mm packed length

61, 66

Corticosteroids

Hypersil C 18, 3 pm

Acetonitrile-10 to 20 mM phosphoric acid, pH 2.1 (35-95:65-5) or acetonitrile10 mM Tris-HCl, pH 8 (35-95:65-5)

350 mm x 50/100 pm i.d.

250 mm packed length

67

Corticosterone, testosterone, androsten-3,17-dione, androstan-3,17-dione, pregnan-3,20-dione

Zorbax ODS, 6 pm

Acetonitrile-10 mM borate buffer, pH 8.0 (65:35)

176 m m x 50 p m i.d.

96 mm packed length

68

Aldosterone, hydrocortisone, testosterone

Hypersil C 18, 3 ~tm

Acetonitrile-4 mM sodium tetraborate, pH 8.0 (20:80)

250 mm x 50 ~tm i.d.

Digoxigenin, gitoxigen, cinobufatalin, digitoxigenin, cinobufagin, bufalin

Spherisorb ODS-1, 3 pm

Acetonitrile-4 mM sodium tetraborate, pH 8.0 (70:30) or acetonitrile-water + 0.1% formic acid (70:30)

500 mm x 50 ~tm i.d. or 450 mm x 100 ~tm i.d.

69 250 mm packed length

70 c~

4~ t.,a

Acetonitrile-1.6 mM sodium tetraborate, pH 9.25, 5 mM SDS (60:40)

320 mm x 100 ~tm i.d.

Hydrocortisone, testosterone, 17-~-methyltestosterone, progesterone

Chromspher, nonporous 1.5 ~tm

Estriol, hydrocortisone, estradiol, estrone, testosterone,

Zorbax ODS, 1.8 ~tm

Acetonitrile-0.8 mM sodium 320 mm x 50 jam i.d. tetraborate, 5 mM SDS (80:20)

Spherisorb ODS-1, 3 ~m

Acetonitrile-Tris, pH 7.8 (80:20)

Hydrocortisone, prednisolone, Poly(AMPS-co-IPAAm) hydrocortisone-21-acetate, hydrogel testosterone

100 mM Tris- 150 mM boric acid, pH 8.1

238 mm packed length

71 ~.,~~

~..~~

240 mm packed length

72

350 mm • 50 ~tm i.d.

75 mm packed length

73

650 mm x 75 ~tm i.d.

500 mm packed length

74

Hydrocortisone, prednisolone, Hypersil C18, 3 ~m betamethasone, betamethasone dipropionate, clobetasol butyrate, fluticasone propionate, clobetasone butyrate, betamethasone- 17-valerate

Acetonitrile-2 mM phosphate, 270 mm • 50 ~m i.d. pH 7.8 (80:20)

200 mm packed length

75

Steroids (neutral or conjugated) and their dansylhydrazine derivatives

Polyacrylic gel-based macroporous particles

Acetonitrile-water-240 mM ammonium formate buffer, pH 3 (55:40:5) (also gradient elution)

350 mm x 100 ~tm i.d.

250 mm active length, laser induced fuorescence detection or coupling with electrospray-ion-trap mass spectrometry

76

Hydrocortisone derivatives

Spherisorb ODS-1, 3 ~tm

Acetonitrile-5 mM phosphate buffer, pH 7 (60:40)

335 mm x 100 ~tm i.d.

165 mm packed length

26

17-ot-methyltestosterone, 4-pregnen-20c~-ol-3-one, progesterone De-esterified steroids, budesonine, steroid A

TABLE 10.1 (continued)

ta~

Compound

Stationary phase

Mobile phase

Capillary dimensions

Note

Dexamethasone, betamethasone 17-valerate, fluticasone propionate

Hypersil Duet C 18/SCX mixed mode, 3 gm

Acetonitrile-25 mM ammonium acetate, pH 4.0 (8O:2O)

400 mm x 1O0 pm i.d.

250 mm packed length, 77 coupling with microelectrospray-mass spectrometer

Ref.

Nitro and nitroso compounds

Nitrobenzene, 2,4-dinitroSpherisorb ODS-2, 3 pm toluene and other benzene derivatives along with phenol, phenyl propanol and thiourea N-nitrosodiethanolamine

Acetonitrile-4 mM Tris, pH 435 mm x 50 pm i.d. 9.2 (60:40) or gradient elution to 80:20

C 18 modified etched capillary 0.3 mol/1 Acetic acid containing T-aminobutyric acid (0.375 mol/1), pH 4.41

158 mm effective length 78

470 mm x 50 gm i.d.

250 mm effective length 79

330 mm x 1O0 pm i.d.

250 mm packed length, 80 stereoisomer separation

Amines

Fluvoxamine (primary amine) Hypersil ODS, Spherisorb ODS-1, 3 gm

Acetonitrile-phosphate buffer, pH 7.0 (60:40), 6 mM hexylamine

o~

Amino

acids

Dansylated amino acids

Polyacrylamide hydrogel (linear) column, 10 ~tm ethylenechlorotrifluoroethylene

10 m M Tris, 100 m M H3PO4 or water-acetonitrile (10:90) with TFA (0.15%)

Dansylated and non-dansylated amino acids

Dns-L-leucine molecular imprinting

Dansylated phenylalanine

420 mm x 100 gm i.d.

270 mm packed length

81

Acetonitrile-acetic acid-water 500 mm x 50 gm i.d. (90:5:5)

250 mm packed length, chiral separation

82

Dns-L-phenylalanine molecular imprinting

Acetonitrile-10 m M phosphate, pH 7 (10:1)

1 0 0 0 m m x 25 ~ m i.d.

850 mm packed length, chiral separation

83

Dansylated phenylalanine

Dns-L-phenylalanine molecular imprinting

Acetonitrile-100 m M acetate (80:20)

250 mm x 75 lam i.d.

250 mm packed length, chiral separation

10

DNP amino acids

[3-CD bonded silica particles, 5 pm

Methanol-10-20 m M triethylamine acetate buffer, pH 4.71 (10-25:90-80)

400 mm x 50 pm i.d.

210 mm packed length, chiral separation

84

PTH amino acids

Zorbax ODS, 3.5 pm

Acetonitrile-5 m M phosphate, 207 mm x 50 pm i.d. pH 7.55 (gradient from 30:70 to 60:40)

127 mm packed length

68

PTH amino acids

Monolithic packing of sintered Zorbax ODS, 6 pm

Acetonitrile-5 m M sodium phosphate, pH 7.5 (30:70)

330 mm x 75 pm i.d.

The sorbent was reoctadecylated; 230 mm packed length

85

PTH amino acids

Zorbax ODS, 3.5 gm

Gradient elution (A: 2 m M aqueous ammonium acetate, pH 7.0; B: 2 m M a m m o n i u m acetate in water-acetonitrile 1:9) from 30 to 90% B

1 5 0 m m x 75 p m i.d.

coupling with electrospray ionization and time-of-flight mass spectrometry

86

4~

t~

ta,9

TABLE 10.1 (continued)

Stationary phase

Mobile phase

Phenylalanine, tyrosine, phenylglycine, tryptophan, serine

L-Phenylalanine anilide molecular imprinting

Acetonitrile-acetic acid-water 500 mm x 75 gm i.d. (80:10:10)

250 mm packed length, 87 chiral separation

Phenylalanine, tyrosine, p-fluorophenylalanine, phenylglycine, phenylalanine amide, DOPA

Phenylalanine or phenylalanine amide molecular imprinting

Acetonitrile-acetic acid-water 400 mm x 75 pm i.d. (90:5"5)

200 mm packed length, 88 chiral separation

N-derivatized amino acids (DNZ, FMOC), their enantiomers

Silica gel (Kromasil) modified with a basic tert.-butyl carbamoyl quinine chiral selector, 3 ~tm

Acetonitrile or methanol-50 mM acetic acid (80:20), pH 6 (titrated with triethylamine or ammonia)

335 mm x 100 (or 75) pm i.d.

Acetonitrile-5 mMphosphate buffer, 7 (80:20)

300 mm (active length) Enantiomer separation x 100 pm i.d.

N-(3,5-dinitrobenzoyl)leucine Monoliths (2-hydroxyethyl diallyl amide methacrylate carbamate copolymerized with four different chiral selectors)

Capillary dimensions

Note

Ref.

Compound

250 mm packed length, 89 chiral separation by weak anion-exchange type chiral stationary phase 90

N-derivatized amino acids, their enantiomers

CEC-Hypersil, 3 ~tm

r

| 4a.

Non-aqueous: methanol, 20 mM ammonium acetate, 80 mM acetic acid, 5 mM tert.butyl carbamoyl quinine; Aqueous: methanol-100 mM ammonium acetate (80:20), pH 6.0, 5 mM tert.-butyl carbamoyl quinine

335 mm x 100 gm i.d.

250 mm effective 91 length, chiral separation

,.,,, ~ t% ~,,~~

Phenylthiohydantoin-amino acids

Chromspher ODS, 1.5 gm

Phosphate buffer pH 7.2, 5 mM SDS, 5% acetonitrile and 5% tetrahydrofuran

325 mm x 100 lam i.d.

240 mm packed length

Dansylated amino acids

]3-Cyclodextrin-bonded positively charged polyacrylamide gel

200 mM Tris, 300 mMboric acid buffer, pH 8.1

550-700 mm x 75 jam i.d.

350 mm effective 93 length, chiral separation

Benzyloxycarbonyl, N-(3,5dinitrobenzyloxycarbonyl), 9-fluorenylmethoxycarbonyl, benzoyl, acetyl and N-(2,4dinitrophenyl) derivatized amino acids and profens

WAX (weak anion-exchange) Acetonitrile-methanol (80"20)+400 mM acetic type CSP (tert.acid+4 mM triethylamine butylcarbamoylquinine as chiral selector on Hypersil silica gel), 3 gm

335 mm x 100 ~tm i.d.

250 mm effective 94 length, chiral separation

Tryptophan and dinitrobenzoyl leucine enantiomers

Chirobiotic T (teicoplanin), 5 gm

Acetonitrile-2 mM (or 5 mM) 330 mm x 100 gm i.d. Na2HPO4, pH 7, 3.5 or 2.3

245 mm effective 95 length, chiral separation

DNZ-leucine, DNB-leucine, Fmoc-Leucine

Silica based (Kromasil 1005gm, Hypersil 120-3gm, Micra NPS-1.5~tm) weak anion-exchange-type chiral stationary phases

Acetonitrile or methanol-100 mM MES (80:20)

335 mm x 75 or 100 gm i.d.

92

250 mm effective 96 length, chiral separation

TABLE 10.1 (continued)

Compound

to

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

60 mm packed length

97

Peptides, proteins

Peptide map, cytochrome c tryptic digest

ODS, 1.5 ~m, 100 A pore size A: 0.1% TFA in water, pH 2, (Polymicro Technologies) B" as A containing 50% (by vol.) acetonitrile; gradient 0-100% B over 10 min

200 mm x 100 ~tm i.d.

Nonapeptide

Nucleosil 100-5C8, 3 gm

Methanol-4 mM ammonium acetate, pH 8.0 (40:60)

250 mm x 220 ~tm i.d.

98

Cytochrome c, bovine, tryptic Vydac C 18, 3 ~tm digest, chicken albumin, tryptic digest

Acetonitrile-0.07% TFA in water (gradient from 0:100 to 40:60)

60 mm x 180 ~tm i.d.

99

Vydac C 18, 3 ~tm Bradykinin, angiotensin II, angiotensin I, Met-enkephalinArg-Phe, neurotensin

0.07% TFA in wateracetonitrile (75:25)

120 mm x 180 ~tm i.d.

99

100

[~-Lactoglobulin, bovine

Vydac C18, 3 ~tm

0.04% TFA in acetonitrilewater (gradient from 0:100 to 75:25)

60 mm x 180 ~tm i.d.

Tetrapeptides, pentapeptides

Gigaporous PLSCX, 8 gm

Acetonitrile-25 mM phosphate, pH 3.5 (40:60)

3 7 0 m m x 180~tmi.d.

280 mm packed length

101

Tyrosine-containing peptides

Polyacrylamide poly(ethylene Acetonitrile-10 mM Tris, glycol) macroporous packing 15 mMboric acid, pH 8.2 (47:53)

250 mm x 100 gm i.d.

205 mm packed length

36

Lysozyme, angiotensins r r~

C 18, etched and modified column; diol open tubular column

30 m M phosphate, pH 2.14, 30 m M citric acid, pH 3-25 m M [3-alanine, 30 mM acetic acid, pH 4.14

450 mm x 50 pm i.d.

250 mm packed length

102 ~,~~ t~

4~ too

Cytochrome c, lysozyme, myoglobin, ribonuclease A

C 18, etched and modified column; diol open tubular column

30 m M phosphate, pH 2.14, 30 m M citric acid, pH 3-25 m M [3-alanine, 30 mM acetic acid, pH 4.14

450 mm x 50 pm i.d.

250 mm packed length

103

Enkephalin methylester, enkephalin amide

Gromsil ODS-2, 1.5 pm

Acetonitrile-0.07 ml/1 TFA in water (80:20)

400 mm x 100 pm i.d.

230 mm packed length

104

Tetrapeptide, C- and N-protected

Spherisorb ODS-1, 3 ~tm

Acetonitrile-50 mM Tris, pH 7.8 (80:20)

335 mm x 50 gm i.d.

250 mm packed length

63, 64

Mixture of peptides and proteins (lysozyme, angiotensin I and III, bradykinin, ribonuclease A)

C 18-modified etched capillary 30 m M citric acid-24.5 m M [3-alanine, pH 3.0

450 mm x 50 lam i.d.

250 mm effective length 105

Lysozymes

Diol or C 18-modified etched capillaries

30 mMPhosphate, pH 2.14 or 450 mm x 50 ~tm i.d. 30 m M citric acid-19 m M Tris, pH 3.0 for C 18 capillary or 30 m M acetic acid-25 m M [3-alanine, pH 4.41 for diol capillary

250 mm effective length 106

Angiotensins

Diol or C 18-modified etched capillaries

30 mMphosphate, pH 2.14

250 mm effective length 106

450 mm x 50 pm i.d.

TABLE 10.1 (continued)

Mobile phase

Capillary dimensions

Compound

Stationary phase

Carbonic anhydrase, myoglobin, amylase, trypsinogen, ribonuclease A, chymotrypsinogen A, mesityl oxide

Polyaspartic acid immobilized Salt or pH gradient elution in 400 mm x 75 pm i.d. phosphate buffer (pH 6.0) or on the capillary wall isocratic elution with 100 mM NaC1 in 10 mM phosphate (pH 6-8)

Peptide mapping ovalbumin, tryptic digest

C 18 COMOSS microfabricated column

Acetonitrile-10 mM potassium phosphate, pH 9.0 (1:3)

Oxytocin, desmopressin, cerbetocin and related synthetic peptides

CEC Hypersil C8, CEC Hypersil C 18, 5 gm, Spherisorb mixed mode C 18/SCX phase 3 pm

Triethylamine buffers, pH 3.0 335 mm • 100 pm i.d. adjusted with phosphoric acid with different proportion of acetonitrile

Basic proteins and peptides

0, 10, 20, 30% Acetonitrile in 470 mm x 20 pm i.d. PLOT (porous-layer open 20 mM aqueous sodium tubular) column, i.e., fused ilica capillary with 2 gm thick phosphate, pH 2.5 polymer layer (highly crosslinked in situ polymerized vinylbenzyl chloride and divinylbenzene)

1.5 pm wide, 10 ~tm deep rectangular channel, 45 mm long

Note

Ref.

250 mm active length

107

700 s needed for run completion

108

250 mm packed length

109

400 mm effective length 110

Recombinant human growth hormone, tryptic digests o~

Vydac 218TPB5 (C18), 5 gm, A: 0.1% TFA/water, B: 300 A 0.09% TFA/acetonitrile, gradient from 0 to 60% B

250 mm x 100 gm i.d.

4~

Electrically assisted 111 capillary HPLC, coupling with electrospray ionizationmass spectrometry

Cytochrome c

Capillary etched with liquid crystals (cholesteryl or cyanopentoxy modified)

60 m M Citric acid and 50 mM 700 or 500 mm x 50 13-alanine, pH 3.00 gm i.d

450 or 250 mm effective length, open tubular CEC

112

Lysozymes, cytochromes c, aspartame

Silica capillary was etched and chemically (C18) modified

pH 3.7

515 or 580 mm x 20 l.tm i.d.

220 or 260 mm effective length, open tubular electrochromatography

113

Ribonuclease A, insulin, c~-lactalbumin, myoglobin

Methacrylic monolith with tertiary amino functions

30% Acetonitrile in 60 m M aqueous sodium phosphate, pH 2.5

390 mm x 50 lam i.d.

290 mm effective length 114

Hypersil ODS, 3 ~tm

30% Methanol, 10% acetonitrile in 5 m M aqueous ammonium acetate

250 mm • 25 gm i.d.

CEC/MS (quadrupole) coupling

Acetonitrile-5 m M acetic acid, 2-3 mM triethylamine, pH 5.0 buffer (8:92)

250 mm • 100 ~tm i.d.

effective column length 116 not specified, 20-26~

Nucleotides, nucleosides Adenosine-styrene oxide adducts, neutral and positively charged; inosine-styrene oxide adducts, neutral and positively charged

CEC Hypersil C 18, 3 gm Nucleosides (adenosine, cytidine, guanosine,thymidine and uridine)

115

-m,

taO

TABLE 10.1 (continued)

O',

Compound

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

Purine and pyrimidine bases and their nucleosides (adenosine, adenine, cytidine, cytosine, guanosine, guanine, inosine, thymine, uridine, uracil)

ODSS (octadecyl sulfonated silica), 10 gm

Acetonitrile-4.8 m M sodium acetate, pH 4.5 (40:60)

270 mm x 20.5 gm i.d

205 mm effective length 117

AMP, ADP and ATP

Nucleosil 100-C 18, 5 gm

Methanol-2 m M dibutylamine, pH 5.0 (10:90)

200 mm x 220 gm i.d.

118

d(GATGCATAGG-OH) and by-products, dC3-dC 11

Gromsil ODS-2, 5 gm

Acetonitrile-10 m M triethylamine acetate (12:88), acetonitrile-10 m M ammonium acetate, gradient from 0"100 to 10:90

500 mm x 50-100 gm i.d.

300 mm packed length

119

Thymine, cytosine, adenine, guanine, adenosine

Capillary etched with liquid crystals (cholesteryl or cyanopentoxy modified)

60 mM Phosphoric acid and 38 m M Tris, pH 2.14

700 or 500 mm x 50 ~tm i.d

450 or 250 mm effective length, open tubular CEC

112

Nucleosides (adenosine, cytidine, guanosine, uridine, thymidine)

Hypersil Phenyl, 3 ~tm, 120 A Acetic acid-ammonia, pH 5acetonitrile (95:5)

270 mm x 75 ~tm i.d

200 or 70 mm bed length

120

Alkaloids

Morphine alkaloids 4~

~.,~~

Nucleosil 100-C 18, 5 ~tm

Acetonitrile-2 m M ammonium acetate (40:60)

150 mm x 220 gm i.d.

118

YMC C30, 5 gm

N,N-Dimethylformamideacetonitrile-methanol (29:70"1) with 2.5 m M lithium acetate

360 mm x 180 ~tm i.d.

300 mm effective length 121

Macrocyclic antibiotics (lactone, from Streptomyces $541)

CEC Hypersil, 3 gm

Acetonitrile-5 m M borate, pH 9 (80:20)

430 mm x 50 gm i.d.

122

Tetracyclines

Etched and modified C 18 open tubular column

Methanol-30 m M citrate containing 24.5 m M 13alanine, pH 3.0 (40:60)

250 mm x 50 gm i.d.

123, 124

Cephalosporins (cefuroxime, axetil)

CEC Hypersil, 3 lam

Acetonitrile-5 m M borate, pH 9 (80:20)

400 mm x 50 lam i.d.

Cephalosporins (cefuroxime, axetil)

Spherisorb ODS-1, 3 gm

Acetonitrile-10 m M Na2HPO4, pH 9.5 (80:20)

400 mm x 50 lam i.d.

Tetracyclines

C 18-modified etched capillary 30 m M phosphate-19 mM Tris, pH 2.14, 40% methanol

Vitamins

|

4~

Retinyl esters

Antibiotics

450 mm x 50 gm i.d.

For MS assay

43 43

250 mm effective length 105

--..I

TABLE 10.1 (continued)

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

Triazine herbicides

Hypersil C 18, 3 pm, Hypersil C8, 3 gm, Spherisorb C6/SCX, 3 ~tm

Acetonitrile-25 m M sodium acetate, pH 8 (50:50)

335 mm x 100 pm i.d.

250 mm packed length

30

Primicarb

Spherisorb ODS-1, 3 pm

Acetonitrile-20 m M aqueous Tris, pH 9.0 (60:40)

330 mm x 50 pm i.d.

245 mm packed length

50

Pesticides

Hypersil ODS C 18, 3 ~tm

Acetonitrile-25 m M H E P E S buffer, pH 7.55 (75:25)

490 mm x 100 pm i.d.

400 mm effective length 125

Urea herbicides

Zorbax ODS, 5 ~tm

270 mm x 100 pm i.d. Methanol-5 mMNaH2PO4, pH 6.0 (70:30) or acetonitrile5 mMNaH2PO4, pH 6.0

200 mm effective 126 length; preconcentration

Compound Pesticides

(5o:5o) 245 mm packed length

127

335 mm • 100 ~tm i.d. Acetonitrile-20 m M sodium dihydrogen phosphate, pH 4.0 (60:40)

250 mm packed length

128

Acetonitrile-13 m M T F A , pH 3.5 (60:40)

252 mm packed length

60

Primicarb

CEC Hypersil C18, 3 ~tm

Acetonitrile-5 m M aqueous Tris, pH 8.6 (42:58, 12:88)

Cinosulfuron

Synchropak (C 18), non-endpacked

Cinosulforon and by-products

Gromsil ODS-O AB, 3 gm

330 mm x 50 pm i.d.

337 mm x 75 pm i.d.

c~

Polychlorinated dibenzo-pdioxins

4~ i

4~

ODS C 18, 3 pm

Spherisorb ODS-1, 3 gm, Herbicides (desisopropylaimmobilized trazine, desethylatrazine, simazine, cyanazine, atrazine, sebutylazine, propazine, terbutylazine, 2-hydroxyterbutylazine, 2-hydroxyatrazine)

Acetonitrile-25 mM Tris, pU 8.5 (75-80"25-20)

450 mm x 100 ~tm i.d.

Acetonitrile-25 mM Tris.HCl buffer, pH 8 (56:44)

335 mm x 100 pm i.d.

250 mm effective 31 length, influence of the packing immobilization

Channel depth 5.2 lam

length not specified, chip arrangement

130

Polyether ketone (PEEK) capillary

40

340 mm packed length

129 r..~~

Dyes Coumarin dyes (C440, C450, C460, C480)

Octadecylsilane or 10 mM Borate with 29% octadecyldiisobutyl(dimethyl- acetonitrile, pH 8.4 amino)silane surface modified capillary

Food dyes (E 102, E 110, E122, E123, E124)

Nucleosil 10-5C 18, 5 gm

Textile dyes (azo- and anthra- Hypersil C 18, 3 gm quinone compounds) Carotenoid isomers

Methanol-10 mM ammonium 200 mm x 250 pm i.d. acetate, pH 8.5 (20:80) Acetonitrile-4 mM borate, pH 8.0 (80:20)

Polymeric C30 (Rainin 30 nm Isocratic: methanol-methylpore size or ProntoSIL 30 nm tert.-butyl ether-buffer pore size), 3 pm (35:60:5), acetone-buffer (99:1, 95:5, 90:10, 85:15) when buffer is 1 mMborate buffer. Gradient elution: acetone-1 mM sodium borate buffer, gradient from 80:10 to 99:1

25 mm x 75 pm i.d. 250 mm x 98 gm i.d.

131 330 mm total length

132

~..,~

4~ O

TABLE 10.1 (continued)

Compound

Drugs (for steroid drugs s e e

Stationary phase also

Ref.

Mobile phase

Capillary dimensions

Note

Acetonitrile-buffer, pH 2.5 (75"25)

340 mm x 100 lam i.d.

250 mm to the detector, 133 temp. 25~ comparison with HPLC and CZE

Steroids)

Drugs of abuse: amphetamine, CEC Hypersil C8, 3 pm metaphetamine, procaine, cocaine, heroin, quinine, noscapine and thiourea Drugs of abuse: phenobarbital, testosterone, cannabinol, testosterone propionate, A9-tetracannabinol, A9-tetracannabinolic acid

CEC Hypersil C8, 3 pm

Acetonitrile-25 mM phosphate buffer, pH 2.5, gradient from 60:40 to 75:25

340 mm x 100 pm i.d.

250 mm to the detector, 133 comparison with HPLC and CZE

Dexamethasone, hydrocortisone, prednisolone acetate, cortisone acetate

Hypersil C8, 3 lam

Acetonitrile-5 mM sodium tetraborate, pH 9.0 (80:20)

370 mm x 50 pm i.d.

190 mm packed length

Hydrocortisone, hydrocortisone- 17-valerate, hydrocortisone- 17-butyrate, betamethasone- 17-valerate, clobetasol- 17-propionate and preservatives (parabenes, benzyl alcohol, sorbic acid, chlorocresol)

Innovatech SCX/C 18, Hypersil ODS-1, ODS-2, 3 pm

250 mm x 50 ~tm i.d. Acetonitrile-50 mM sodium dihydrogen phosphate, pH 3.5 (30:70)

134

135

"x

4~

Acidic and basic drugs analysed simultaneously (amphetamine, metamphetamine, procaine, cocaine, heroin, quinine, noscapine)

CEC Hypersil C8

Acetonitrile-50 mM phosphate buffer, pH 2.5 (75:25) (75, 60, 45 and 30% acetonitrile also tested)

340 mm x 100 ~tm i.d.

250 mm to the detector, 136 20~

~,~~ r~

ta, a |

4~

~,~~

Acidic and basic drugs CEC Hypersil C8 (phenobarbital, diazepam, methaqualone, testosterone, cannabinol, testosterone propionate, 9-tetrahydrocannabinol, 9-tetrahydrocannabinolic acid)

Acetonitrile-50 mM 340 mm x 100 ~m i.d. phosphate buffer, pH 2.5 (60:40) with 2 gl/ml hexylamine for 1 min, then linear gradient to acetonitrile25 mM phosphate buffer with 2 gl/ml hexylamine (75:25)

250 mm to the detector, 136 stepped gradient, comparison with MECC

Acidic and basic drugs (see above)

CEC Hypersil C8

Acetonitrile-phosphate 340 mm x 100 gm i.d. buffer, pH 2.0 with 3.4 gl/ml hexylamine (2:98) for 20 min, then linear gradient to acetonitrile-phosphate buffer with 3.4 lal/ml hexylamine (65:35) and hold for 10 min

250 mm to the detector, 136 complex stepped gradient

Tripredane and related compounds

Spherisorb ODS-1, 3 lam

Acetonitrile-50 mM Tris buffer, pH 7.8 (80:20)

Barbital, butethal, phenobarbi- Hypersil C8, Hypersil C 18, tal, amylobarbital, secoHypersil phenyl, 3 tam barbital, hexobarbital

250 mm x 50 gm i.d.

(a) Acetonitrile-50 mM MES, 350 mm x 100 ~tm i.d. pH 6.1-water (60-40:20:2040); (b) methanol or acetonitrile50 mM phosphate, pH 4.5water (40-50:20:40-30)

63 250 mm packed length

137, 138

TABLE 10.1 (continued)

O b,.)

Mobile phase

Capillary dimensions

Stationary phase

Bendroflumethiazide

Spherisorb ODS-1, Spherisorb Acetonitrile-10-50 mM 500-550 mm • 50 ~tm 260-350 mm to the detector SCX, 3 ~tm NaH2PO4, pH 3.5-9.8 (70:30) i.d.

Barbiturates

Permethyl-[3-cyclodextrinMethanol-5 mM phosphate modified silica gel (Nucleosil, buffer, pH 7.0 (1:4) 5 ~tm)

Chlorthalidone, hydroflumethiazide, bendroflumethiazide, bumetanide

Spherisorb ODS- 1

Acetonitrile-5 mMNaH2PO4, 330 mm x 100 pm i.d. pH 2.3 (gradient from 40:60 to 60:40)

Hydroflumethiazide, methylchlothiazide, metolazone, epitizide, bendrofluazide

Hypersil C 18, 3 ~tm

Acetonitrile-5 mM ammonium acetate in water, gradient from 50:50 to 20:80

Bendroflumethiazide, nortriptyline, chlomipramine, methdilazine, imipramine, desipramine

Spherisorb ODS-1, 3 ~tm

550 mm x 50 ~tm i.d. Acetonitrile-10-50 mM Na2HPO4, pH 5.7-9.8 (70:30)

350 mm packed length

65

Bendroflumethiazide, nortriptyline, chlomipramine, methdilazine, imipramine, desipramine

Spherisorb SCX, 3 gm

Acetonitrile-50 mM Na2HPO4, pH 3.5 (70:30)

500 mm x 50 ~tm i.d.

260 mm packed length

65

400 mm x 100 gm i.d.

Note

Ref.

Compound

65

235 mm effective length, chiral separation, pressuresupported CEC

139

250 mm to the detector

140

61, 66

460 mm x 50 ~tm i.d.

Nitrazepam, diazepam

4~

Cloxazolam, nitrazepam, clotiazepam, diazepam

460 mm x 50 gm i.d.

Hypersil C 18, 3 jam

5 mM Ammonium acetate in acetonitrile-water, gradient from 50:50 to 80:20

Etched cholesterol modified open tubular column

Acetonitrile-10 mMTris-HC1, 700 mm x 50-75 gm pH 7.3 (0-20:100-80) i.d.

66 r..~,

550 mm packed length

141

r~

|

4~

Nonsteroidal antinflammatory CEC Hypersil C18/SCX, 3 ~tm Acetonitrile-50 mM drugs (ketoprofen, naproxen, Na2HPO4-water (60:20:20) flurbiprofen, indomethacin, ibuprofen) Bumetanide, flurbiprofen, p-hydroxybenzoic acid

Hypersil C 18 or Hypersil C 18/SCX, 3 gm

Acetaminophen, caffeine, ace- GROM-SIL1 100 ODS-0 AB, tylsalicylic acid 5 gm (Thomapyrin |

Antiviral drugs (suramin)

Nucleosil 100 C 18, 5 ~tm

Bare silica Micra, 3 ~tm Amino group containing drugs (codeine phosphate, ephedrine hydrochloride, thebaine, berberine hydrochloride, jatrorrizine hydrochloride, cocaine hydrochloride)

Acetonitrile-50 mM Na2HPO4, pH 2.3-2.5-water (40:20:40)

210 mm x 50 ~tm i.d.

138

210-230 mm x 50 ~tm i.d.

63, 138

200 mm x 250 ~tm i.d. 2 mM Borate in D 2 0 deuterated acetonitrile (80:20) or 1 mMborate in D 2 0 deuterated acetonitrile (gradient from 100:0 to 70:30) Methanol-2 mM dibutylamine, pH 5.0 (10:90)

200 mm x 220 gm i.d.

Acetonitrile-5-20 mM TrisHC1, pH 7.5-10 (90:35-10)

270 mm x 65-75 ~tm i.d.

coupling with NMR

142

118 200 mm packed length

143

4~ O

4~

TABLE 10.1 (continued)

Compound

Stationary phase

Mobile phase

Capillary dimensions

Isradepin and and byproducts

Hypersil C18, 3 ktm

Acetonitrile-2 mM sodium tetraborate, pH 8.7 (80:20)

143 mm x 50 ktm i.d.

144

Methanol-5 mM borate buffer, pH 8.5 (60:40)

260 mm x 50 ~m i.d.

145

Methanol-4 mM ammonium acetate, pH 5 (40:60)

235 mm x 220 ~tm i.d.

98

NPS ODS-2, 1.5 ~tm Cardiac glycosides (digoxigenin, digoxin, digitoxigenin)

Acetonitrile-25 mM Tris, pH 8 (50:50) made 0.3% to SDS

260 mm x 100 ~tm i.d.

Nucleosil 5C8, 3 gm 2-Phenylethylamine derivatives (epinephrine, DOPA, 2-amino-3-hydroxy-3-phenylpropanol, ephedrine)

Methanol-5 mMNaH2PO4, pH 3.1 (60:40)

155 mm • 100 jam i.d.

Antiepileptic drugs (ethosuc- Spherisorb ODS-1, 3 ~tm cinimide, primidon, carbamazepine- 10,11-diol, carbamazepine- 10,11-epoxide, carbamazepine, phenytoin Sulfanilamide, sulfaflurazol, sulfadicramide

Nucleosil 100-5C8, 3 gm

Note

245 mm packed length

Ref.

60

145

c~

Chiral drugs and chiral compounds (clomipramine, ~ chlorodiazepoxide, diazepam, .~ temazepam, doxepine, terbu"~" taline, clenbuterol, homota~ k atropine, tropicamide, ,~ 3,5-dinitrobenzoyl alanine methyl ester) o~

450 mm x 50 gm i.d. Silanized/hydrosilylated inner Phosphate pH 2.14, Tris surface of the capillary pH 3.0, citric acid + 13-alanine pH 3.7, lactic acid + 13-alanine pH 4.41, ,/-aminobutyric acid pH 6.0 (38-60 mM)

PM-13-CD, 5 gm Barbiturates (mephobarbital, hexobarbital, pentobarbital, 1-methyl-5-(2-propyl)-5(n-propyl)barbituric acid, 5-ethyl- 1-methyl-5-(n-propyl) barbituric acid), benzoin, c~methyl-ot-phenylsuccinimide, gluthethimide, methylthiohydantoin-proline, methyl mandelate

Methanol-5 mM phosphate, pH 7 (20:80)

400 mm x 100 9m i.d.

250 mm effective length, open tubular chromatography, chiral separations

146 .,,,o

~o r~

235 mm packed length, chiral separation

147

210 mm packed length, 84 chiral separation

Hexobarbital

13-CD, 5 pm

Acetonitrile-4 mM phosphate, 400 mm x 50 ~tm i.d. pH 6.8 (5:95), methanol-5 mM triethylamine acetate, pH 4.71 (15:85)

Chlorthalidone

Spherisorb $30DS-1, 3 gm

10 mM HP-I3-CD in acetonitrile-5 mM phosphate, pH 6.5 (20:80)

520 mm x 50 ~tm i.d.

262 mm packed length, chiral separation

148

Chlorthalidone

Hydroxypropyl-[3-CD bonded Acetonitrile-5 mMphosphate, 580 mm x 50 gm i.d. silica, 5 ~tm pH 6.5 (25:75)

272 mm packed length, chiral separation

148

TABLE 10.1 (continued)

Compound

Stationary phase

Mianserin

Hydroxypropyl-13-CD bonded Acetonitrile-10 mM silica, 5 pm phosphate, pH 7.5 (50:50)

Ibuprofen, cicloprofen, 1-phenylethanol, 1,1'binaphthyl-2,2'-dihydrogenphosphate, flurbiprofen, carprofen, etodolac, warfarin, hexobarbital

Chirasil-Dex coated column, 0.2 lam

Alkylated barbituric acids, 6chloro-c~-methylcarbazole-2acetic acid, 1-phenylethanol, 1-phenyl- 1-propanol, otmethy 1-2,3,4,5,6-pentafluo robenzyl alcohol, 1-(2naphthyl)-ethanol,

Mobile phase

Capillary dimensions

Note

Ref.

550 mm x 50 lam i.d.

219 mm packed length, chiral separation

148

20 mM Borate-phosphate, pH 800 mm x 50 pm i.d. 7, or 20 mM Tris-HC1, pH 7

Chiral separation

149, 147, 150, 151

Chirasil-Dex coated column, 0.15 pm

Acetonotrile-20 mM borate700-980 mm x 50 ~tm phosphate buffer, pH 7 i.d., or 620 mm • (10:90) or methanol-20 mM 25 gm i.d. borate-phosphate buffer, pH 7 (38-3:62-97)

600-880 mm packed 152 length, chiral separation or 500 mm packed length, chiral separation

ODS C18, 5 gm

20 mM Sodium phosphate, pH 3.0-12 mM 13-CD-4-5 mM sodium 1-heptanesulfonate

170-230 mm packed 153 length, chiral separation

1-(p-biphenyl)-ethanol Salsolinol

200-290 mm x 75 gm i.d.

c~

Hexobarbital, pentobarbital, isofamide, cyclophophamide, disopyramide

c~1-Acid glycoprotein-bonded 1-Propanol-5 mM phosphate, silica, 5 ~tm pH 6.5 (2:98) or 2-propanol2 mM phosphate, pH 5.5 (2:98) or 2-propanol-4 mM phosphate, pH 6.5 (2:98)

420 mm x 50 gm i.d.

170 mm packed length, chiral separation

154

Temazepam, benzoin

Human serum albuminbonded silica, 7 gm

2-Propanol-4 mM phosphate, pH 7 (5-7.5:95-92.5)

420 mm x 50 lam i.d.

170 mm packed length, chiral separation

155

Hexobarbital, warfarin

Vancomycin-coated silica, 3 lam

Acetonitrile-0.1% triethylamine acetate, pH 4 (80:20) or acetonitrile-0.1% triethylamine acetate, pH 5 (2O:8O)

500 mm x 1O0 gm i.d.

400 mm packed length, chiral separation

156

Glutethimide, 1-(9-anthryl)2,2,2-trifluoro-ethanol, mephobarbital, aminoglutethimide

3,5-dimethylphenylcarbamoyl Acetonitrile-40 mM cellulose and p-methylphosphate, pH 7 (15-40:85benzoyl cellulose coated 6O) column, 0.25 gm

570 mm x 50 gm i.d.

500 mm packed length, chiral separation

157

Acebutolol, alprenolol, atenolol, metoprolol, pindolol, prenalterol, propranolol

(R)-propranolol molecular imprinting phase

Acetonitrile-100 mM phosphoric acid and triethanolamine to pH 3.0 (8O:2O)

350-900 mm x 75 gm i.d.

265-815 mm packed 158 length, chiral separation

Propranolol, metoprolol

(R)-propranolol or (S)metoprolol molecular imprinting phase

Acetonitrile-2-4 mM acetate, pH 3.0 (80:20)

350 mm x 75 lam i.d.

265 mm packed length, chiral separation

4~ t~ !

4~

159

4~

TABLE 10.1 (continued)

Compound

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

Terbutaline, benzoin

Charged polyacrylamide gel

50 mg/ml Poly [3-cyclodextrin, 50 mg/ml carboxymethyl ~-cyclodextrin, 5% Tween 20, 200 mM Tris, 300 mMboric acid buffer, pH 9.0

700 mm x 75 pm i.d.

350 mm effective 160 length, chiral separation

Terbutaline, propranolol, benzoin

[3-Cyclodextrin-bonded charged polyacrylamide gel

200 mM Tris, 300 mM boric acid buffer, pH 9.0

700 mm x 75 pm i.d.

161 350 mm effective length, chiral separation

Benzodiazepines

Electropak phenyl, 3 pm

Tris.HC1, pH 8-acetonitrile (40:60)

470 mm x 75 pm i.d.

400 mm packed length

Benzoin acetate, methylbenzoin, Tr6ger's base, trans-stilbene oxide and 1, l'-binaphthyl-2,2'-diol

Aminopropyl silica gel (LiChrospher 1000, 5 gm) coated with helically chiral poly(diphenyl-2pyridylmethyl methacrylate) (PDPM)

Methanolic solution of ammonium acetate (2.5 mM, pH 4.5)

300 mm x 1O0 pm i.d.

163 200 mm effective length, chiral separation

Mephobarbital, hexobarbital, MTH proline, methyls of mecoprop, diclofop and fenoxaprop, barbiturates, chlorinated alkyl phenoxypropanoates, etc.

Silica gel (Nucleosil, 5 gin) coated with Chirasil-Dex

Methanol-20 mM MES buffer, pH 6 (1"1, 2:3 or 7:3)

290 (400) mm x 1O0 lam i.d.

200 (250) mm effective 53 length, pressuresupported chiral separation

162

o~

"x I%

4~

Opiate drugs

C 18, 1.5 ~tm

Thalidomide and its metabolites

LiChrospher 100 RP- 18, 5 pm 5 mM Acetonitrileammomium acetate, pH 6 (60:40)

too

10mMTris, 5mMSDS, 20% 280 mm x 75 tam i.d. acetonitrile, pH 8.3 330 mm x 100 lam i.d.

150mmeffectivelength 164 245 mm packed length, chiral separation

165 r~

Neutral polar pharmaceuticals Reversed-phase non-porous silica (NPS C-18, NPS ODS1, NPS PolyEncap B 1, NPS TAS- 1), 1.5 ~tm

Acetonitrile-2 mMphosphate, 335 mm x 75 pm i.d. pH 3.0 (60:40, 50:50 or 40:60)

250 mm effective length; comparison of various 1.5 lam nonporous phases

166

Benzodiazepines (temazepam, Capillary etched with liquid oxazepam, clonazepam, diaze- crystals (cholesteryl) pam, nitrazepam)

60 mM Phosphoric acid and 38 mMTris, pH 2.14

700 mm x 50 pm i.d

450 mm effective length, open tubular CEC

112

Theophylline, caffeine, acetaminophen, hydrochlorothiazide, 13-hydroxyethyltheophylline, phenylbutazone and theobromine

Silica EP-75-20-3-Si, 3 ~tm

Isopropanol-hexane-1 mM Tris (52:40:8, pH 8)

270 mm x 75 pm i.d.

200 mm effective length 167

Thalidomide and its metabolites

Aminopropyl silica coated Methanol-ethanol (75:25) with Chiralpak AD and/or OD containing 2.5 mM ammomium acetate

335 mm x 100 gm i.d.

250 mm effective 168 length, chiral separation

Clenbuterol, salbutamol, methadone, polynuclear aromatic hydrocarbons

MOS Hypersil, 3 gm

25 mM Ammonium acetate 160 m m x 75 p m i.d. buffer, pH 5.0, in acetonitrilewater (90:10) or 2 mM Tris, pH 8.0 in acetonitrile-water (80:20)

169

TABLE 10.1 (continued)

Compound

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

Indapamide, lormetazepam, transstilbene oxide, benzoin

3,5-Dimethylphenylcarbamoyl cellulose immobilized on silica gel, 7 pm

6.6 or 0.8 mM citrate buffer (pH 5 or 6)-acetonitrile (30:70, 40:60 or 50:50)

270 mm x 100 lam i.d.

190 mm effective 170 length, chiral separation

[3-adrenergic blocking agent and various enantiomers

Vancomycin bonded on LiChrospher diol silica, 5 pm

Methanol-acetonitrile-acetic 355 mm x 75 l.tm i.d. acid-triethylamine (80-20:2080:0.1-0.3:0.1-0.4)

265 mm effective 171 length, chiral separation

Tricyclic antidepressants

Custom made molecular imprint polymer sorbents

330 mm x 100 ~tm i.d. Acetonitrile-10 mM sodium acetate, pH 3.0 (98:2) with 0.02% trifluoroacetic acid and 0.015% triethylamine

225 mm packed length

Model drug mixture (procaine, timolol, ambroxol, metoclopramide, thiourea, naproxene, antipyrine)

Spherisorb ODS-1, 3 lam, CEC Hypersil C 18, 3 pm or CEC Hypersil C 8 , 3 lam

Acetonitrile-25 mM phosphate, 0.2% hexylamine, pH 2.5 (80:20)

335 mm x 100 pm i.d.

250 mm effective length 56

Carbovir, ranitidine, ondansetron, imipramine, amitriptyline, clomipramine

Water symmetry shield RP-8, 3.5 ~tm

Acetonitrile-100 mMTris, pH 9.0 (70:30)

330 mm x 50 pm i.d.

245 mm packed length

Anti-inflammatory drugs, non-steroidal (etodolac and derivatives)

330 mm x 100 gm i.d. LiChrospher 100 RP-18, 5 gm Acetonitrile-75 (10) mM ammonium formate, pH 2.5 or 3.0 (40:60 or 50:50)

172

173

245 mm packed length, 174 coupling with electrospray ionisationmass spectrometry

~--

Drug test mixture (aminoChiralcel OD, glutethimide, 2,2'-diaminoChiralpak AD, 6,6'-dime thylbiph en yl econa- Chiralcel OJ zole, etozolin, glutethimide, indapamide, metomidate, piprozolin, trans-stilbene oxide, Tr6ger's base)

Methanolic or ethanolic ammonium acetate (10 mM), apparent pH 7.7

Benzodiazepines (nitrazepam, Cholesteryl bonded silica, nimetazepam, estazolam, 6.5 gm, 300 A brotizolam, clotiazepam, oxazolam, haloxazolam, cloxazolam, medazepam) Benzodiazepines (nitrazepam, Cholesteryl modified etched clotiazepam, cloxazolam, capillary medazepam)

305 mm x 100 gm i.d.

220 mm active length, chiral separation

175

Acetonitrile-5 mM Tris.HC1 200 mm (effective buffer, pH 7.3 (35:65 or 25:75) length) x 100 gm i.d.

Comparison with ODS packing

176

Acetonitrile-10 mMTris.HC1 buffer, pH 7.3 (10-20:90-80)

75 gm i.d.

Open tubular CEC, comparison with packed capillary CEC

176

250 or 400 mm packed length

177

t~

Plant constituents

Cannabinoids (cannabigerol, cannabidiol, cannabinol, A-9-tetrahydrocannabinol, A-8-tetrahydrocannabinol, cannabichromene, A-9-tetrahydrocannabinolic acid)

Hypersil C 18 or Hypersil C8, 3 gm

6.25-25 mM phosphate, pH 2.57 (67-75:35-25)

340 or 490 mm x 100 gm i.d.

Morphine alkaloids

Nucleosil 100-C 18, 5 gm

Acetonitrile-2 mM ammonium acetate (40:60)

150 mm x 220 ~tm i.d.

118

TABLE 10.1 (continued) bO

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

Iodide, iodate and perrhenate

Nucleosil SB, 5 gm

5 mMPhosphate buffer, pH2.6

600 mm x 75 ~tm i.d.

400 mm packed length

178

Alkali metals and ammonium

Fused silica capillary 5 mMimidazole-10 mMMES 370 mm x 75 ~tm i.d. suspension of cation exchange [2-(N-morpholino)ethaneparticles (slid-phase reagent, sulfonic acid], pH 6.15 SPR)

305 mm effective length, ion-exchange electrochromatography

179

Compound

Inorganic compounds

Applications

413

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Applications

417

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419

203 M.R. Euerby, C.M. Johnson and J. Mole, Fison Pharmaceuticals, unpublished results, 1994 204 A. Dermaux and P. Sandra, Electrophoresis, 20 (1999) 3027.

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Index of Compounds Separated A Acebutolol 407 Acenaphthene 344, 373,375,377 Acenaphthylene 344, 372, 373,375 Acetaldehyde 379 Acetaminophen 80, 403,409 Acetone 347, 372, 373,375, 379 Acetophenone 187, 347, 378, 379 Acetylaminofluorene deoxyguanosine adduct 311, 312 Acetylsalicylic acid 80, 320, 381,403 Acidic analytes 96, 97, 100, 105, 131 drugs 364, 401 Acids 98, 100, 104 see also individual acids Adenine 259, 363,396 Adenosine 259, 362, 363,395,396 Adenosine-styrene oxide adducts 395 ADP 126, 396 [3-Adrenergetic blocking agents 331, 410 Adrenosterone 385,386 Alanine and derivatives 264, 276, 299-301, 303,305,359 Albumin, tryptic digest 392 Aldehydes 60 Aldopentoses 380 Aldosterone 124, 306, 385, 386 Aliphatic hydrocarbons 343 Alkali metals 412 Alkaloids 397 see also individual alkaloids Alkylbenzenes 173, 174, 177, 212, 372, 374-376 see also individual alkylbenzenes Alkyl benzoates 382 Alkylphenones 187, 379 see also individual alkylphenones Alkyl phenoxypropanoates, chlorinated 383,408

Alprenolol 407 Ambroxol 75, 410 Ambroxolol 367 Amines 104, 388,403 Aminoglutethimide 330 Amino acids 100, 283,333,358-360, 389-391 acetyl 391 benzoyl 391 benzyloxycarbonyl 391 dansyl derivatives 389, 391 dinitrobenzoyl methyl esters 264 N-(3,5-dinitrobenzyloxycarbonyl) derivatives 391 2,4-dinitrophenyl derivatives 389, 391 enantiomers 389-391 9-fluorenylmethoxycarbonyl derivatives 391 phenylthiohydantoin (PTH) derivatives 80, 276, 299-301,303-305, 358, 359, 389, 391 see also individual amino acid derivatives Aminoglutethimide 407, 411 2-Amino-3-hydroxy-3-phenylpropanol 404 2-Aminopyridine 219 Amitriptyline 128, 199, 410 Ammonium 412 AMP 126, 396 Amphetamine 364, 365,400, 401 Amylase 394 Amylbenzene 157, 215,220, 376 Amylobarbital 123, 401 Amylparaben 348 Androstan-3,17-dione 356, 357, 386 5-c~-Androstan-17-one 204 Androsten-3,17-dione 356, 357, 386 Androstenedione 123,385 Androsterone 124, 204, 205 Angiotensins 209, 260, 392, 393 Anhydrotetracycline 370

422 Anilines 212, 219 Anions, see individual anions; Acidic analytes Anthracene 174, 192, 196, 344, 372, 373, 375-377 Anthraquinone dyes 124 1-(9-Anthryl)-2,2,2-trifluoro-ethanol 407 Antibiotics 258, 368-370, 397 see also individual antibiotics Anti-depressants 121, 122, 126, 128, 199 see also individual anti-depressants Anti-epileptics 126, 404 see also individual anti-epileptics Anti-inflammatory drugs, non-steroidal 73, 403,410 Antipyrine 75,367, 410 Antiviral drugs 126, 403 see also individual antiviral drugs Aromatic acids 219 Aromatic compounds 97, 196 Aromatic hydrocarbons 60 Asparagine and derivatives 276, 299-301, 303-305,359 Aspartame 258, 395 Atenolol 407 ATP 126, 396 Atrazine 399 Azo dyes 124 B

Barbital 123,401 Barbiturates 96, 97, 123,402, 405,408 see also individual barbiturates Barbituric acids 327 alkylated 406 Bases 98, 104 see also individual basic compounds Basic analytes 96, 97, 105,267, 331 drugs 364, 401 peptides 394 proteins 394 Bendrofluazide 123,402 Bendroflumethiazide 125,402 Benz[a]anthracene 349 Benzaldehyde 215,220, 236, 237, 376, 377,379 Benzamide 91 Benz[a]anthracene 373,374

Index

1,2-Benzanthracene 372 Benzene 115,215,220, 344, 346, 373 derivatives 215,374-377, 388 Benzoate derivatives 76, 79 Benzo[g]chrysene adduct with deoxyguanosine 311, 312 Benzodiazepines 124, 263,264, 307, 368, 408,409, 411 see also individual azepines Benzo[a]pyrene 174, 175,344, 373 Benzo[b]fluoranthene 344, 373 Benzo [g]chrysene- 11,12-dihydrodiol- 13,14epoxide-DNA adducts 59, 60 B e n z o [ g h i ] p e r y l e n e 344, 373 Benzo[k]fluoranthene 344, 373 Benzoic acid 99, 131, 219 derivatives 384, 385 Benzoin 327, 330, 405,407, 408, 410 acetate 333,408 Benzophenone 376, 377, 379 Benzyl alcohol 91,157, 189, 215,220, 236, 237, 376, 377, 379, 400 Benzylamine 90, 92, 93, 99 Berberine 403 Beta-estradiol 124 Betamethasone 124, 386, 387 dipropionate 124, 387 17-valerate 387, 388,400 Bile acids 201,206, 207, 357 1,1 '- Binaphthy1-2,2'-dihydrogenphosphate 406 1,1 '-Binaphthyl-2,2'-diol 408 Biotin 342 Biphenyl 67, 90, 91,157, 321,377, 379, 384, 385 1-(p-Biphenyl)-ethanol 406 Bradykinin 260, 392, 393 Bromides 128 o-Bromobenzoic acid 131 p-Bromobenzoic acid 131 4-Bromobenzoic acid 219 Brotizolam 411 Budesonide 125 Budesonine 387 Bufadienolide steroids 308 Bufalin 125,386 Bumetanide 402, 403 Bupivacaine 333

423

Subject Index

2-Butanone 379 Butethal 123, 401 N-Butylaniline 219 Butylbenzene 115, 215,220, 346, 372, 374, 377 Butylbenzoic acid 324 Butyl-p-hydroxybenzoic acid 384 Butylparaben 348, 372, 376, 377 Butyraldehyde 379 Butyrophenone 187, 347, 378, 379

Caffeine 80, 93, 99, 258,403,409 Cannabichromene 411 Cannabidiol 411 Cannabigerol 411 Cannabinoids 97, 123,364, 411 see also individual cannabinoids Cannabinol 400, 401, 411 Carbamazepine 404 Carbamazepine- 10,11-diol 404 Carbamazepine- 10,11-epoxide 404 Carbohydrates 348-350 see also individual carbohydrates Carbonic anhydrase 394 Carboxylic acids 320, 381-385 see also individual carboxylic acids Carbovir 410 Cardenolide steroids 308 Cardiac glycosides 404 Carprofen 406 Catechol 196, 378 Cefuroxime axetil 125, 126, 307, 397 Cephalosporin 125, 126, 307, 397 Cerbetocin 394 Chlomipramine 402 Chlorides 128 6-Chloro-a-methylcarbazole-2-acetic cid 406 Chlorobenzenes 212 4-Chlorobenzoic acid 219 Chlorocresol 400 Chlorodiazepam 263 O,L-Chlorodiazepoxide 263,405 Chlorotetracycline 370 Chlorthalidone 335,402, 405 Cholic acid 204 Chrysene 373

Chymotrypsinogen 194, 211,394 Cicloprofen 406 Cinobufagin 125,386 Cinobufatalin 125, 386 Cinosulfuron 369, 371,398 by-products 369, 371,398 Clenbuterol 405,409 Clobetasol butyrate 124, 387 Clobetasol- 17-propionate 400 Clobetasone butyrate 387 Clomipramine 405, 410 Clonazepam 409 Clotiazepam 403, 411 Cloxazolam 403, 411 Cocaine 364, 365,400, 401,403 Codeine phosphate 403 1,3,5-Collidine 219 Conalbumin 100 Corticosteroids 386 see also individual corticosteroids Corticosterone 357, 386 Cortisone 386 acetate 400 Coumarin dyes 399 see also individual categories of dyes Cresols 378 Crotonaldehyde 379 Cyanazine 399 Cyclophophamide 407 Cytidine 362, 363,395,396 Cytochrome c 194, 211, 261,262, 361, 393,395 tryptic digest 72, 309, 310, 321,322, 392 Cytosine 259, 363,396

De-esterified steroids 387 Dehydroisoandrosterone 204, 205 Desethylatrazine 399 Deoxyribonucleic acids, see DNA Desipramine 402 Desisopropylatrazine 399 Desmopressin 394 Detamethasone 127 Dexamethasone 124, 127, 385,386, 388, 400

424

2,2'-Diamino-6,6'-dimethylbiphenyl 330, 411 Diazepam 124, 263,365,401,403,405, 409 Dibenz[a,h]anthracene 344, 373 Dibucaine 384 1,4-Dichlorobenzene 376 Diclofop methyl 327, 408 Diethylphthalate 67, 321,384, 385 Digitalis glycosides 357 Digitoxigenin 125,386, 404 Digoxigenin 125,386, 404 Digoxin 404 2',5'-Dihydroxyacetophenone 378 3,5-Dihydroxybenzoic acid 219 3,4-Dihydroxyphenylalanine (DOPA) 404 2',5'-Dihydroxypropiophenone 378 Dimethylnaphthalene 157 Dimethylphthalate 67, 321,384, 385 3,5-Dinitrobenzoic acid 131 N-(3,5-Dinitrobenzoyl)leucine diallylamide enantiomers 227 N-(3,5-Dinitrobenzyloxycarbonyl) amino acids 391 2,4-Dinitrophenyl amino acids derivatives 358 2,4-Dinitrotoluene 388 Dinucleotides 100 see also Nucleotides Diphenhydramine 90, 92 Disopyramide 407 Diuretics 60, 123,125,307 see also individual diuretics DNA adducts 59, 60, 311 DNB-alanine 405 DNB-leucine 228,229, 331,391 DNZ-amino acids 390 DNZ-leucine 230, 391 DOPA, s e e 3,4-Dihydroxyphenylalanine Doxepine 405 Drugs 235,258-260, 266, 363-368, 400chiral 263,264 metabolites 258 of abuse 364, 400 Dyes 399

Index

E

Econazole 330, 411 Eicosenoic acid 384 Enkephalines 216, 392, 393 Ephedrine 403,404 4-Epianhydrotetracycline 370 Epinephrine 404 4-Epitetracycline 370 Epitizide 123,402 Equiline 204 Erucic acid 384 Estazolam 411 Estradiol 302, 304, 385,387 Estriol 387 Estrone 204, 205,387 Ethosuccinimide 126, 404 N-Ethylaniline 219 Ethylbenzene 115,215,220, 346, 372, 374,376,377 5-Ethyl- 1-methyl-5-(n-propyl) barbituric acid 405 Ethylnaphthalene 157 Ethylparaben 348, 372, 376, 377 Etodolac 406, 410 Etozolin 330, 411 Explosives 119 F

FAPEs, s e e Fatty acids, phenacyl esters Fatty acids 125,350-354, 383,384 methyl esters 302, 350, 384 phenacyl esters 350, 351, 353, 354, 383 Fenoxaprop methyl 327, 408 Fluocortolone 124, 385,386 Fluoranthene 119, 344, 372, 373,375-377 Fluorene 174, 192, 344, 349, 372-377 Fluorides 128 p-Fluorophenylalanine 390 Flurbiprofen 403,406 Fluticasone 385 propionate 127, 387, 388 Fluvoxamine 388 Fmoc-amino acids 390 Fmoc-leucine 331,332, 391 Folic acid 320, 381 Food dyes 127, 399

425

Subject I n d e x

Formaldehyde 379 Formamide 356, 357, 359 Formates 128 Fullerenes 374 G Gitoxigen 386 Glucose 348, 350 Glucuronides, aromatic 380 Glutamine and derivatives 276, 299-301, 303,305,359 Gluthethimide 327, 330, 405, 411 Glycine and derivatives 276, 299-301,303, 305, 359 conjugates 204, 207 Glycosphingolipids 381 Growth hormone, tryptic digest 395 Guanine 259, 363,396 Guanosine 362, 363,395, 396 H

Halogenated hydrocarbons 345 Haloxazolam 411 Herbicides 399 Heroin 364, 365,400, 401 Hexaldehyde 379 Hexanophenone 187 Hexobarbital 121,123,331,401,405-408 Hexylbenzene 175, 176 Hexylparaben 372, 376, 377 Homoatropine 405 Hydrocarbons 343-348, 372-377 Hydrochlorothiazide 409 Hydrocortisone 124, 306, 385-387, 400 derivatives 387 Hydrocortisone- 17-butyrate 400 Hydrocortisone- 17-valerate 400 Hydroflumethiazide 402 Hydrophobic compounds 320 Hydroquinone 196, 379 ethers 196, 379 1113-Hydroxyandrosterone 204, 205 19-Hydroxy-4-androsterone-3,17-dione 204 2-Hydroxyatrazine 399 p-Hydroxybenzoic acid 93,320, 381,384 4-Hydroxybenzoic acid 219 p-Hydroxybenzoic acid 403

c~-Hydroxyethyl-naphthalene 330 13-Hydroxyethyltheophylline 409 17-ct-Hydroxyprogesterone 385 20-c~-Hydroxyprogesterone 385 Hydroxyquinone 378 2-Hydroxyterbutylazine 399 I

Ibuprofen 403,406 IgG, see Immunoglobulin G Imipramine 402, 410 Immunoglobulin G 342 Indapamide 330, 410, 411 Indeno[1,2,3-c,d]pyrene 344, 373 Indomethacin 403 Inorganic compounds 100, 128, 412 see also individual inorganic compounds Inosine 362, 363,396 Inosine-styrene oxide adducts 395 Insecticides 346 Insulin 209, 395 Iodate 99, 412 Iodide 99, 412 4-Iodobenzoic acid 219 Isofamide 407 Isoleucine and derivatives 299-301,358, 359 Isradepin 124, 404 by-products 124

Jatrorrizine 403 K

Ketones 60, 347, 348 Ketoprofen 403

a-Lactalbumin 395 [3-Lactoglobulin 392 Lanthanides 100 Leucine and derivatives 228-230, 299-301, 331,358, 359, 390, 391 see also Amino acids Leucine enkephaline 216

426 Lipids 353 Lorazepam 330, 410 Lysozyme 194, 211, 261,264, 265, 361, 393,395 M

Macrocyclic antibiotics 397 Macrocyclic lactones 125 Malonic acid, sulfate 382 Maltohexaose 348, 350 Malto-oligosaccharides 348-350, 352, 380 Mecoprop methyl 327, 408 Medazepam 411 Mephobarbital 120, 327-329, 405,407, 408 Mepivacaine 333 Methacrolein 379 Methadone 409 Methamphetamine 364, 365,400, 401 Methaqualone 365,366, 401 Methdilazine 402 Methionine enkephaline 216 Methyl amitriptyline 199 Methyl benzoate 376 Methylbenzoin 333,408 Methylchlothiazide 402 Methyl-p-hydroxybenzoic acid 384 Methyl-mandelate 327, 405 Methylnaphthalene 192, 236, 237, 375 Methylparaben 189, 348 c~-Methyl-2,3,4,5,6-pentafluorobenzyl alcohol 406 a-Methyl-a-phenyl-succinimide 327, 405 Methylprednisolone 386 1-Methyl-5-(2-propyl)-5-(n-propyl)barbituric acid 405 17-a-Methyltestosterone 387 Methylthiohydantoin-proline 405 Metoclopramide 75,367, 410 Metolazone 123,402 Metomidate 330, 411 Metoprolol 333,407 Mianserin 406 Monosaccharides 381 see also individual carbohydrates Morphine alkaloids 126, 397, 411 MTH-proline 327, 408 Myoglobin 261,361,393-395

Index

N

Naphthalene 119, 174, 192, 344, 349, 372377 [3-Naphthol 189, 196 1-(2-Naphthyl)ethanol 327, 406 Naproxene 75,367, 403, 410 Neurotensin 392 Neutral analytes 126 compounds 104, 128 drugs 364 Nicotinamide 320, 381 Nicotinic acid 320, 381 Nimetazepam 411 Nitrated hydrocarbons 344, 345 Nitrazepam 124, 403,409, 411 Nitro compounds 388 Nitrobenzene 375,388 p-Nitrobenzoic acid 131 Nitrogen-containing heterocycles 345 p-Nitrophenyl-~-D-glucopyranosides 349, 352, 380 p-Nitrophenyl-ot-o-maltoside 352, 380 p-Nitrophenyl-a-D-maltopentaoside 352, 380 p-Nitrophenyl-ot-D-maltotetraoside 352, 380 p-Nitrophenyl-a-D-maltotrioside 352, 380 p-Nitrophenylglycosides 349 Nitroso compounds 388 N-Nitrosodiethanolamine 388 Nonapeptides 392 see also Peptides Non-ionised (non-polar) analytes 104, 302 Norethindrone 123,385 Nortriptyline 90, 92, 128, 199, 402 Noscapine 364, 365,400, 401 Nucleic acids 100, 311, 312 see also DNA; RNA Nucleobases 100, 259, 363,396 Nucleosides 97, 125,362, 363,395,396 Nucleotides 126, 128, 396 O Octanophenone 187 Oils, vegetable 125 Oleic acid 384

Subject Index

Oligonucleotides 277 see also Nucleotides Oligopeptides 201, 210 see also Peptides Oligosaccharides 201,202, 348, 351,380 see also individual carbohydrates Ondansetron 410 Oxazepam 263,409 Oxazolam 411 Oxygen-containing heterocycles 345 Oxytocin 394 Ovalbumin, tryptic digest 394 P

PAHs, see Polyaromatic hydrocarbons Palmitoleic acid 384 Parabens 127, 348,372, 376, 400 Paracetamol, metabolites 80 Pentapeptides 392 Pentobarbital 405,407 Pentylbenzene 115,346, 372, 374 Pentylparaben 372, 376, 377 Peptide drugs, metabolites 68 Peptides 72, 73, 125, 131,132, 201,209, 210, 216, 260-262, 277, 309, 310, 321, 322, 359-362, 391-395 C-protected 393 N-protected 393 Perchlorates 128 Perrhenate 412 Pesticides 346, 369, 398, 399 Pharmaceuticals 97, 104, 258-260 diverse 65 Phenanthrene 174, 192, 196, 344, 349, 372377 Phenobarbital 123,365,366, 400, 401 Phenols 97, 196, 212, 372, 378, 388 Phenothiazine 376 Phenylalanine and derivatives 299, 300, 334, 358, 359, 389, 390 amide 390 Phenylbutazone 409 T-Phenyl-T-butyrolactone 327 Phenylenediamines 212 1-Phenylethanol 406 2-Phenylethylamine derivatives 404 Phenylglycine 390 2-Phenylmethyl-1-naphthol 378

427 Phenyl propanol 388,406 2-Phenylpropionic acid 382 Phenylthiohydantoin-amino acids 80, 276, 299-301,303-305,358, 359, 389, 391 Phenytoin 126, 404 Phthalate esters 382 Pindolol 407 Piprozolin 330, 411 Polyaromatic hydrocarbons (PAHs) 59, 78, 116, 127, 173, 174, 177, 343-345,372377, 379, 409 see also individual polyaromatic hydrocarbons Polychlorinated dibenzo-p-dioxins 399 Polystyrene standards 224, 376 Prednisolone 124, 386, 387 acetate 400 Pregnan-3,20-dione 356, 357, 386 4-Pregnen-20~-ol-3-one 387 Prenalterol 407 Preservatives 97 Primicarb 398 Primidon 126, 404 Procainamide 90-92 Procaine 75,364, 365,367, 400, 401, 410 Profens 391 Progesterone 123,204, 356, 385,387 metabolites 356 Proline and derivatives 299-301,327, 359 Propazine 399 Propionaldehyde 379 Propiophenone 378 Propranolol 333,384, 407, 408 Propylbenzene 115, 215,220, 346, 372, 374,376 Propyl-p-hydroxybenzoic acid 384 Propylparaben 348, 372, 376, 377 Prostaglandins 126, 381 Protein(s) 100, 194, 201, 211,225,260262, 264, 309, 361,392-395 digests 309, 310, 361,392 sequencing 283 see also individual proteins PTH-amino acids, see Phenylthiohydantoinamino acids Purine bases 128, 363,396 see also Nucleobases Pyrene 174, 196, 344, 349, 373-375,377

428 9-(1-Pyrene)nonanol 137 Pyridine 378 Pyrimidine bases 128, 363,396 see also Nucleobases

Q Quinine 364, 365,400, 401 R

Ranitidine 410 Resorcinol 189, 196, 378 Retinyl esters 123,383,397 Ribonucleases 194, 211, 261,264, 361, 393-395 t-RNA 100, 128, 312 Ropivacaine 333 S

S-oxidation compounds 124 Salbutamol 409 Salsolinol 335,406 Sebutylazine 399 Secobarbital 123, 401 Serin 390 Serotonin 258,265 Simazine 399 Softeners 346 Sorbic acid 400 Steroid A 125 Steroid drugs 355 glycosides 357 hormones 357 Steroids 124, 125, 188, 201,203-205,307, 308,355-358,385-388 see also individual steroids trans-Stilbene oxide 330, 333,408, 410, 411 Sugars 201 Sulfadicramide 404 Sulfaflurazol 404 Sulfanilamide 404 Sulfides 128 Sulfonates, aliphatic 128 Sulphur-containing heterocycles 345 Suramin 126 Sweeteners, artificial, see Aspartame

lndex

T Taurine conjugates 204, 207 Temazepam 263,405,407, 409 Terbutaline 408 Terbutylazine 399 o-Terphenyl 67, 321,384, 385 Testosterone 123, 124, 306, 357, 365, 385387, 400, 401 decanoate 383 esters 353 isocaproate 383 phenyl propionate 383 propionate 365,383,400, 401 1,2,4,5-Tetrachlorobenzene 376 Tetracyclines 258, 370, 397 A-8-Tetrahydrocannabinol 411 see also Cannabinoids A-9-Tetrahydrocannabinol 365,400, 401, 411 see also Cannabinoids A-9-Tetrahydrocannabinolic acid 365,400, 401, 411 see also Cannabinoids Tetrapeptides 392, 393 see also Peptides Textile dyes 124, 399 Thalidomide 330-332, 409 metabolites 330, 409 Thebaine 403 Theobromine 409 Theophylline 258, 409 Thiazide diuretics 307 Thiocyanates 128 Thiourea 67, 75, 90, 91, 119, 157, 175, 176, 215,220, 236, 237, 320, 321, 362, 367, 376, 377, 379, 381,384, 385, 388, 400, 410 Thomapyrin 80, 403 Threonine and derivatives 276, 299-301, 303,305,359 Thymidine 362, 395,396 Thymine 259, 363,396 Timolol 75,367, 410 Tipredane 125,369, 385 p-Tolualdehyde 379 o-Toluic acid 131 Toluene 115,215,220, 346, 372, 374, 376

429

Subject Index

2-Toluic acid 219 Triamcinolone 386 acetonide 386 Triazine herbicides 123 see also individual herbicides 1,3,5-Trichlorobenzene 376 Tricyclic anti-depressants 126, 410 see also individual anti-depressants Triglycerides 125,352-355,382, 383 1,2,3-Trimethylbenzene 375 Triphenylene 174 Tripredane 401 Tr6ger's base 330, 333,408,411 Tropicamide 405 Trypsinogen 394 Tryptamine 258,265 Tryptophan and derivatives 299, 300, 331, 359, 390, 391 Tyrosine and derivatives 276, 299-301, 303,305,359, 390

Tyrosine-containing peptides 392 see also Peptides

Uracil 363,396 see also Nucleobases Urea herbicides 398 see also individual herbicides Uridine 362, 363,395,396 see also Nucleosides V Valeraldehyde 379 Valine and derivatives 264, 299-301,358 see also Amino acids Vitamins 302, 397 W Warfarin 330, 406, 407

This Page Intentionally Left Blank

431

JOURNAL

OF C H R O M A T O G R A P H Y

LIBRARY

A Series of Books Devoted to Chromatographic and Electrophoretic Techniques and their Applications Although complementary to the Journal of Chromatography, each volume in the Library Series is an important and independent contribution in the field of chromatography and electrophoresis. The library contains no material reprinted from the journal itself.

O t h e r V o l u m e s in this s e r i e s Volume 1

Chromatography of Antibiotics (see also Volume 26) by G.H. Wagman and M.J. Weinstein

Volume 2

Extraction Chromatography edited by T. Braun and G. Ghersini

Volume 3

Liquid Column Chromatography. A Survey of Modern Techniques and Applications edited by Z. Deyl, K. Macek and J. Jan~ik

Volume 4

Detectors of Gas Chromatography by J. t~ev~ik

Volume 5

Instrumental Liquid Chromatography. A Practical Manual on HighPerformance Liquid Chromatographic Methods (see also Volume 27) by N.A. Parris

Volume 6 Volume 7

Isotachophoresis. Theory, Instrumentation and Applications by F.M. Everaerts, J.L. Beckers and Th.P.E.M. Verheggen Chemical Derivatization in Liquid Chromatography by J.F. Lawrence and R.W. Frei

Volume 8

Chromatography of Steroids by E. Heftmann

Volume 9

HPTLC - High Performance Thin-Layer Chromatography edited by A. Zlatkis and R.E. Kaiser

Volume 10

Gas Chromatography of Polymers by V.G. Berezkin, V.R. Alishoyev and I.B. Nemirovskaya

Volume 11

Liquid Chromatography Detectors

Volume 12

Affinity Chromatography (see also Volume 55)

by R.P.W. Scott by J. Turkov~ Volume 13

Instrumentation for High-Performance Liquid Chromatography edited by J.F.K. Huber

Volume 14

Radiochromatography. The Chromatography and Electrophoresis of Radiolabelled Compounds by T.R. Roberts

Volume 15

Antibiotics. Isolation, Separation and Purification edited by M.J. Weinstein and G.H. Wagman

432

Volume 16

P o r o u s Silica. Its Properties and Use as Support in Column Liquid Chromatography by K.K. Unger

Volume 17

75 Years of C h r o m a t o g r a p h y - A Historical Dialogue edited by L.S. Ettre and A. Zlatkis

Volume 18

E l e c t r o p h o r e s i s . A Survey of Techniques and Applications P a r t A: T e c h n i q u e s P a r t B: Applications edited by Z. Deyl

Volume 19

Chemical D e r i v a t i z a t i o n in Gas C h r o m a t o g r a p h y by J. Drozd

Volume 20

E l e c t r o n C a p t u r e . Theory and Practice in Chromatography edited by A. Zlatkis and C.F. Poole

Volume 21

E n v i r o n m e n t a l P r o b l e m s Solving u s i n g Gas a n d Liquid Chromatography by R.L. Grob and M.A. Kaiser

Volume 22

C h r o m a t o g r a p h y . Fundamentals and Applications of Chromatographic and Electrophoretic Methods (see also Volume 51) P a r t A: F u n d a m e n t a l s

P a r t B: Applications edited by E. Heftmann Volume 23

C h r o m a t o g r a p h y of Alkaloids P a r t A: T h i n - L a y e r C h r o m a t o g r a p h y by A. Baerheim-Svendsen and R. Verpoorte P a r t B: Gas-Liquid C h r o m a t o g r a p h y a n d H i g h - P e r f o r m a n c e Liquid C h r o m a t o g r a p h y by R. Verpoorte and A. Baerheim-Svendsen

Volume 24

C h e m i c a l M e t h o d s in Gas C h r o m a t o g r a p h y by V.G. Berezkin

Volume 25

M o d e r n Liquid C h r o m a t o g r a p h y of M a c r o m o l e c u l e s by B.G. Belenkii and L.Z. Vilenchik

Volume 26

C h r o m a t o g r a p h y on Antibiotics. Second, Completely R e v i s e d Edition by G.H. Wagman and M.J. Weinstein

Volume 27

I n s t r u m e n t a l Liquid C h r o m a t o g r a p h y . A Practical Manual on High-Performance Liquid Chromatographic Methods. Second, Completely Revised Edition by N.A. Parris

Volume 28

M i c r o c o l u m n H i g h - P e r f o r m a n c e Liquid C h r o m a t o g r a p h y by P. Kucera

Volume 29

Q u a n t i t a t i v e C o l u m n Liquid C h r o m a t o g r a p h y . A Survey of Chemometric Methods by S.T. Balke

433

Volume 30

M i c r o c o l u m n S e p a r a t i o n s . Columns, Instrumentation and Ancillary Techniques by M.V. Novotny and D. Ishii

Volume 31

G r a d i e n t E l u t i o n in C o l u m n Liquid C h r o m a t o g r a p h y . Theory and Practice by P. Jandera and J. ChurfiSek

Volume 32

The Science of C h r o m a t o g r a p h y . Lectures Presented at the A.J.P. Martin Honorary Symposium, Urbino, May 27-31, 1985 edited by F. Bruner

Volume 33

Liquid C h r o m a t o g r a p h y Detectors. Second, C o m p l e t e l y R e v i s e d Edition by R.P.W. Scott

Volume 34

P o l y m e r C h a r a c t e r i z a t i o n by Liquid C h r o m a t o g r a p h y by G. G16ckner

Volume 35

O p t i m i z a t i o n of C h r o m a t o g r a p h i c Selectivity. A Guide to Method Development by P.J. Schoenmakers

Volume 36

Selective Gas C h r o m a t o g r a p h i c D e t e c t o r s by M. Dressler

Volume 37

C h r o m a t o g r a p h y of Lipids in Biomedical R e s e a r c h a n d Clinical Diagnosis edited by A. Kuksis

Volume 38

P r e p a r a t i v e Liquid C h r o m a t o g r a p h y edited by B.A. Bidlingmeyer

Volume 39A

Selective S a m p l e H a n d l i n g a n d D e t e c t i o n in H i g h - P e r f o r m a n c e Liquid C h r o m a t o g r a p h y . P a r t A by R.W. Frei and K. Zech

Volume 39B

Selective S a m p l e H a n d l i n g a n d D e t e c t i o n in H i g h - P e r f o r m a n c e Liquid C h r o m a t o g r a p h y . P a r t B by K. Zech and R.W. Frei

Volume 40

A q u e o u s Size-Exclusion C h r o m a t o g r a p h y by P. Dubin

Volume 41

H i g h - P e r f o r m a n c e Liquid C h r o m a t o g r a p h y of B i o p o l y m e r s a n d Biooligomers Part A: Principles, Materials and Techniques P a r t B: S e p a r a t i o n of I n d i v i d u a l C o m p o u n d s Classes by O. Mike~

Volume 42

Q u a n t i t a t i v e Gas C h r o m a t o g r a p h y for L a b o r a t o r y Analyses a n d Online P r o c e s s C o n t r o l by G. Guiochon and C.L. Guillemin

Volume 43

N a t u r a l P r o d u c t s Isolation. Separation Methods for Antimicrobials, Antivirals and Enzyme Inhibitors edited by G.H. Wagman and R. Cooper

Volume 44

Analytical Artifacts. GC, MS, HPLC, TLC and PC by B.S. Middleditch

434

Volume 45A

C h r o m a t o g r a p h y and Modification of N u c l e o s i d e s Analytical Methods for Major and Modified N u c l e o s i d e s HPLC, GC, MS, NMR, UV and FT-IR edited by C.W. Gehrke and K.C.T. Kuo

Volume 45B

C h r o m a t o g r a p h y a n d Modification of N u c l e o s i d e s Biological Roles a n d F u n c t i o n of Modification edited by C.W. Gehrke and K.C.T. Kuo

Volume 45C

C h r o m a t o g r a p h y a n d Modification of N u c l e o s i d e s Modified N u c l e o s i d e s in C a n c e r a n d Normal Metabolism Methods and Applications edited by C.W. Gehrke and K.C.T. Kuo

Volume 45D

C h r o m a t o g r a p h y a n d Modification of N u c l e o s i d e s C o m p r e h e n s i v e D a t a b a s e for RNZ and DNA N u c l e o s i d e s Chemical, Biochemical, Physical, Spectral and S e q u e n c e edited by C.W. Gehrke and K.C.T. Kuo

Volume 46

Ion C h r o m a t o g r a p h y : Principles and Applications by P.R. Haddad and P.E. Jackson

Volume 47

Trace Metal Analysis and Speciation edited by I.S. Krull

Volume 48

S t a t i o n a r y P h a s e s in Gas C h r o m a t o g r a p h y by H. Rotzsche

Volume 49

Gas C h r o m a t o g r a p h y in Air Pollution Analysis by V.G. Berezkin and Yu.S. Drugov

Volume 50

Liquid C h r o m a t o g r a p h y in B i o m e d i c a l Analysis edited by T. Hanai

Volume 51

C h r o m a t o g r a p h y , 5th edition. Fundamentals and Applications of Chromatographic and Related Differential Migration Methods Part A: F u n d a m e n t a l s and Techniques P a r t B: Applications edited by E. Heftmann

Volume 52

Capillary E l e c t r o p h o r e s i s . Principles, Practice and Applications by S.F.Y. Li

Volume 53

H y p h e n a t e d Techniques in Supercritical Fluid C h r o m a t o g r a p h ~ and Extraction edited by K. Jinno

Volume 54

C h r o m a t o g r a p h y of Mycotoxins. Techniques and Applications edited by V. Betina

Volume 55

Bioaffinity C h r o m a t o g r a p h y . Second, completely revised edition by J. Turkov~

Volume 56

C h r o m a t o g r a p h y in the P e t r o l e u m I n d u s t r y edited by E.R. Adlard

Volume 57

Retention and Selectivity in Liquid C h r o m a t o g r a p h y . Prediction, Standardisation and Phase Comparisons edited by R.M. Smith

435

Volume 58

C a r b o h y d r a t e Analysis edited by Z. E1 Rassi

Volume 59

Applications of Liquid C h r o m a t o g r a p h y / Mass S p e c t r o m e t r y in Environmental Chemistry edited by D. Barcel5

Volume 60

A d v a n c e d C h r o m a t o g r a p h i c a n d E l e c t r o m i g r a t i o n M e t h o d s in BioSciences edited by Z. Deyl, I. Mikgik, F. Tagliaro and E. Tesa~ov~

Volume 61

P r o t e i n Liquid C h r o m a t o g r a p h y edited by M. Kastner

Volume 62

Capillary E l e c t r o c h r o m a t o g r a p h y edited by Z. Deyl and F. Svec

This Page Intentionally Left Blank

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Capillary Electrophoresis of Nucleic Acids. Introduction to the Capillary Electrophoresis

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Capillary Electrophoresis of Nucleic Acids. Practical Applications of Capillary Electrophoresis

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Capillary Flows with Forming Interfaces

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Capillary Electrophoresis in Chiral Analysis

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Capillary Flows with Forming Interfaces

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Capillary Electrophoresis. Methods and Protocols

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Capillary Electrophoresis: Methods and Protocols

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Capillary Electrophoresis in Polymer Analysis

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Clinical Applications of Capillary Electrophoresis

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Practical Capillary Electrophoresis, Second Edition

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Gravity-Capillary Free-Surface Flows

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Chemometric Methods in Capillary Electrophoresis

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Capillary Electrophoresis Guidebook: Principles, Operation, and Applications

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Capillary Electrophoresis of Proteins and Peptides

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Capillary Fluid Exchange: Regulation, Functions, and Pathology

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