Retinoids: Methods and Protocols (Methods in Molecular Biology)

ME T H O D S

IN

MO L E C U L A R BI O L O G Y

Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

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Retinoids Methods and Protocols

Edited by

Hui Sun Department of Physiology, Jules Stein Eye Institute, and Brain Research Institute, David Geffen School of Medicine, University of California, Los Angeles, CA, USA

Gabriel H. Travis Departments of Ophthalmology and Biological Chemistry, Jules Stein Eye Institute, David Geffen School of Medicine, University of California, Los Angeles, CA, USA

Editors Hui Sun Department of Physiology Jules Stein Eye Institute and Brain Research Institute David Geffen School of Medicine 650 Charles Young Drive South University of California Los Angeles CA 90095, USA [email protected]

Gabriel H. Travis Departments of Ophthalmology and Biological Chemistry Jules Stein Eye Institute 100 Stein Plaza David Geffen School of Medicine University of California Los Angeles CA 90095, USA [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60327-324-4 e-ISBN 978-1-60327-325-1 DOI 10.1007/978-1-60327-325-1 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2010928266 © Springer Science+Business Media, LLC 2010 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface Technical advancement is a major driving force in the experimental sciences, and retinoid research is no exception. Ancient Egyptians recognized that fresh liver can cure night blindness. More than 3000 years later, vitamin A was identified as the essential ingredient in liver. Since then, the pace of discovery has accelerated due to the advent of new techniques, especially during the recent decades. The molecular mechanism for vitamin A’s physiological function was first elucidated in vision. Today, the biological functions of vitamin A have been found in almost all vertebrate organs, and its multitasking ability has continued to surprise researchers. In addition to vision, known biological functions of vitamin A include its roles in embryonic growth and development, immune competence, reproduction, maintenance of epithelial surfaces, and proper functioning of the adult brain. At the biochemical level, vitamin A derivatives serve distinct functions as photoreceptor chromophores, as transcriptional regulators through the control of nuclear hormone receptors, and as translational regulators, a function discovered recently. Since vitamin A derivatives have potent biological activities, especially in their effects on cellular growth and differentiation, imbalances in vitamin A homeostasis are associated with a wide range of pathological conditions, such as visual disorders, cancer, infectious diseases, diabetes, teratogenicity, and skin diseases. New biological functions are still being discovered for vitamin A derivatives. For example, it was recently discovered that retinol inhibits adipogenesis. Retinol, but not retinoic acid, has the ability to maintain the pluripotency of embryonic stem cells. Retinoic acid plays surprising roles in regulating protein translation in neurons. Certain tissues have the ability to accumulate surprisingly high concentrations of retinoids under physiological conditions. For example, when channelrhodopsin, which uses all-trans-retinal as its chromophore, and rhodopsin, which uses 11-cis-retinal as its chromophore, are expressed in different regions of the mouse brain in the optogenetic technique to study neural circuits, they become light sensitive without the addition of exogenous retinoid. The physiological functions of retinoids in the adult brain are beginning to emerge, including their roles in sleep, learning, and memory. The purpose of this book is to summarize recent technical tools to help researchers in diverse fields to uncover more surprises in the future. The target audience of this book includes both beginning researchers and experienced researchers who would like to learn new techniques. All chapters were written by experts on the subjects. Topics cover diverse techniques for both in vitro and in vivo studies. A special chapter provides advice on the practical use of diets in both animal and human research on vitamin A. Biochemical techniques include the detection and quantitation of retinoids using HPLC, mass spectrometry, and fluorescence and techniques to study visual pigments, retinoid isomerase, a membrane transporter for retinoid, A2E, retinoic acid catabolism, and cellular vitamin A uptake. Biophysical techniques include fluorescence anisotropy of retinol binding protein, electrophysiology to study retinoid cycle in vision, visualization of retinoid in native tissues, two-photon microscopy to study retinoid transport, and epifluorescence to study

v

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Preface

retinol in photoreceptor cells. Cell biological techniques include cell culture models for studying retinoid transport and the role of retinol in embryonic stem cell culture. Hui Sun Gabriel H. Travis

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

v

Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

ix

1.

Quantification of Endogenous Retinoids . . . . . . . . . . . . . . . . . . . . . Maureen A. Kane and Joseph L. Napoli

1

2.

Culture of Highly Differentiated Human Retinal Pigment Epithelium for Analysis of the Polarized Uptake, Processing, and Secretion of Retinoids . . . . . Jane Hu and Dean Bok

55

Feeder-Independent Culture of Mouse Embryonic Stem Cells Using Vitamin A/Retinol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jaspal S. Khillan and Liguo Chen

75

In Vitro Assays of Rod and Cone Opsin Activity: Retinoid Analogs as Agonists and Inverse Agonists . . . . . . . . . . . . . . . . . . . . . . . . . . . Masahiro Kono and Rosalie K. Crouch

85

Physiological Studies of the Interaction Between Opsin and Chromophore in Rod and Cone Visual Pigments . . . . . . . . . . . . . . . Vladimir J. Kefalov, M. Carter Cornwall, and Gordon L. Fain

95

3.

4.

5.

6.

Measurement of the Mobility of All-Trans-Retinol with Two-Photon Fluorescence Recovery After Photobleaching Yiannis Koutalos

. . . . . . . . . 115

7.

Microfluorometric Measurement of the Formation of All-Trans-Retinol in the Outer Segments of Single Isolated Vertebrate Photoreceptors . . . . . . . 129 Yiannis Koutalos and M. Carter Cornwall

8.

HPLC / MSN Analysis of Retinoids . . . . . . . . . . . . . . . . . . . . . . . . 149 James E. Evans and Peter McCaffery

9.

Binding of Retinoids to ABCA4, the Photoreceptor ABC Transporter Associated with Stargardt Macular Degeneration . . . . . . . . . . . . . . . . . 163 Ming Zhong and Robert S. Molday

10.

Fluorescence-Based Technique for Analyzing Retinoic Acid . . . . . . . . . . . . 177 Leslie J. Donato and Noa Noy

11.

The Interaction Between Retinol-Binding Protein and Transthyretin Analyzed by Fluorescence Anisotropy . . . . . . . . . . . . . . . . . . . . . . . 189 Claudia Folli, Roberto Favilla, and Rodolfo Berni

12.

Assay of Retinol-Binding Protein–Transthyretin Interaction and Techniques to Identify Competing Ligands . . . . . . . . . . . . . . . . . . . . 209 Nathan L. Mata, Kim Phan, and Yun Han

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Contents

13.

Molecular Biology and Analytical Chemistry Methods Used to Probe the Retinoid Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229 Marcin Golczak, Grzegorz Bereta, Akiko Maeda, and Krzysztof Palczewski

14.

Visualization of Retinoid Storage and Trafficking by Two-Photon Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 247 Yoshikazu Imanishi and Krzysztof Palczewski

15.

Reverse-Phase High-Performance Liquid Chromatography (HPLC) Analysis of Retinol and Retinyl Esters in Mouse Serum and Tissues . . . . . . . . 263 Youn-Kyung Kim and Loredana Quadro

16.

Detection of Retinoic Acid Catabolism with Reporter Systems and by In Situ Hybridization for CYP26 Enzymes . . . . . . . . . . . . . . . . . . . . 277 Yasuo Sakai and Ursula C. Dräger

17.

Diet in Vitamin A Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . 295 A. Catharine Ross

18.

Experimental Approaches to the Study of A2E, a Bisretinoid Lipofuscin Chromophore of Retinal Pigment Epithelium . . . . . . . . . . . . . . . . . . . 315 Janet R. Sparrow, So Ra Kim, and Yalin Wu

19.

Analysis of the Retinoid Isomerase Activities in the Retinal Pigment Epithelium and Retina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 329 Gabriel H. Travis, Joanna Kaylor, and Quan Yuan

20.

Techniques to Study Specific Cell-Surface Receptor-Mediated Cellular Vitamin A Uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341 Riki Kawaguchi and Hui Sun

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363

Contributors GRZEGORZ BERETA • Department of Pharmacology, School of Medicine, Case Western Reserve University, 10900 Euclid Ave, Cleveland, OH 44106-4965, USA RODOLFO BERNI • Department of Biochemistry and Molecular Biology, University of Parma, Via G.P. Usberti 23/A, 43100 Parma, Italy DEAN BOK • Jules Stein Eye Institute and Department of Neurobiology, David Geffen School of Medicine, University of California, Los Angeles, CA, USA LIGUO CHEN • Department of Microbiology and Molecular Genetics, 3501 Fifth Avenue, University of Pittsburgh, Pittsburgh, PA 15261, USA M. CARTER CORNWALL • Department of Physiology and Biophysics, Boston University School of Medicine, Boston, MA, USA ROSALIE K. CROUCH • Department of Ophthalmology, Medical University of South Carolina, 167 Ashley Ave, Room 511, Charleston, SC 29425, USA LESLIE J. DONATO • Department of Biochemistry, University of Wisconsin-Madison, Madison, WI, USA URSULA C. DRÄGER • Eunice Kennedy Shriver Center for Mental Retardation, University of Massachusetts Medical School, Waltham, MA, USA JAMES E. EVANS • Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, MA 01655, USA GORDON L. FAIN • Department of Physiological Science and Jules Stein Eye Institute, University of California, Los Angeles, CA, USA ROBERTO F AVILLA • Department of Biochemistry and Molecular Biology, University of Parma, Via G.P. Usberti 23/A, 43100 Parma, Italy CLAUDIA F OLLI • Department of Biochemistry and Molecular Biology, University of Parma, Via G.P. Usberti 23/A, 43100 Parma, Italy MARCIN GOLCZAK • Department of Pharmacology, School of Medicine, Case Western Reserve University, 10900 Euclid Ave, Cleveland, OH 44106-4965, USA YUN HAN • Sirion Therapeutics, San Diego, CA, USA JANE HU • Jules Stein Eye Institute and Department of Neurobiology, David Geffen School of Medicine, University of California, Los Angeles, CA, USA YOSHIKAZU IMANISHI • Department of Pharmacology, School of Medicine, Case Western Reserve University, 10900 Euclid Ave, Cleveland, OH 44106-4965, USA MAUREEN A. K ANE • Department of Pharmaceutical Sciences, University of Maryland, Baltimore, MD, USA RIKI KAWAGUCHI • Department of Physiology, Jules Stein Eye Institute, and Brain Research Institute, David Geffen School of Medicine, University of California, Los Angeles, CA 90095-1751, USA JOANNA KAYLOR • Jules Stein Eye Institute, UCLA School of Medicine, Los Angeles, CA, USA VLADIMIR J. KEFALOV • Department of Ophthalmology and Visual Sciences and Department of Anatomy and Neurobiology, Washington University School of Medicine, St. Louis, MO, USA

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Contributors

JASPAL S. KHILLAN • Department of Microbiology and Molecular Genetics, 3501 Fifth Avenue, University of Pittsburgh, Pittsburgh, PA 15261, USA SO RA KIM • Departments of Ophthalmology and Pathology and Cell Biology, Columbia University, New York, NY 10032, USA YOUN-KYUNG KIM • Department of Food Science and Rutgers Center for Lipid Research, School of Environmental and Biological Sciences, Rutgers University, 65 Dudley Road, New Brunswick, NJ 08901, USA MASAHIRO KONO • Department of Ophthalmology, Medical University of South Carolina, 167 Ashley Ave, Room 511, Charleston, SC 29425, USA YIANNIS KOUTALOS • Departments of Ophthalmology and Neurosciences, Medical University of South Carolina, 167 Ashley Avenue, Charleston, SC 29425, USA NATHAN L. M ATA • Sirion Therapeutics, San Diego, CA, USA AKIKO MAEDA • Department of Pharmacology, School of Medicine, Case Western Reserve University, 10900 Euclid Ave, Cleveland, OH 44106-4965, USA PETER MCCAFFERY • Institute of Medical Sciences, School of Medical Sciences, Foresterhill, Aberdeen, Scotland ROBERT S. MOLDAY • Department of Biochemistry and Molecular Biology, 2350 Health Sciences Mall, University of British Columbia, Vancouver, BC V6T 1Z3, Canada JOSEPH L. NAPOLI • Department of Nutritional Science and Toxicology, University of California, Berkeley, CA, USA NOA NOY • Departments of Pharmacology and Nutrition, Case Western Reserve University School of Medicine, 10900 Euclid Ave, SOM Rm W333, Cleveland, OH 44106, USA KRZYSZTOF PALCZEWSKI • Department of Pharmacology, School of Medicine, Case Western Reserve University, 10900 Euclid Ave, Cleveland, OH 44106-4965, USA KIM PHAN • Sirion Therapeutics, San Diego, CA, USA LOREDANA QUADRO • Department of Food Science and Rutgers Center for Lipid Research, School of Environmental and Biological Sciences, Rutgers University, 65 Dudley Road, New Brunswick, NJ 08901, USA A. CATHARINE ROSS • Department of Nutritional Sciences and Huck Institute for the Life Sciences, Pennsylvania State University, University Park, PA 16802, USA YASUO SAKAI • Department of Plastic Surgery, Osaka University School of Medicine, Osaka, Japan JANET R. SPARROW • Departments of Ophthalmology and Pathology and Cell Biology, Columbia University, New York, NY 10032, USA HUI SUN • Department of Physiology, Jules Stein Eye Institute, and Brain Research Institute, David Geffen School of Medicine, University of California, Los Angeles, CA 90095-1751, USA GABRIEL H. TRAVIS • Jules Stein Eye Institute, UCLA School of Medicine, Los Angeles, CA, USA YALIN WU • Departments of Ophthalmology and Pathology and Cell Biology, Columbia University, New York, NY 10032, USA QUAN YUAN • Jules Stein Eye Institute, UCLA School of Medicine, Los Angeles, CA, USA MING ZHONG • Department of Biochemistry and Molecular Biology, University of British Columbia, Vancouver, BC, Canada

Chapter 1 Quantification of Endogenous Retinoids Maureen A. Kane and Joseph L. Napoli Abstract Numerous physiological processes require retinoids, including development, nervous system function, immune responsiveness, proliferation, differentiation, and all aspects of reproduction. Reliable retinoid quantification requires suitable handling and, in some cases, resolution of geometric isomers that have different biological activities. Here we describe procedures for reliable and accurate quantification of retinoids, including detailed descriptions for handling retinoids, preparing standard solutions, collecting samples and harvesting tissues, extracting samples, resolving isomers, and detecting with high sensitivity. Sample-specific strategies are provided for optimizing quantification. Approaches to evaluate assay performance also are provided. Retinoid assays described here for mice also are applicable to other organisms including zebrafish, rat, rabbit, and human and for cells in culture. Retinoid quantification, especially that of retinoic acid, should provide insight into many diseases, including Alzheimer’s disease, type 2 diabetes, obesity, and cancer. Key words: Retinoid, retinoic acid, retinal, retinaldehyde, retinol, retinyl ester, mass spectrometry, LC/MS/MS, HPLC.

1. Introduction Retinoid homeostasis involves balance among several retinoids in multiple tissues effected through dietary intake, storage, mobilization, transport, and metabolism (see Fig. 1.1) (1–5). Lecithin:retinol acyltransferase (LRAT) and in skin diacylglycerol acyltransferase-2 (DGAT2) and retinyl ester hydrolases (REH) mediate storage and mobilization of vitamin A (retinol), respectively. Metabolism activates retinol into retinoic acid (RA) by a reversible and rate-limiting dehydrogenation of retinol into retinal, catalyzed by short-chain retinol dehydrogenases (SDR), followed by an irreversible dehydrogenation H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_1, © Springer Science+Business Media, LLC 2010

1

2

Kane and Napoli Typical tissue levels

O O (CH2)nCH3 all-trans-retinyl palmitate and other RE REH

0.5–500 nmol/g

LRAT/DGAT2

OH all-trans-retinol (vitamin A)

Rdh (SDR )

0.05–50 nmol/g

Rrd (SDR )

O H 0.01–0.5 nmol/g

all-trans-retinal Raldh (Aldh)

O 9 4

OH

13

18

all-trans-retinoic acid

0.00001–0.05 nmol/g (0.5–50 pmol/g)

Cyp

O OH O

4-oxo-retinoic acid

+

Regulation of transcription and translation

O OH

CH2OH 18-OH-retinoic acid

Fig. 1.1. Structures of analytes in the central pathway of retinoid metabolism. Typical in vivo levels of each analyte are listed. Ranges reflect variation among tissues and dietary conditions.

by retinal dehydrogenases (RALDHs) into RA. A number of cytochrome P450 (CYP) enzymes catabolize RA to polar metabolites (6, 7). All-trans-RA (atRA) mediates a multitude of systemic effects, including development, nervous system function, immune response, cell proliferation, cell differentiation, and reproduction, by regulating transcription of hundreds of genes through binding to retinoic acid receptors (RAR) α, β, γ and peroxisome proliferator-activated receptor (PPAR)β/δ (8–11). Expression loci of specific retinoid-binding proteins, enzymes, and receptors, which contribute to RA generation, signaling, and catabolism, indicate that RA concentrations in vivo are temporally/spatially controlled to produce the individual actions of vitamin A (6, 7, 12–15).

Quantification of Endogenous Retinoids

3

Various approaches have been used to determine effectors of retinoid metabolism, including genetic alteration of retinoid-binding proteins (16–21), enzymes (22–25), and receptors (8, 26, 27); dietary manipulation of vitamin A intake (28); and exposure to xenobiotics (29–34). Quantifying how manipulation of retinoid metabolism effects the flux of retinoids through metabolic paths, the availability of substrate for RA production, and/or endogenous RA levels will provide insight into retinoid homeostasis and metabolism and, thereby, function. Dysfunctions in retinoid homeostasis have been linked to dyslipidemia, diabetes, obesity, cancer, and Alzheimer’s disease, but these data have not necessarily been accompanied by robust quantification of retinoid concentrations in vivo (35–43). Quantification of RA and/or other retinoids would seem essential to elucidate mechanisms by which retinoids contribute to disease. Quantification of retinoids requires attention to proper handling and, in some cases, isomeric distribution. The susceptibility of retinoids to isomerization and oxidation is well documented and necessitates care during sample collection, handling, and storage (44–51). Resolution of isomers is important in RA analyses, as isomers of RA have different affinities for nuclear receptors and, therefore, may afford different biological actions. atRA activates retinoic acid receptors (RAR) (8, 52) and peroxisome proliferator-activated receptor, type β/δ (9, 10). 9-cisRA (9cRA) activates both RAR and retinoid X receptors (RXR) (26). Retinoids may exert additional biological effects through dimerization of RXR with an array of type II nuclear receptors, such as thyroid hormone, peroxisome proliferator-activated, vitamin D, liver X, farnesoid X, pregnane, constitutively activated, and the small nerve growth factor-induced clone B subfamily of nuclear receptors (26). 13-cis-RA (13cRA) does not activate RAR or RXR directly but induces dyslipidemia and insulin resistance, most likely through conversion into atRA (35, 53–56). Tissues and serum contain 9,13-di-cis-RA (9,13dcRA), which may reflect conversion from 13cRA and/or 9cRA, and does not activate RAR or RXR (57–59). The disparity in endogenous abundance often demands attention to analytical methodology. RE storage levels (∼high micromolar) can differ by as much as six orders of magnitude from endogenous RA levels (∼low nanomolar) (49–51). Previously, analytical limitations of direct RA quantification have hindered the complete investigation of retinoid metabolism essential for understanding retinoid function and its relationship to disease risk. A number of recent analytical efforts have provided assays with the necessary sensitivity, specificity, and/or isomeric resolution to quantify endogenous RA levels (and other retinoids) in tissue and serum, and thus, indirect methods of quantification should be avoided (48–51, 60, 61).

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Indirect methods used as substitutes for direct RA measurement and analytically robust assays, such as in vitro reporter assays or transgenic RA reporter mouse strains, lack specificity, lack means of quantification, and/or have produced contradictory results (62–64). These non-instrumental methods, based on reporter gene expression, have not been developed into analytically rigorous assays, are not specific for all-trans-RA (e.g., 3,4didehydro-RA, 9cRA, 4-oxo-RA, 4-hydroxy-RA, and 4-hydroxyretinol all produce signals), are not quantitative, and can give both false-positive and false-negative results (63–65). Additionally, because reporter detection systems reflect RAR activation, they cannot evaluate retinoid presence in real time and may reflect the longer term consequences of receptor activation, after the retinoid has been catabolized. Administration of a super-physiological dose of retinol to raise RA levels to a level detectable by UV absorbance is also problematic. An example of this approach dosed as much as 50 mg/kg, ∼300-fold greater than the recommended daily intake of retinol for a mouse (30, 66, 67). Superphysiological doses such as this induce an artificial environment where serum atRA levels are raised ∼1600-fold higher than typical steady-state values of ∼2.5 pmol/ml, likely overwhelming normal metabolism and eliciting retinoid toxicity responses (30, 49, 50, 66, 67). Each of the retinoid detection methods described in the literature, including LC-MS/MS, LC-MS, HPLC-UV, GC-MS, and ECD, has different sensitivities, effectiveness with various biological matrices, benefits, and limitations. A comparison of these methodologies is provided (see Table 1.1). HPLC with UV detection has the benefit of ease and economics, but has

Table 1.1 Comparison of atRA limits of detection (LOD) for validated assays References

atRA LODa

Detection

Assay application/ demonstrated matrices

Napoli (45)

120 fmol

GC/MS

Serum, plasma, cells

Wang et al. (60)

(∼211 fmol)b

LC/MS

Prostate

Schmidt et al. (48)

186 fmol

LC/UV

Tissues, serum

Sakhi et al. (71)

26.6 fmol

LC/ECD

Tissues (embryo)

Ruhl (79)

23.3 fmol

LC/MS/MS

Serum, cell pellets (no tissues)

Kane et al. 2005 (49)

10 fmol

LC/MS/MS

Tissues, serum, cells

Gundersen et al. (61)

6.6 fmol

LC/MS/MS

Plasma only

Kane et al. 2008 (50)

0.5 fmol

LC/MS/MS

Tissues, serum, cells

a atRA LOD expressed as mol on column and defined as S/N=3 b Estimated based on a listed LOQ of 702 fmol (30)

Quantification of Endogenous Retinoids

5

LOQ ∼0.4–1 pmol and does not provide mass identification (45, 51, 68). Recent advances in column technology and column switching capabilities have assisted in lowering detection limits (48, 69). HPLC with electrochemical detection has sensitivity in the femtomolar range, but lacks the definite mass identification of analytes of MS, is subject to interference from other analytes, and has solvent/electrode/flow-dependent sensitivity (70–73). GC/MS affords sensitivity, with a lower limit of detection <250 fmol, but GC/MS requires derivatization for RA detection (74, 75). LC/MS-based assays offer mass identification of analytes, but do not have the potential sensitivity or the enhanced specificity of selected reaction monitoring (SRM) (76–79). Triple-quadrupole LC/MS/MS offers the most effective RA detection at this time with sensitivity, specificity, no requirement for derivatization, and definite mass identification (49, 50). Future directions for retinoid analysis may include utilization of greater sensitivity MS/MS instrumentation, incorporation of high-throughput methodology (61), quantification of localized areas and/or individual cellular populations (e.g., areas isolated by laser capture microdissection or subcellular fractionation) (80, 81), and simultaneous quantification of retinoids with other biologically significant metabolites to enable retinoid pathway analysis in concert with the evaluation of other metabolic pathways (82). Described here are all procedures needed for accurate and reliable quantification of retinoids, including detailed descriptions of how to properly handle retinoids, prepare standard solutions, harvest tissues, collect samples, homogenize, extract, separate, and detect retinoids. Strategies for optimizing extraction of retinoids from biomatrices, chromatographic resolution, and detection sensitivity are provided, as are the steps necessary for evaluating the performance of retinoid assays. Retinoid assays described here for mice also have proven applicable to other organisms including zebrafish, rat, rabbit, and human.

2. Materials 2.1. Proper Handling and Transfer of Retinoids, Retinoid Solution Preparation

1. Disposable glass containers (vials, etc.) for solutions (Fisher Scientific) 2. Disposable glass pipettes (Pasteur and graduated) for transfer (Fisher Scientific) 3. Calibrated glass syringes (Hamilton) for measuring small volumes of retinoid solutions 4. –20◦ C freezer for short-term (less than 1–2 weeks) storage of solutions and prepared samples

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Kane and Napoli

5. UV–Vis spectrometer 6. Quartz cuvettes 7. Disposable glass vials/containers 8. Disposable glass Pasteur pipettes 9. Calibrated glass syringes for measuring retinoid volumes (Hamilton) 2.2. Sample Collection

1. Animals should be of similar strain, size, age, diet, and/or fasting state 2. Yellow room lights or darkened room with desk lamp with yellow bulb 3. Dissecting tools 4. Eppendorf tubes or similar to collect tissue 5. Balance to weigh tissue 6. Liquid nitrogen to flash freeze samples 7. –80◦ C freezer for storage of samples (tissue samples can be stored 1–2 weeks with no degradation)

2.3. Homogenization

1. Saline (0.9% NaCl) 2. Ground glass hand homogenizers (Duall, size 22 or similar, Kontes) and/or motorized homogenizer (Heidolph or similar) with ground glass mortar/pestle (Kontes) and/or Polytron stainless steel motorized homogenizer 3. Disposable 1 ml graduated glass pipettes to measure homogenate 4. 16 × 150 mm disposable glass culture tubes 5. UV–Vis spectrometer, disposable polystyrene cuvettes, Bradford reagent (Bio-Rad) for protein determination

2.4. Internal Standard

1. Internal standard solution at concentration approximately similar to analyte detection amount to be delivered to each sample in ∼5–10 μl DMSO, ethanol, acetonitrile, or other compatible solvent

2.5. Conversion of Retinal into Retinal Oxime and Extraction

1. 0.1 M O-ethylhydroxylamine in 0.1 M HEPES, pH 6.5 2. Hexane 3. 16 × 150 mm disposable glass culture tubes 4. Vortex mixer 5. Disposable 10 ml glass pipettes 6. Nitrogen gas solvent evaporator (Organomation Associates Inc. model N-EVAP 112, Berlin, MA) or similar with disposable gas delivery tips 7. Disposable 53/4 glass Pasteur pipettes for nitrogen gas solvent evaporator gas delivery tips

Quantification of Endogenous Retinoids

2.6. RA/Retinol/RE/Polar Metabolite Extraction and Resuspension

7

1. 1. 0.025 M KOH in 100% ethanol 2. 4 M HCl 3. Internal standard solutions (make approximately same concentration as analyte of interest) 4. Hexane 5. 16 × 150 mm disposable glass culture tubes 6. Disposable 10 ml glass pipettes 7. Vortex mixer 8. Dynac centrifuge (Becton Dickinson) or similar for 16 × 150 mm culture tubes 9. Nitrogen gas solvent evaporator (Organomation Associates Inc. model N-EVAP 112, Berlin, MA) or similar with disposable gas delivery tips 10. Disposable 53/4 glass Pasteur pipettes for nitrogen gas solvent evaporator gas delivery tips 11. Ice bucket(s) and ice 12. Resuspension solvent(s): acetonitrile, hexane with 0.4% IPA, and/or retinal isomer mobile phase 13. Disposable 9 glass Pasteur pipettes for transfer of resuspended samples to inserts 14. Limited volume glass inserts for HPLC vials (available from Agilent, Waters, etc.)

2.7. Separations 2.7.1. Reverse-Phase Retinoic Acid Separations (See Sections 3.13.1 and 3.13.2)

1. Ascentis alkyl amide C16 column (Supleco): 2.1 × 100 mm, 3 μm (gradient 1), and/or Ascentis alkyl amide C16 column (Supelco): 2.1 × 150 mm, 3 μm (gradient 2) 2. Supelcosil ABZ+PLUS Supelguard (Supelco, 2.1 × 20 mm, 5 μm)

cartridge

column

3. Pre-column filter 4. Acetonitrile with 0.1% formic acid 5. Water with 0.1% formic acid 6. Methanol with 0.1% formic acid 7. Acetonitrile (for column storage) 2.7.2. Normal-Phase Retinoic Acid Separation (See Section 3.13.3)

1. Zorbax SIL column (Agilent) 4.6 × 250 mm, 5 μm

2.7.3. Total Retinal Separation (See Section 3.14.1)

1. Zorbax SB-C18 column (Agilent) 4.6 × 100, 3.5 mm

2. Hexane with 0.4% isopropyl alcohol

2. Acetonitrile 3. Water

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Kane and Napoli

4. Water with 10% formic acid 5. 1,2-Dichloroethane 2.7.4. Retinal Isomer Separation (See Section 3.14.2)

1. Two Zorbax SIL column (Agilent) 4.6 × 250 mm, 5 μm

2.7.5. Total Retinol, Total Retinyl Ester Separation (See Section 3.15.1)

1. Zorbax SB-C18 column (Agilent) 4.6 × 100, 3.5 mm

2. 11.2% ethyl acetate, 2% dioxane, 1.4% 1-octanol, 85.4% hexane

2. Acetonitrile 3. Water 4. Water with 10% formic acid 5. 1,2-Dichloroethane

2.7.6. Retinol Isomer Separation (See Section 3.15.2)

1. Zorbax SIL column (Agilent) 4.6 × 250 mm, 5 μm

2.7.7. Polar Metabolite Separation (See Section 3.16)

1. Zorbax SB-C18 column (Agilent) 4.6 × 100, 3.5 mm

2. Hexane with 0.4% isopropyl alcohol

2. Acetonitrile 3. Water 4. Water with 10% formic acid

2.8. Analysis Instrumentation

1. HPLC with autosampler (Agilent 1200 series or comparable) and triple-quadrupole mass spectrometer (API3000/API-4000, Applied Biosystems, or comparable) for endogenous RA analysis 2. HPLC with autosampler and UV detector (Agilent 1200 series or comparable) for endogenous retinol, retinal, and RE analysis

3. Methods 3.1. Retinoid Handling

1. Handle retinoids under yellow (or red) light ONLY (see Note 1). Retinoid degradation due to light exposure is illustrated in Fig. 1.2. Handling under yellow light includes all procedures of the following: • Standard solution preparation (see Note 2) • Tissue collection/dissection(see Note 3) • Tissue dissection requiring a microscope (see Note 4) • Homogenization • Extraction • Resuspension

Quantification of Endogenous Retinoids

9

2500 2000

A

B

1500

+hν

atRA

1000

Intensity

500 0

800

C

D

600

+hν

4,4-dimethyl-RA

400 200 0

10

12

14

16

18

10 12 20 22 Retention Time (min)

14

16

18

20

22

Fig. 1.2. Similar isomerization rates of atRA (a and b) and 4,4-dimethyl-RA (c and d). (a) and (c) show standard solutions (∼50 nM) exhibiting mild isomerization after 10 min of exposure to standard fluorescent room lights. (b) and (d) show standard solutions with severe isomerization after 30 min exposure to sunlight. Note the similar rate of production of isomers from the all-trans forms for both retinoids (t R ≈16.2 min for atRA; t R ≈19.8 for 4,4-dimethyl-RA) to cis-isomers (t R =10–16 min for RA and 12–18 min for 4,4-dimethyl-RA). Adapted from Ref. (49).

2. Use only glass containers, pipettes, and calibrated syringes to handle retinoids (see Note 5) 3. Clean calibrated syringes before and after use by flushing 15–20 times with acetone and/or ethanol (see Notes 6 and 7) 4. Use fresh pipettes for transfer of materials to prevent crosscontamination among samples. This includes transfer of • Standard solutions • Homogenate • Hexane extracts • Resuspended samples 5. Use fresh pipettes for nitrogen delivery to each sample when evaporating solvent (see Note 8) 6. Clean homogenizers with water followed by ethanol and/or acetone before and after use to remove retinoid residue (see Note 9)

10

Kane and Napoli 0.8 at-retinol

Absorbance (AU)

0.6

atRA

at-retinal

0.4

0.2

0.0 250

300

350 400 Wavelength (nm)

450

Fig. 1.3. Absorbance spectra of select retinoids in ethanol.

3.2. Retinoid Standard Preparation and Quantification

It is inappropriate to prepare and quantify retinoids by weight. Absorption spectroscopy is a simple and rapid way to accurately quantify retinoid concentrations (83). Sample absorbance spectra for several retinoids are shown in Fig. 1.3. 1. Dissolve a small amount of retinoid in the appropriate solvent (see Table 1.2, Note 10). 2. Collect the full absorbance spectrum (250–450 nm) (see Notes 11 and 12). 3. Calculate the concentration using Beer’s law, A=εbc (see Note 13). • A, absorbance at maximum (see maxima for selected retinoids listed in Table 1.2). • ε, molar absorptivity (M−1 cm−1 ). • b, path length (cm). Path length is 1 cm in a standard (1 × 1 cm) cuvette. • c, concentration (M). 4. Dilute or evaporate an appropriate aliquot and re-dissolve in the desired solvent (if necessary).

3.3. Purification of Retinoid Reagents

3.3.1. Chromatographic Purification of Retinol

Commercially available retinol can have retinal contamination (as high as 1–10%) that can influence experimental results when treating cells or subcellular fractions. 1. A concentrated solution of retinol should be prepared in hexane with 5% acetone. 2. Inject retinol and purify using a Zorbax Sil 9.4 × 250 mm column (Agilent) and 5% acetone in hexane at 5 ml/min.

Quantification of Endogenous Retinoids

11

Table 1.2 Optical properties of select retinoidsa,b,d Compound

Solvent

λmax

ε

E1% 1 cm

References

All-trans-retinol

Ethanol

325

[52,770]

1845

(84)

Hexane

325

[51,770]

1810

(84)

9-cis-retinol

Ethanol

323

42,300

[1477]

(85)

9,13-di-cis-retinol

Ethanol

324

39,500

[1379]

(85)

13-cis-retinol

Ethanol

328

48,305

1689

(85, 86)

11-cis-retinol

Ethanol

319

[34,890]

1220

(84)

Hexane

318

[34,320]

1200

(84)

All-trans-retinyl acetate

Ethanol

325

51,180

1560

(87)

Hexane

325

[52,150]

1590

(88)

All-trans-retinyl palmitatec

Ethanol

325

[49,260]

940

(88)

All-trans-retinal

Ethanol

383

[42,880]

1510

(84)

Hexane

368

[48,000]

1690

(84)

9-cis-retinal

Ethanol

373

36,100

[1270]

(86)

9,13-di-cis-retinal

Ethanol

368

[32,380]

1140

(89)

13-cis-retinal

Ethanol

375

[35,500]

1250

(84)

Hexane

363

[38,770]

1365

(84)

Ethanol

380

[24,935]

878

(84)

Hexane

365

[26,360]

928

(84)

All-trans-retinoic acid

Ethanol

350

45,300

[1510]

(85)

9-cis-retinoic acid

Ethanol

345

36,900

[1230]

(85)

9,13-di-cis-retinoic acid

Ethanol

346

34,500

[1150]

(85)

13-cis-retinoic acid

Ethanol

354

39,750

[1325]

(85)

11-cis-retinoic acid

Ethanol

342

[27,780]

926

(89)

11-cis-retinal

4-oxo-retinoic acid 4-oxo-13-cis-retinoic acid

Ethanol

360

[58,220]

1854

(90)

Hexane

350

[54,010]

1720

(90)

Ethanol

361

[39,000]

1242

(91)

λmax , maximum wavelength (nm) ε, molar absorptivity (M−1 cm−1 ) E, mass attenuation coefficient (ml g−1 cm−1 ) a Adapted from Furr (83) b Values in brackets are calculated from corresponding ε or E 1% values 1 cm c Medium- and long-chain fatty-acyl esters of retinol have identical molar extinction coefficients (ε) (92) d ε and E 1% values are given for the maximum wavelength 1 cm

3. Collect retinol which should elute at ∼20 min (± depending on acetone concentration). 4. Evaporate solvent (see Note 14) and resuspend in a solvent of choice (see Table 1.2).

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5. Determine the concentration of the solution by measuring its absorbance and using Beer’s law (see Section 3.2). 6. Dilute and/or evaporate with nitrogen and resuspend in another solvent (if necessary). 3.3.2. Retinol Purification by Reduction of Retinal

Retinal can be readily reduced to retinol by NaBH4 and the excess NaBH4 can be neutralized by reaction with acetone to form 2propanol. 1. Mix retinol (with retinal contamination) and NaBH4 in THF with 5% MeOH (see Note 15) with NaBH4 in excess of retinal (at least ∼10-fold excess of estimated retinal level). 2. After 15–30 min of reaction time, add an excess of acetone to react with the added amount of NaBH4 (forming 2-propanol). 3. Blow off solvents with nitrogen or another inert gas. 4. Wash the hexane layer by resuspending in H2 O/hexane (∼1:1), vortex – mixing, and removing the (upper) hexane layer. Keep the hexane layer. 5. Add more H2 O to the hexane layer and repeat washing of hexane layer at least three times. 6. Quantify retinol concentration by absorbance spectroscopy (see Section 3.2). 7. Dilute and/or evaporate with nitrogen and resuspend in another solvent (if necessary).

3.4. Tissue Sample Preparation

3.4.1. Tissue Harvest/Sample Collection

There are a number of factors which can affect endogenous retinoid levels that should be controlled when assaying animals: (1) dietary vitamin A, (2) age, and (3) fasting. (1) Dietary vitamin A. Dietary vitamin A affects retinoid levels. Standard lab chow has a copious amount of vitamin A (∼15–30 IU vitamin A/g). Copious dietary vitamin A mutes or eliminates changes in endogenous retinoids due to genetic manipulation (or other treatment). Diet with a sufficient level of vitamin A (4IU vitamin A/g) is more appropriate for studies of endogenous retinoids (66). Mice often need to be bred more than one generation on a diet of 4IU vitamin A/g to stabilize endogenous retinoid levels. (2) Age. Mice should be age matched within a group and similar between compared groups. Age ± 0.5–1 month is usually acceptable. (3) Fasting. Metabolic state can affect retinoid levels. Groups of compared mice should be treated identically in terms of fasting/not fasting. 1. Harvest tissue under yellow lights (see Notes 1 and 3 and Section 3.1). If a dissecting microscope is used, a yellow filter should be used with the light source (see Note 4).

Quantification of Endogenous Retinoids

13

2. Euthanize mice according to institutional guidelines. 3. Collected blood by cardiac puncture using a 281/2-guage needle or similar. Allow blood to clot on ice for ∼30 min and then centrifuge at 10,000×g for 10 min (preferably at 4◦ C) to isolate serum. Draw off serum using a 1 ml calibrated glass pipette to measure the volume. Serum can be flash frozen in liquid nitrogen and stored at −80◦ C until assay. 4. Dissect and weigh tissue, place in a microcentrifuge tube, and flash freeze in liquid nitrogen. Store tissues at −80◦ C until assay. 5. If tissue sample is too small to be weighed accurately (localized region, embryo, cells), protein content can be determined and retinoid expressed per gram protein instead of per gram tissue (see Section 3.5.3). 6. Typical tissue harvest amounts from adult mice are listed in Table 1.3. Amounts provide enough tissue in most cases for multiple analyses. Small sample sizes may require pooling. Amount of tissue needed for an analysis should be tested and optimized (see Section 3.11). 7. Measurements from approximately 10–20 mg of tissue produce rigorous data from limited samples (e.g., embryo, localized areas, cells), whereas routine tissue assay amounts from adult mice range from approximately 40–115 mg. 3.4.2. Tissue Storage and Retinoid Stability During Storage

RA is susceptible to degradation even if stored at −80◦ C. Precautions must be taken to assure sample handling prior to analysis does not produce artifactual changes in endogenous retinoid levels. More abundant endogenous retinoids, such as retinol and RE, are less susceptible to but not devoid of storage-induced degradation. 1. Keep samples frozen as tissue until immediately before homogenization. 2. Collect and flash freeze in liquid nitrogen as described in Section 3.4 under yellow lights. 3. Shield from light exposure at all times during storage. 4. Tissues can remain in liquid nitrogen in an appropriate storage container until assay or remove samples from liquid nitrogen and place at −80◦ C until assay. 5. Analysis within several days is preferable, but is possible for up to 1–2 weeks without significant degradation. Same day or next day analysis of frozen samples produces the highest quality data. 6. Most samples stored over 1 month may show measurable loss, especially of RA.

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Table 1.3 Typical tissue harvest amounts from adult mouse∗∗ Tissue harvest amount

Volume of saline for homogenization

Extract amount for RA/retinol/RE

Extract amount for retinal

Blood (isolated serum)

200–600 μl (100–300 μl)

N/A None

N/A 100–300 μl

N/A 100–300 μl

Liver

0.2 g

2 ml

0.5–0.75 ml

0.5–1 ml

Kidney

1–2 kidney

1–2 ml

0.75–1 ml

0.5–1 ml

Testis

1–2 testis

1–2 ml

1 ml

0.25–1 ml

White adipose

0.2 g

1–1.5 ml

0.75–1 ml

0.75–1 ml

Brown adipose

0.1–0.2 g

1 ml

1 ml

–

Muscle

0.1–0.2 g

1 ml

1 ml

–

Skin

0.3–0.4 g

1–1.5 ml

1–1.5 ml

–

Spleen

Whole spleen

1 ml

1 ml

–

Pancreas

Whole pancreas

1 ml

0.3–1 ml

0.3–1.0 ml

Mammary gland Retinae

0.1–0.2 g Both retinae, pool ∼4 retinae

1–1.5 ml 1 ml

1 ml 1 ml

– –

Brain (whole)

Entire brain

2 ml

0.5–0.75 ml

–

Hippocampus

2 hippocampi

1 ml

1 ml

–

Cortex

0.1–0.2 g

1–1.5 ml

0.75–1 ml

–

Cerebellum

Whole region

1 ml

1 ml

–

Striatum

Whole region

1 ml

1 ml

–

Thalamus

Whole region

1 ml

1 ml

–

Olfactory bulb

Whole region

1 ml

1 ml

–

Hypothalamus

Whole region, pool 2–3 mice

1 ml

1 ml

–

Tissue

–, not determined N/A, not applicable ∗∗ Amounts listed are guidelines. Extract conditions should be optimized for each tissue. See Section 3.11 for optimization strategies

7. Test stability of each sample/tissue type, if storage before analysis will take place (see Note 16). 8. Do not store homogenates (see Note 17). 9. Generally, do not freeze, thaw, and then refreeze tissues (unless stability has been verified). 10. Thaw all samples (regardless of type) on ice.

Quantification of Endogenous Retinoids

3.5. Tissue Homogenization and Preparation

3.5.1. Tissue Homogenization

15

Retinoids are susceptible to degradation during homogenization. Use the most gentle homogenization possible. Keep all samples on ice during homogenization. Homogenization methods, starting with the most gentle, include (1) hand homogenization with a ground glass homogenizer, (2) motorized homogenizer driving the pestle with a ground glass homogenizer, and (3) a Polytron motorized hand-held stainless steel homogenizer. Most tissues can be homogenized by hand. Very fibrous tissues may need motorized pestle or Polytron. 1. Perform all homogenization under yellow lights (see Note 3). 2. Clean homogenizers before use (see Section 3.1). 3. Homogenize on ice with ice-cold saline (0.9% NaCl) to make an approximately 10–25% homogenate (for approximate amounts of saline for different tissue types, see Table 1.3). 4. When using a hand homogenizer, use a slow motion and use as few strokes as possible to achieve a homogenate. When using a Heidolph homogenizer (or similar motor-driven homogenizer), use approximately five strokes at lowest possible speed that will achieve homogenization. When using a Polytron, use the lowest possible speed to achieve homogenization (∼20–25% of maximum) pulsing the homogenizer on and off in ∼5–10 bursts of short duration (see Notes 18, 19, and 20). 5. Aliquot homogenate with graduated 1 ml glass pipettes into 16 × 150 mm culture tubes. 6. Extract samples immediately after homogenization (see Notes 17 and 21).

3.5.2. Subcellular Fractionation

Retinoids can be quantified in subcellular fractions. 1. Perform all procedures under yellow lights (see Note 3). 2. Harvest tissue as described in Section 3.4. 3. Tissues should be homogenized (to make ∼25% homogenate) in 10% sucrose, 10 mM Tris-HCl, 1 mM EDTA, 1.5 mM DTT, pH 7.4 at 1240 rpm in a Dounce homogenizing vial, Kontes, size 22. 4. Isolate subcellular fractions at 4◦ C by ultracentrifugation. 5. Freeze fractions at −80◦ C until assay (see Note 17). 6. Determine protein content as described in Section 3.5.3.

3.5.3. Protein Determination

If sample is too small to be weighed (e.g., localized area, embryo parts, cells, subcellular fraction), determine the total protein

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content of the sample as homogenate using the Bradford method or another method to assess total protein content. In this case, retinoids will be expressed as retinoid per gram protein instead of retinoid per gram tissue. Bradford assay is a dye-binding assay used to measure total protein content, where absorbance of the protein–dye solution is proportional to protein amount (see Notes 22, 23, and 24). 1. Dilute Bradford reagent with water immediately before use so that it is 20% concentrated Bradford reagent. 2. Aliquot 1 ml diluted Bradford reagent into disposable polystyrene cuvettes. 3. Add 20 μl protein to diluted Bradford reagent, mix well, and let react for 5–10 min. 4. Measure absorbance at 595 nm (see Note 25). 5. Calculate protein concentration from a standard curve. Use protein concentration to determine total gram of protein in homogenate aliquot. 3.6. Internal Standards

An internal standard improves accuracy by establishing extraction efficiency, handling loss, and revealing handling-induced isomerization/degradation. An internal standard should have similar chemical properties comparable to the analyte of interest, including structure, extraction efficiency, and chromatographic behavior. More than one internal standard may be added to follow multiple analytes. 1. Establish performance of chosen internal standard before use (see Section 3.22.5). 2. Internal standard solutions are prepared (see Sections 3.1 and 3.2) at a concentration that delivers an amount comparable to the analyte of interest. 3. Addition volume is usually 5–20 μl, delivered in ethanol, acetonitrile, or DMSO. 4. Internal standard(s) are added to the homogenate with a calibrated glass syringe and the sample is vortex mixed (see Notes 26 and 27). 5. Samples are then extracted as described in Section 3.8.

3.7. Conversion of Retinal to Retinal Oxime Derivatives

(O-ethyl)hydroxylamine is used to convert retinal into a stable oxime for accurate quantification (see Fig. 1.4). Retinal has

CHO retinal

+ CH3CH2ONH2 O-ethylhydroxylamine

Fig. 1.4. Conversion of retinal into retinal-(O-ethyl)oxime.

N-OC2H5 retinal (O-ethyl) oxime

Quantification of Endogenous Retinoids

17

a reactive aldehyde group susceptible to promiscuous reactions in the sample matrix, preventing efficient extraction. Reaction with hydroxylamine or (O-alkyl)hydroxylamines produces syn- and anti- isomers (93–95). It is more desirable to generate retinal oximes from (O-alkyl)hydroxylamines (e.g., (O-ethyl)hydroxylamine) instead of the nonalkylated hydroxylamine because the anti-retinaloxime isomer of the nonalkylated hydroxylamine tends to co-elute with retinol and/or elute as a broad asymmetrical peak in both reverse and normal-phase HPLC. Additionally, the variable contribution from the synretinaloxime prevents retinal quantification with only the synisomer. 1. Perform all procedures under yellow lights (see Note 3). 2. Collect and homogenize tissues as described in Sections 3.4 and 3.5 and aliquot homogenate (see Table 1.3) into 16 × 150 mm disposable glass culture tubes. 3. To an appropriate amount of homogenate or serum (see Table 1.3), add 1–2 ml methanol and 0.5–2 ml of 0.1 M O-ethylhydroxylamine in 0.1 M HEPES (pH 6.5). 4. Vortex samples well and let stand for 15 min at room temperature. 5. Extract retinal with hexane (see Section 3.8.4). 3.8. Extraction

3.8.1. Acid–Base Extraction: Retinol, RE, RA, and Polar Metabolites

Most samples are extracted with a two-step acid–base extraction that recovers multiple retinoids: RA, retinol, RE, and RA polar metabolites. Retinal extraction requires pre-treatment to convert reactive retinal to a stable product (see Section 3.7). Extractions can be altered to target only one type of analyte, if desired. 1. Carry out all procedures under yellow lights (see Note 3). 2. Collect samples and homogenize as described (see Sections 3.4 and 3.5). 3. Aliquot homogenate into 16 × 150 mm disposable glass culture tubes (see Table 1.3). 4. Determine total protein content (if necessary) before extraction (see Section 3.5.3). 5. Add internal standard to homogenate/serum and vortex mix (see Section 3.6). 6. Add from 1 to 3 ml of 0.025 M KOH in ethanol to tissue homogenates/serum and vortex mix (at least 10 s). 7. If sample requires addition of acetonitrile to facilitate protein precipitation, add acetonitrile here (usually ∼1 ml acetonitrile) and vortex mix (at least 10 s). 8. Add 10 ml of hexane to homogenate/serum and vortex mix (at least 10 s).

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9. Centrifuge for 1–3 min at ∼1,000 × g to facilitate phase separation. 10. Draw off the top (organic) phase containing nonpolar retinoids (retinol and retinyl ester(s)) to a new 16 × 150 mm disposable glass culture tube (see Note 28). 11. To the bottom (ethanolic aqueous) layer of homogenate/ serum, add 4 M HCl (60–240 μl) and vortex mix (at least 10 s). 12. Add 10 ml hexane and vortex mix (at least 10 s). 13. Centrifuge for 1–3 min at ∼1,000 × g to facilitate phase separation. 14. Draw off top (organic) phase containing RA and polar retinoids to a new 16 × 150 mm disposable glass culture tube. It is important not to disturb the bottom acidcontaining layer! Any of the acid-aqueous layer drawn off with the hexane will destroy RA and/or cause major isomerization (see Fig. 1.5). 15. Evaporate organic phases under nitrogen with gentle heating at ∼25–30◦ C in a water bath (see Note 28). 16. Keep evaporated samples on ice until resuspension (see Section 3.9, resuspension). 3.8.2. RA-Only Extraction from Serum

If only RA quantification from serum is desired, a one-step acetonitrile extraction is possible. 1. Follow Section 3.8.1, acid–base extraction: steps 1, 2 and 3. 2. Add 1 ml of acetonitrile and 60 μl of 4 M HCl and vortex mix (at least 10 s). 3. Add 10 ml of hexane and vortex mix. 4. Centrifuge for 1–3 min at ∼1,000 × g to facilitate phase separation. 5. Draw off top (organic) phase containing RA and polar retinoids to a new 16 × 150 mm disposable glass culture tube. It is important not to disturb the bottom acid-containing layer! Any of the acid-aqueous layer drawn off with the hexane will destroy RA and/or cause major isomerization (see Fig. 1.5). 6. Evaporate organic phases under nitrogen with gentle heating at ∼25–30◦ C in a water bath (see Note 28). 7. Keep evaporated samples on ice until resuspension (see Section 3.9).

3.8.3. Retinol and RE Extraction

If only retinol and RE are desired for quantification, use Section 3.8.1, acid–base extraction: omitting steps 8, 9, 10, and 11.

Quantification of Endogenous Retinoids 1600

A

2500

2

1200 1000

1500 1000 500 0

800

14

600 400

16 18 20 Ret. Time (min)

5

4

6

200 Intensity

1

2000 Intensity

1400

19

3

0 1600

2500

B Intensity

1400 1200 1000

2000 1500

1

1000 500 0

800

14 16 18 20 Ret. Time (min)

600

5

6

400

4 3

200

2

0 10

12

14 16 18 Retention Time (min)

20

22

Fig. 1.5. Application of internal standard with mouse kidney illustrating acid-induced isomerization. (a) Chromatogram for atRA with correct sample handling. Inset: 4,4dimethyl-RA (internal standard) shows only all-trans form. (b) Chromatogram for atRA illustrating handling-induced isomerization by acid contamination. Inset: cis-isomers (arrows) of 4,4-dimethyl-RA. Note the decrease in atRA and increase in cis-isomers concurrent with cis-isomers in the internal standard. Peak identities: (1) 4,4-dimethylRA, (2) atRA, (3) 9cRA, and (4) 13cRA. Peaks 5 and 6 most likely indicate 9,13-di-cis-RA and 11-cis-RA, respectively. Adapted from Ref. (49).

3.8.4. Retinal (Oxime) Extraction

Retinal oxime derivatives are extracted efficiently with hexane (>95% recovery in most cases). 1. Follow Section 3.7 to prepare retinal (O-ethyl) oxime derivatives. 2. Add 10 ml hexane to homogenate and vortex mix (at least 10 s). 3. Centrifuge for 1–3 min at ∼1,000 × g to facilitate phase separation (see Note 29). 4. Draw off top (organic) layer to a new 16 × 150 mm disposable glass culture tube. 5. Evaporate organic phases under nitrogen with gentle heating at ∼25–30◦ C in a water bath (see Note 28). 6. Keep evaporated samples on ice until resuspension (see Section 3.9).

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Kane and Napoli

Several factors are important to consider when selecting a resuspension solvent for analysis of retinoids. These include (1) solubility of retinoid in resuspension solvent, (2) compatibility of resuspension solvent with HPLC mobile phase, (3) volume of resuspension solvent necessary to make amount of analyte in injection volume appropriate to linear range of the analysis method, and (4) stability of sample in resuspension solvent. 1. Perform all procedures under yellow lights (see Note 3).

3.9. Resuspension

2. Extract and evaporate samples (see Section 3.8). 3. Add appropriate volume of solvent for the desired analyte and separation method (see Tables 1.4 and 1.5 and Notes 30, 31, 32, and 33). 4. Vortex mix for 10–20 s. 5. Transfer sample with a 9 Pasteur pipette to a low-volume glass insert for analysis. 6. Analyze according to analyte (see Sections 3.12–3.21).

1. Place samples in amber vials and keep shielded from light.

3.10. Extracted Sample Storage

2. Samples in acetonitrile are stable at room temperature for 2 days; at 4 days a 20% loss occurs (at room temperature) (49). Cooling of the autosampler helps preserve sample quality.

Table 1.4 Typical resuspension and injection volumes∗∗

Tissue type

RA resuspend (µl)

RA inject (µl)

Retinol RE resuspend (µl)

Retinol RE inject (µl)

Retinal resuspend (µl)

Retinal inject (µl)

Serum/plasma

60

20–30

120

100

120

100

Livera

60

20–30

500

100 and 10–20d

120

100

Adipose and high lipid content tissuesb

60

20–30

200

100

150–200

100

All other tissues

60

20–30

120

100

120

100

Small samplesc

40–60

20–30

110–120

100

110–120

100

a See Note 32 b See Note 33 c See Note 35 d See Note 41 ∗∗ Volumes listed are guidelines. Extract conditions should be optimized for each tissue

Quantification of Endogenous Retinoids

21

Table 1.5 Typical resuspension solvents∗∗ Separation method

Sections

Resuspension solvent

RA isomers (gradient 1)

3.13.1

Acetonitrile

RA isomers (gradient 2)

3.13.2

Acetonitrile

RA isomers (normal phase)

3.13.3

Hexane with 0.4% isopropyl alcohol

Total retinal

3.14.1

Acetonitrile

Retinal isomers

3.14.2

Retinal isomer mobile phase (11.2% ethyl acetate, 2% dioxane, 1.4% 1-octanol, 85.4% hexane)

Total retinol and total RE

3.15.1

Acetonitrile

Retinol isomers

3.15.2

Hexane with 0.4% isopropyl alcohol

Polar metabolites

3.16.1, 3.16.2

Acetonitrile or acetonitrile/water mixture up to 50% water

∗∗ Solvents listed are guidelines. Extract conditions should be optimized for each tissue

3. If samples are not analyzed immediately, store at −20◦ C (preferable) or 4◦ C. 4. Resuspended samples (in acetonitrile) stored at −20◦ C remain unchanged for ∼5–7 days (49–51). 5. Test stability of each sample type if samples are to be stored for any length of time before analysis (see Note 16). 3.11. Sample Preparation Optimization Strategies

Attention to sample preparation provides dividends in sensitivity and accuracy, especially in the low-femtomole range. The sample preparation described here, although not extensive, purifies the matrix sufficiently to enhance accuracy and the lives of guard and analytical columns. The simpler sample preparations available are adequate only for samples with less complicated matrixes, such as serum from normal subjects, or certain cell culture extracts (61, 71). Most tissues (e.g., liver, kidney, testis) and/or serum from metabolically altered subjects present a more complex matrix, which requires sufficient matrix cleanup to prevent deterioration of assay performance (see Note 34). The following experimental variable should be optimized: 1. Amount of tissue extracted. Small tissues/limited tissue amounts may require pooling of multiple tissue samples for analysis. Large tissue samples/tissues abundant in retinoids require only a fraction of the entire tissue for analysis. 2. Amount of homogenate extracted. Too little extracted will yield low signal, whereas too much sample extracted will yield poor extraction efficiency and/or high background. 3. % homogenate. Dilution of homogenate can improve extraction efficiency. For very small samples it is often easier to

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handle a more dilute sample to minimize transfer losses (addition of 0.5–1.0 ml saline for homogenization). 4. Ratio of extraction reagents. The amount of 0.025 M KOH in ethanol, 4 M HCl, and/or hexane effect extraction efficiency. Tissue homogenate typically needs more extraction reagent(s) than cells or dilute/small samples. 5. Precipitation of protein. 0.025 M KOH in ethanol precipitates some protein. Acetonitrile can be added during the KOH step to assist in precipitating additional protein. Tissues that are protein rich can display higher background and/or interfering peaks in LC/MS/MS chromatograms. 6. Resuspension volume. Resuspension volume can be adjusted according to the abundance of the analyte, the size of the sample, and the type of analysis. See Table 1.4 for starting guidelines. 3.12. Separation Methods and Sample Preparation

The separation methods here are optimized according to analyte. Some separation methods allow quantification of multiple species and/or isomeric forms. Additionally, the sample preparation protocol reported here allows analysis of greater than 5,000–10,000 samples (∼6–12 months) before requiring column replacement. Methods that shorten sample preparation modestly report changing guard columns daily and replacing analytical columns frequently, even with the relatively simple matrix of normal serum, which could be quite costly (61).

3.13. RA Isomer Separations

Two reverse-phase separation protocols were developed to resolve RA and its isomers: one predominantly for cultured cells or subcellular fractions (gradient 1) and one predominantly for tissue samples (gradient 2), which have higher background. An alternate normal-phase separation is also provided.

3.13.1. Cell/Subcellular Fraction RA (Gradient 1)

1. Use a high performance liquid chromatograph (HPLC) consisting of a vacuum degasser, binary pump, temperature-controlled column compartment, and a temperature-controlled autosampler. 2. Maintain the column compartment at ∼20–25◦ C and the autosampler at 10◦ C. 3. Inject 20–30 μL. Inject 30 μl for small/dilute/low abundance samples (see Note 35). 4. Use a Supelcosil ABZ+PLUS Supelguard cartridge column (Supelco, 2.1 × 20 mm, 5 μm) before the analytical column. 5. Use a Supelcosil ABZ+PLUS column (Supelco, 2.1 × 100 mm, 3 μm).

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23

6. Use the following solvents: A, H2 O with 0.1% formic acid; B, acetonitrile with 0.1% formic acid (see Note 36). 7. Gradient 1 separation is effected at 400 μl/min with the following linear gradient: 0–5 min, 60% B to 95% B; 5–8 min, hold at 95% B; 8–9 min, 95% B to 60% B; 9–12 min, re-equilibrate with 60% B. 8. Use MS/MS detection (see Section 3.18). 9. Retention times of RA isomers are as follows: 6.8 min (13cRA), 7.5 min (9cRA), 8.0 min (atRA), and 8.8 (4,4dimethyl-retinoic acid) (see Fig. 1.6, Note 37). 10. Quantify each RA isomer from a calibration curve generated from standard amounts of that isomer using the gradient 1 separation. 3.13.2. Tissue RA (Gradient 2)

1. Use a high performance liquid chromatograph (HPLC) consisting of a vacuum degasser, binary pump, temperature-controlled column compartment, and a temperature-controlled autosampler. 2. Maintain the column compartment at 25◦ C and the autosampler at 10◦ C. 3. Inject 20–30 μl. Inject 30 μl for small/dilute/low abundance samples (see Note 35). 4. Use a Supelcosil ABZ+PLUS Supelguard cartridge column (Supelco, 2.1 × 20 mm, 5 μm) before the analytical column. 5. Use an Ascentis RP-Amide column (Supelco, 2.1 × 150 mm, 3 μm). 6. Use the following solvents: A, H2 O with 0.1% formic acid; B, acetonitrile with 0.1% formic acid (see Note 36). 7. Gradient 2 separation is effected at 400 μl/min with the following linear gradient: 0–3 min, hold at 70% B; 3–15 min, 70% B to 95% B; 15–20 min, hold at 95% B; 20–21 min, 95% B to 70%B; 21–25 min, re-equilibrate at 70% B. 8. Use with MS/MS detection (see Section 3.18). 9. Retention times of RA isomers are as follows: 12.8 min (13cRA), 13.8 min (9cRA), 14.3 min (atRA), and 17.1 (4,4-dimethyl-retinoic acid) (see Fig. 1.6, Note 37). 10. Quantify each RA isomer from a calibration curve generated from standard amounts of that isomer using the gradient 2 separation.

3.13.3. Alternate Normal-Phase Separation

1. Use a high performance liquid chromatograph (HPLC) consisting of a vacuum degasser, binary pump, temperaturecontrolled column compartment, and a temperaturecontrolled autosampler.

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10

COOH

RA(gradient 1) Sub-subsection 3.4.1.1

8 Intensity × 10−3

D

13cRA

6

A all-trans-retinoic acid

atRA

9cRA

4 2

COOH

0

13-cis-retinoic acid

4

Intensity × 10−4

2.5

6

8

10

12

RA (gradient 2) Sub-subsection 3.4.1.2

2

B

atRA 13cRA 9cRA

1.5 1

COOH

9-cis-retinoic acid

0.5 0

Absorbance × 103 (350 nm)−3

12 1.2

14

16

18

RA (normal phase) Sub-subsection 3.4.1.3

1.0

COOH

C

9,13-di-cis-retinoic acid

13cRA

0.8

atRA

0.6

COOH

0.4 9cRA

0.2 all-trans-4,4-dimethyl-retinoic acid

0.0 5

7

9 11 13 Retention Time (min)

15

17

Fig. 1.6. RA isomer separations and structures. (a, b) SRM chromatograms of standard solutions: (a) gradient 1, cultured cell/subcellular fraction protocol; (b) gradient 2, tissue protocol. The solid line corresponds to m/z Q1:301/Q3:205 for RA, and the broken line corresponds to m/z Q1:329/Q3:151 for 4,4-dimethyl-RA. 9,13-dcRA elutes between 9cRA and 13cRA (data not shown). (c) HPLC/UV chromatograms of standard solutions monitored at 340 nm. (d) Structures of atRA, its isomers, and the internal standard 4,4-dimethyl-RA. Reprinted with permission from Ref. (50). Copyright © 2008, American Chemical Society; see Section 3.13.

2. Maintain the column compartment at 25◦ C and the autosampler at 10◦ C. 3. Inject 100 μl (for UV detection or less if using MS/MS detection). 4. Use a Zorbax SIL, 4.6 × 250 mm, 5 μm column. 5. Use isocratic 0.4% 2-propanol/hexane at 2 ml/min (see Note 38).

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25

6. A representative chromatogram with HPLC/UV detection at 340 nm is shown; however, separation could be adapted to MS/MS detection (see Fig. 1.6). 7. Retention times of RA isomers are as follows: 10.9 min (13cRA), 12.1 min (9cRA), and 13.1 min (atRA) (see Note 37). 8. Quantify each RA isomer from a calibration curve generated from standard amounts of that isomer using the normalphase separation (see Note 39). 3.14. Retinal

3.14.1. Total Retinal

Retinal quantification requires summing the syn- and the anti-(Oethyl)retinaloxime forms in chromatograms (of samples prepared and extracted as described in Sections 3.7 and 3.8). A reversephase separation quantifies total retinal, whereas a normal-phase separation can separate cis- and trans-isomeric forms of retinal (O-ethyl) oximes. 1. Use a high performance liquid chromatograph (HPLC) consisting of a vacuum degasser, binary pump, temperaturecontrolled column compartment, and a temperaturecontrolled autosampler. 2. Maintain the column compartment at 25◦ C and the autosampler at 10◦ C. 3. Inject 100 μl. 4. Use a Zorbax SB-C18, 4.6 × 100 mm, 3.5 μm column. 5. Analytes are separated at 1 ml/min with a linear gradient from 40% H2 O/60% acetonitrile/0.1% formic acid to 5% H2 O/95% acetonitrile/0.1% formic acid over 5 min. Final conditions were held for 9 min (see Notes 37, 40, and 41). 6. Use UV detection at 368 nm. Retinol can be monitored simultaneously at 325 nm. 7. Retinal (O-ethyl) oximes elute at 6.6 min (anti-) and 10.9 min (syn-), and retinol elutes at 7.2 min (see Fig. 1.7, Note 37). 8. The sum of the syn- and the anti- oximes is used to quantify retinal (O-ethyl) oxime from a calibration curve generated from standard amounts of retinal (O-ethyl) oxime.

3.14.2. Retinal cis- and trans-Isomers

1. Use a high performance liquid chromatograph (HPLC) consisting of a vacuum degasser, binary pump, temperaturecontrolled column compartment, and a temperaturecontrolled autosampler. 2. Maintain the column compartment at 25◦ C and the autosampler at 10◦ C. 3. Inject 100 μl.

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0.03 0.01

0.10 Total Retinal Sub-subsection 3.4.2.1

Retinol

Retinal (anti-)

0.0015

008 Retinal (syn-)

0.06

0.0010

0.04

0.0005

0.02

0.0000

0.00 0.0

2.5

5.0 7.5 Retention Time (min)

Absorbance (325 nm)

Absorbance (368 nm)

0.05

10.0

Fig. 1.7. Retinal separation. HPLC/UV chromatograms of standard solutions using the retinal oxime method showing the anti-retinal-(O-ethyl)oxime (6.6 min), retinol (7.2 min), and the syn-retinal-(O-ethyl)oxime (10.9 min). Solid line, left axis; broken line, right axis. Reprinted from Ref. (51). Copyright © 2008, with permission from Elsevier; see Section 3.14.

4. Use two (2) Zorbax SIL, 4.6 × 250 mm, 5 μm columns, connected in series. 5. Use 11.2% ethyl acetate, 2% dioxane, 1.4% 1-octanol in hexane at 1 ml/min flow rate (see Note 38). 6. Use UV detection at 325 nm. 7. Representative chromatograms can be found in Furr (81) and in Landers and Olson (92) (see Note 37). 8. The sum of the syn- and anti- oximes from each isomer is used to quantify retinal (O-ethyl) oxime from a calibration curve generated from standard amounts of retinal (O-ethyl) oxime. 3.15. Retinol and RE

3.15.1. Total Retinol and RE

The total retinol and RE method is a reverse-phase separation that is effective with tissue quantification, as well as cell systems and subcellular fractions. The retinol isomer method is an isocratic normal-phase separation that can quantify the isomeric distribution of retinol, which may be of interest when investigating precursors to RA isomers. 1. Use a high performance liquid chromatograph (HPLC) consisting of a vacuum degasser, binary pump, temperaturecontrolled column compartment, and a temperaturecontrolled autosampler. 2. Maintain the column compartment at 25◦ C and the autosampler at 10◦ C. 3. Inject 100 μl. Perform a separate 10 μl injection for liver RE quantification to ensure that the RE signal occurred within the linear detection range (see Note 41).

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27

4. Use a guard column or inline filter. 5. Use a Zorbax SB-C18, 4.6 × 100 mm, 3.5 μm column (Agilent). 6. Separate analytes at 1 ml/min with 11% H2 O/89% acetonitrile/0.1% formic acid for 9 min, followed by a linear gradient over 2 min to 100% acetonitrile. Then maintain 100% acetonitrile for 2 min, followed by a linear gradient over 2 min to 5% acetonitrile/1,2-dichloroethane. Hold final conditions for 2 min before returning to initial conditions (see Notes 42 and 43). 7. Use with UV detection at 325 nm. 8. Retention times for analytes: retinol at 4.8 min, retinyl acetate (IS) at 8.9 min, and RE (shown as retinyl palmitate) at 16.5 min (see Fig. 1.8, Notes 37 and 44). 9. Quantify retinol and RE from calibration curves generated from standard amounts of retinol and RE separated using the total retinol and RE method (see Note 45). 1. Use a high performance liquid chromatograph (HPLC) consisting of a vacuum degasser, binary pump, temperaturecontrolled column compartment, and a temperaturecontrolled autosampler.

3.15.2. Retinol Isomers

2. Maintain the column compartment at 25◦ C and the autosampler at 10◦ C. 3. Inject 100 μl. Perform a separate 10 μl injection for liver RE quantification to ensure that the RE signal occurred within the linear detection range (see Note 41). 4. Use a Zorbax SIL, 4.6 × 250 mm, 5 μm column. 5. Resolve analytes using 2 ml/min (see Note 38).

2-propanol/hexane

at

0.07

0.05

Total Retinol/RE

RE

Sub-subsection 3.4.3.1

0.04 0.03 0.02

Retinol 0.01 0.00

B

Retinol isomers

A Absorbance (325 nm)

Absorbance (325 nm)

0.4%

Sub-subsection 3.4.3.2

0.05 0.03

RE REA

at-retinol 9c-retinol

0.01 0.002

13c-retinol

0.001 0.000

0

5

10

15

Retention Time (min)

20

25

0

10

20

30

Retention Time (min)

Fig. 1.8. Retinol and retinol isomer separations. HPLC/UV chromatograms of standard solutions using (a) the total retinol/total RE method and (b) the retinol isomer method. The RE standard shown is retinyl palmitate and REA is retinyl acetate (internal standard). Reprinted from Ref. (51). Copyright © 2008, with permission from Elsevier, see Section 3.15.

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6. Use UV detection at 325 nm. 7. Retention times for retinol isomers: 20.9 min (13c-retinol), 27.0 min (9c-retinol), and 28.9 min (at-retinol). Other retinoids eluted at 2.0 min (retinyl palmitate and other RE) and 3.6 min (retinyl acetate) (see Fig. 1.8, Notes 37 and 46). 8. Quantify retinol isomers from calibration curves generated from standard amounts of each isomer separated using the retinol isomer method. 3.16. Polar Metabolites

3.16.1. Polar Metabolites with 4.6 mm ID column

Polar metabolites can be separated using the same mobile phase solvents as in Sections 3.14 and 3.15 using a gradient of water, acetonitrile, and formic acid. Blumberg et al., White et al., and Taimi et al. provide other separation methods (96–98). The separation method described by Taimi et al. using a 2.1 mm ID column and a flow rate of 0.2 ml/min is compatible with MS/MS detection (98). 1. Use a high performance liquid chromatograph (HPLC) consisting of a vacuum degasser, binary pump, temperaturecontrolled column compartment, and a temperaturecontrolled autosampler. 2. Maintain the column compartment at 25◦ C and the autosampler at 10◦ C. 3. Inject 100 μl. 4. Use a Zorbax SB-C18, 4.6 × 100 mm, 3.5 μm column (Agilent). 5. Separate analytes at 1 ml/min with a linear gradient from 75% H2 O/25% acetonitrile/0.1% formic acid to 1% H2 O/99% acetonitrile/0.1% formic acid over 30 min; hold at 1% H2 O/99% acetonitrile/0.1% formic acid for 7 min; return to initial conditions over 4 min; and hold for four additional minutes to equilibrate. 6. Use UV detection at 355 nm or MS/MS detection (see Section 3.18). 7. RA elutes at 30 min and polar metabolites elute between 18 and 23 min (4-OH-RA, 19.5 min; 4-oxo-RA, 20.1 min) (see Note 37). 8. Calibration curves are generated from standard amounts of each retinoid with the gradient solvent system used.

3.16.2. Polar Metabolites with 2.1 mm ID Column (from Taimi et al. (98))

1. Use a high performance liquid chromatograph (HPLC) consisting of a vacuum degasser, binary pump, temperaturecontrolled column compartment, and a temperaturecontrolled autosampler.

Quantification of Endogenous Retinoids

29

2. Use a Zorbax C18 Eclipse XDB 150 × 2.1, 5 μm column (Agilent). 3. Use water (solvent A), acetonitrile (solvent B), and 10% acetic acid (solvent C). 4. Separate polar metabolites at a flow rate of 0.2 ml/min starting with mixture of solvents A:B:C in the ratio 64:35:1 for 2 min, a linear gradient for 28 min up to 95% of solvent B with a constant flow rate of 1% solvent C, an isocratic hold at 95% B for 10 min, a linear gradient to initial conditions over 5 min, and an equilibration at initial conditions for 5 min. 5. Use MS/MS detection (see Section 3.18) or UV detection at 355 nm. 6. RA elutes at 33.2 min and polar metabolites elute between 15 and 20 min (4-OH-RA, 15.8 min; 4-oxo-RA, 16.9 min; 18-OH-RA, 19.0 min) (see Note 37). 3.17. Separation Optimization Strategies

Proper and sufficient chromatographic separation of analytes is essential to accurate quantification. Species of interest should be baseline resolved (peaks not overlapping) and have good peak shape (sharp peaks) (see Figs. 1.6, 1.7, and 1.8). Use standard solutions of the retinoids of interest to evaluate the separation before unknown analysis. If insufficient resolution is observed, several factors can be optimized to effect an adequate separation: 1. Column stationary phase. Surface chemistry differences lead to different selectivity for analyte separation. Choose column based on the selectivity of the stationary phase for the analytes to be separated. For example, C-16 alkylamide column stationary phases have greater resolving power for reverse-phase separations of RA isomers compared to C-18 stationary phases. C-18 columns work well for reverse-phase retinol, retinal, and retinyl ester separations. Silica columns are effective for normal-phase retinoid separations. 2. Column dimensions. Optimize column diameter and length to effect a separation. For analytical separations, columns with ID of 4.6 mm or smaller are typically used. MS-based detection commonly uses columns with ID of 2.1 mm or smaller. Smaller inner diameter columns not only increase resolution but also increase pressure. Capillary columns require a high pressure pump. Increasing column length increases resolution, but doubling column length results in double the elution time and solvent consumption with a 1.4fold increase in resolution (100). 3. Stationary phase particle size. Smaller diameter stationary phase particles not only increase resolution but also increase pressure. However, greater resolution resulting from smaller stationary phase particles will reduce the column length needed for a given separation.

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4. Mobile phase composition. Mobile phase composition must be compatible with column stationary phase and with the detection method. Mobile phase composition can affect MS ionization efficiency. Mobile phase choice can also influence peak shape, for example, acetonitrile-based retinol separations give sharper peak shapes as compared to methanolbased separations for retinol on a C-18 column (see Fig. 1.9). 5. Mobile phase gradient. Altering the gradient of a separation either in slope or in composition can assist in adjusting the retention time of analytes. 6. pH. The pH of some mobile phases is important and aids in the separation of charged species through ion pairing (see Note 36). 6. Flow rate. Optimum flow rate will give optimum resolution through maximizing plate height (see Note 47). 7. Temperature. Controlling temperature greatly assists in reproducibility of retention times. Elevating or reducing temperature can also be useful in optimizing a separation. 8. Separation mode/type. Switching from reverse phase to normal phase (or vice versa) may be necessary to effect sufficient resolution between species of interest. This modification requires switching column type (stationary phase). Low abundance of endogenous RA requires sensitive detection. MS/MS is currently the most sensitive method of RA detection and is readily coupled to LC separations capable of resolving RA isomers that have varied biological activity in vivo. Typically, quantitative MS/MS uses a triple-quadrupole MS instrument where the parent ion mass is selected in Q1, the parent

3.18. LC/MS/MS Detection

0.010

A

RE

0.008

Absorbance (325 nm)

Absorbance (325 nm)

Methanol

0.006 0.004

Retinol IS

0.002

0.06 0.04 0.02

RE

Acetonitrile

B

IS

0.010

Retinol 0.005

0.000

0.000 0

5

10 15 Retention Time (min)

20

25

0

5

10 15 Retention Time (min)

20

25

Fig. 1.9. Comparison of methanol-based and acetonitrile-based mobile phases for total retinol and RE separation. (a) Methanol based and (b) acetonitrile based. Panels a and b show identical mouse kidney samples separated with the same gradient of methanol or acetonitrile/water/1,2-dichloroethane. Reprinted from Ref. (51). Copyright © 2008, with permission from Elsevier; see Section 3.15.

Quantification of Endogenous Retinoids

31

ion is collisionally fragmented by N2 in Q2, and a product ion mass is selected in Q3 for detection. MS/MS offers appropriate sensitivity for RA detection through background reductions of 100–1000-fold over MS (see Fig. 1.10). MS/MS also imparts specificity by requiring analytes to meet both parent ion and product ion m/z conditions for detection. The sensitivity and background reduction advantage of MS/MS allows for analysis of smaller tissue samples and produces superior chromatograms for quantification (see Fig. 1.10). Information obtained from MS/MS fragmentation is also useful in the identification of unknown molecules. Several reports have shown that positive atmospheric pressure chemical ionization (APCI) has numerous advantages for RA analysis (as well as other retinoids), including favorable ionization efficiency based on the conjugated structure and carboxylic acid group (see Fig. 1.6), greater sensitivity and lower

6

4

2

0 200

225 250 m/z

275

301.1

6 4 2

283.1

201.1

1

177.4

205.0 255.0

0

300

175

200

225 250 m/z

275

300

2500

1.5

C

IS

1.0

D

λUV = 350 nm

m /z = 301.1/205.0

atRA

2000 Intensity

Absorbance × 10–3 (350 nm)

175

MS/MS

B

301.1 Intensity × 10–4

Intensity × 10–5

8

MS

A

atRA

0.5

1500 1000 500 0.04 g Liver

2.0 g Liver

0.0 0

1

2

3

4

5

Retention Time (min)

6

7

0 0

5

10

15

20

Retention Time (min)

Fig. 1.10. Background reduction by MS/MS detection. (a, b) RA mass spectra showing (a) Q1 scan with [M + H]+ (m/z: 301.1); (b) Q3 scan with [M + H]+ and product ions obtained after fragmentation. Both scans were obtained by infusing 200 nM RA at 10 μl/min. Note the reduction in background in (b) compared to (a). (c, d) Comparison of UV detection and MS/MS detection showing (c) UV detection at 350 nm (using a separation similar to alternate normal-phase separation) and (d) MS/MS detection using m/z 301.1 205.0 transition (and separation similar to tissue separation (gradient 2)). Note the reduction in background in (d) compared to (c). Also note the 50-fold lower tissue requirement for MS/MS detection in (d). Reprinted in part with permission from Ref. (50). Copyright © 2008, American Chemical Society; see Section 3.13.

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background than negative APCI, and greater signal intensity and linear dynamic range than electrospray ionization (49, 60, 77). APCI is also less susceptible than ESI to matrix suppression that can interfere with accurate quantification. Negative ESI–MS/MS, however, has been used effectively to identify polar metabolites produced from RA catabolism (96–98). 3.18.1. MS/MS Detection of RA Isomers

1. Use the separation described in Section 3.13.1 or 3.13.2. 2. Use an Applied Biosystems API-4000 triple-quadrupole mass spectrometer (or comparable instrument) equipped with APCI operated in positive ion mode. 3. Operate in multiple reaction monitoring (MRM) mode: monitor RA using an m/z 301.1 [M + H]+ to m/z 205.0 transition; monitor 4,4-dimethyl-RA using an m/z 329.4 [M + H]+ to m/z 151.3 transition, and use a dwell time of 150 ms for both RA and 4,4-dimethyl-RA (see Note 48). 4. The optimum positive APCI conditions on an API-4000 (Applied Biosystems) included the following: collision gas, 7; curtain gas, 10; gas1, 70; nebulizer current, 3; source temperature, 350; declustering potential, 55; entrance potential,10; collision energy, 17; collision exit potential, 5. Source position was vertical 790, horizontal 750 (see Note 49) (50). 5. Data from previous work were acquired with an Applied Biosystems API-3000 triple-quadrupole mass spectrometer equipped with APCI using the conditions described (49). 6. Assess detector capabilities and assay performance prior to quantification; then quantify each analyte from a calibration curve generated from standard amounts of that compound (see Section 3.22). 7. Peak identity/instrument performance for each method should be verified daily by injecting a mixture of authentic retinoid standards (see Note 37).

3.18.2. MS/MS Identification of RA Polar Metabolites (Taimi et al. (98))

1. Use with separation of polar metabolites described in Section 3.16. 2. Use a Micromass Quattro Ultima triple-stage quadruple mass spectrometer (Manchester, UK) in negative ESI mode. 3. Operate in either full mass scan mode (m/z 200–500) or product ion scan mode (m/z 50–400). Metabolites were characterized using MS/MS in the product ion scan mode (see Note 50). 4. Negative ESI conditions were capillary voltage, −3.15 KV; cone voltage, −37 V; desolvation gas (nitrogen) flow, 871 l/h; collision gas pressure, 2.3 × 10−3 torr; collision energy, 25 V.

Quantification of Endogenous Retinoids

3.18.3. Optimization Strategies for MS/MS Detection

33

Optimization of MS/MS conditions for maximum signal are essential to effective quantification efforts. Infusing a standard solution (∼1 nM–1 μM) via a syringe pump (∼1–100 μl/min) is necessary to tune the instrument properly before analysis (see Note 51). Optimize conditions by infusion and confirm with the chromatographic conditions used during quantification. 1. Instrument type. The model, vendor, and instrument type all affect the potential sensitivity. Different instruments will have different “base” sensitivities. 2. Molecular ion. Infuse each analyte to confirm the molecular ion obtained (see Note 52). 3. Ionization method. Whereas positive APCI has been reported to be most sensitive for retinoids, different ionization modes should be investigated for their sensitivity on a particular instrument. 4. Source position. Optimize the physical position of the source in relation to the orifice. This is essential when analytes are of low abundance. 5. Ionization conditions. Optimize all ionization conditions including gas flows, various voltages, and collision/fragmentation conditions. 6. Solvent composition. Optimize solvent composition (including mobile phase solvents and modifiers) which can affect sensitivity. 7. Product ion. Examine product ions produced by collisionally activated precursor ion fragmentations with N2 for intensity and background levels.

3.19. HPLC/UV

UV detection after HPLC separation of retinoids offers analysis specificity because very few compounds absorb at wavelengths characteristic of retinoids. The intrinsic absorption of most compounds in the sample milieu is significantly more blue (maxima at shorter wavelength) than that for retinoids. Additionally, UV detection of retinoids has specificity through structure-dependent absorbance maxima (see Table 1.2). Benefits of UV detection also include simplicity and cost-effectiveness (compared to MS-based detection methods). Whereas single wavelength and diode array detection (DAD) are effective for in vitro assay retinoid quantification and quantitation of abundant endogenous retinoids (retinal, retinol, RE) in vivo, they lack the necessary sensitivity for endogenous RA detection. Endogenous RA concentrations in the assay tissue amounts described here (see Table 1.3) are up to several orders of magnitude below the limit of detection and/or limit of quantification for both DAD and single-wavelength detection (50).

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3.19.1. Typical UV Detection Conditions

1. Use separations described or other (see Sections 3.13, 3.14, 3.15, 3.16, and 3.17). 2. Acquire absorbance spectra for analytes of interest in the mobile phase solvent to determine appropriate detection wavelength (see Table 1.2). 3. Use either a single-wavelength UV detector set at/near the absorbance maxima or use a DAD to collect an appropriate wavelength range to cover the entire absorbance spectra (see Note 53). 4. Assess detector capabilities and assay performance prior to quantification; then quantify each analyte from a calibration curve generated from standard amounts of that compound (see Section 3.22). 5. Peak identity/instrument performance for each method should be verified daily by injecting a mixture of authentic retinoid standards (see Note 37).

3.19.2. Optimization Strategies for UV Detection

1. To maximize signal. Choose a wavelength close to the absorbance maximum in the mobile phase solvent to maximize signal. 2. To maximize specificity. Choose a wavelength (not necessarily the maximum) that does not overlap or minimally overlaps with other retinoid species. 3. DAD vs. single wavelength. Single wavelength is often more sensitive than DAD; however, because DAD acquires the entire absorbance spectra, compound identity can be confirmed by its spectral signature.

3.20. GC/MS

For GC/MS detection, consult work by Napoli (45).

3.21. ECD

For detection by electrochemical detection consult work by Hagen et al., Sakhi et al., and Ulven et al. (70–72).

3.22. Essential Assay Characterization and Application

Several experiments must be performed to verify assay performance in order to obtain reliable, reproducible retinoid quantification data. Assay characterization includes determination of limits of detection (LOD), limits of quantification (LOQ), linear range, reproducibility, accuracy and precision, recovery, and handling-induced degradation.

3.22.1. LOD and LOQ

The LOD is defined by a signal/noise ratio of 3:1 and the LOQ is defined by a signal/noise ratio of 10:1. LOD and LOQ provide sensitivity measures. Determine for each analyte, with each method, and on each instrument.

Quantification of Endogenous Retinoids

35

1. Prepare a series of standard solutions on the day of use from a stock solution with a spectrophotometrically verified concentration (see Section 3.2). 2. Collect replicate data (at least triplicate) for each concentration. 3. Assess concentration of analyte that results in S/N=3 for LOD and S/N=10 for LOQ (see Fig. 1.11). 3.22.2. Linear Range

The concentration range of the standard solutions should span several orders of magnitude and encompass the amount of analyte that will be encountered in a physiological sample. 1. Prepare a series of standard solutions on the day of use with spectrophotometrically verified concentrations (see Section 3.2). 2. Collect replicate data (at least triplicate) for each concentration. 3. Plot average peak area as a function of concentration (see Fig. 1.11). 4. Exclude data that deviate from linearity at the high and low ends of the concentration range, if necessary. The remaining linear data define the working linear range. 5. Use linear regression to obtain the best fit line to the data and assess goodness of fit according to r2 (see Note 54).

100

4.0 × 104

A

1.0 × 103 Peak Area

B 3.5 × 104

75

3.0 × 104

25

LOQ

0

25

LOD

Peak Area

Intensity

50

7.5 × 102 5.0 × 102 2.5 × 102 0

2.5 × 104

0

2.0 × 104

10 20 30 40 50 fmol atRA

1.5 × 104 1.0 × 104 5.0 × 103

0 11.5 13.0 14.5 Retention Time (min)

0 0

200

400

600

800

1000

fmol atRA

Fig. 1.11. Assay characterization. (a) LOD and LOQ for atRA using gradient 2, tissue protocol (see Section 3.22.1). LOD is 0.75 fmol and LOQ is 0.125 fmol. (b–d) Representative calibration curve for atRA obtained using the cultured cell protocol (r2 , 0.999) (see Section 3.22.2). (b) Full linear range. (b, inset) Zoom in on data less than 50 fmol to show the functionality and linearity of the assay with low fmol. Reprinted in part with permission from Ref. (50). Copyright © 2008, American Chemical Society see Sections 3.13 and 3.22.

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6. Use the slope of the line to determine unknown concentrations of analyte according to peak area. 3.22.3. Accuracy/Precision

Accuracy is the agreement between applied and measured amounts and should be assessed at different areas of the linear range (low, mid, and high concentration). Precision is measured by the instrumental coefficient of variance (CV) which is obtained by repeat measurements of standard samples on the same day (see Note 55). 1. Prepare a series of standard solutions on the day of use with spectrophotometrically verified concentrations similar in concentration to that which will be encountered in a physiological sample (see Section 3.2). 2. Collect replicate data (at least triplicate) for each concentration. 3. Assess agreement between applied (prepared concentration) and measured (via calibration curve) amounts for accuracy. 4. Assess instrumental CV from repeat injections of the same sample concentration (see Note 56).

3.22.4. Recovery/Handling During Preparation

3.22.5. Evaluation of Internal Standard Performance

Percentage recovery reflects extraction efficiency and handling losses. The ability of an internal standard to reflect the loss and extraction efficiency of an endogenous retinoid must be evaluated before use in a quantitative assay. An internal standard should have a structure similar to the analyte, similar chromatographic behavior, and similar extraction efficiency. For example, 4,4-dimethyl-RA has a structure similar to atRA (see Fig. 1.6), has similar chromatographic behavior (see Fig. 1.6), and has similar extraction efficiency. Internal standard should be used at a level similar to the level of analyte and evaluated for each matrix (see Note 57). 1. Obtain sample (tissue, serum, cells, etc.) in sufficient quantity to perform multiple measurements (see Section 3.4). 2. Evaluate 3–10 samples per matrix type; allocate several samples as for background/blank determination. 3. Add internal standard to homogenized samples (see Section 3.6). 4. Add a known amount of exogenous retinoid to the same homogenate samples (comparable to endogenous level of analyte). 5. Extract and resuspend samples (see Sections 3.8 and 3.9). 6. Prepare several “100%” standard samples with the same amount of internal standard added to homogenate in the appropriate resuspension volume. Also prepare “100%” samples for exogenous retinoid.

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37

7. Analyze samples with desired separation and detection method (see Sections 3.12–3.21). 8. Calculate % of internal standard recovered and % of exogenous retinoid recovered. Amount of endogenous “background” should be accounted for in % exogenous retinoid recovery. Interferences from the biological matrix should also be evaluated in the “blank” samples (see Note 58). 9. Compare the % recovery of internal standard and the % recovery of retinoid of interest. Values need to be similar for use as an internal standard in quantitation. 3.22.6. Quantitation with an Internal Standard

1. Evaluate internal standard for ability to reflect endogenous retinoid of interest before use (see Section 3.22.5). 2. Add internal standard(s) to each homogenized sample and vortex well (5–10 s) (see Section 3.6). 3. Multiple internal standards can be used to reflect multiple analytes. 4. Extract and resuspend samples (see Sections 3.8 and 3.9). 5. Prepare several “100%” standard samples with the same amount of internal standard added to homogenate in the appropriate resuspension volume. 6. Analyze samples with desired separation and detection method (see Sections 3.12–3.21). 7. Calculate % of internal standard recovered in each sample. 8. Use % recovery value to obtain absolute amount of retinoid in sample.

3.22.7. Evaluation of Extraction Efficiency Without an Internal Standard

Quantitative analysis using a very efficient extraction in the sample preparation can be effective without an internal standard. Evaluate extraction efficiency with physiological levels of added exogenous retinoid. 1. Obtain sample (tissue, serum, cells, etc.) in sufficient quantity to perform multiple measurements (see Section 3.4). 2. Evaluate 3–10 samples per matrix type; allocate several samples as for background/blank determination. 3. Add a known amount of exogenous retinoid to homogenized sample (comparable to endogenous level of analyte). 4. Extract and resuspend samples (see Sections 3.8 and 3.9). 5. Prepare several “100%” standard samples with the same amount of exogenous retinoid added to homogenate in the appropriate resuspension volume. 6. Analyze samples with desired separation and detection method (see Sections 3.12–3.21).

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7. Calculate % of exogenous retinoid recovered. Amount of endogenous “background” should be accounted for in % exogenous retinoid recovery. 8. % recovery of retinoid of interest should consistently be >90–95% for extractions without an internal standard (see Notes 59 and 60). 3.22.8. Quantitation Without an Internal Standard

1. Evaluate extraction efficiency for retinoids of interest prior to sample analysis (see Section 3.22.5). 2. Extract and resuspend samples (see Sections 3.8 and 3.9). 3. Analyze samples with desired separation and detection method (see Sections 3.12–3.21). 4. Assume extraction losses to be negligible to obtain absolute amount of retinoid in sample (see Notes 59 and 60).

3.22.9. Evaluation of Handling-Induced Degradation by an Internal Standard

An internal standard may also act as an indicator of handlinginduced degradation, such as isomerization of RA (see Note 61). 1. Obtain sample (tissue, serum, cells, etc.) in sufficient quantity to perform multiple measurements (see Section 3.4). 2. Evaluate 3–10 samples per group; include a control group. 3. Add internal standard(s) to each homogenized sample and vortex well (5–10 s) (see Section 3.6). 4. Expose to mild and severe degradation stress (e.g., light, acid). 5. Extract and resuspend samples (see Sections 3.8 and 3.9). 6. Prepare several “100%” standard samples with the same amount of exogenous retinoid added to homogenate in the appropriate resuspension volume. 7. Analyze samples with desired separation and detection method (see Sections 3.12–3.21). 8. Evaluate degradation of internal standard compared to degradation of retinoid (see Note 61, Figs. 1.2 and 1.5).

3.22.10. Reproducibility

Reproducibility of the assay is evaluated in terms of the intraassay (same day) and inter-assay (different day) CV (see Note 55). Samples should be prepared using all procedures of the assay to reflect assay variability, including sample preparation and chromatographic analysis.

3.22.11. Intra-assay CV

1. Obtain sample (tissue, serum, cells, etc.) in sufficient quantity to perform multiple measurements (see Section 3.4). 2. Prepare multiple samples on the same day. For example, 3–10 samples prepared separately from a single minced mouse or rat liver.

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39

3. Homogenize, extract, and resuspend samples individually (see Sections 3.5, 3.6, 3.7, 3.8, and 3.9). 4. Collect data according to desired method on the same day (see Sections 3.12–3.21). 5. Quantify the amount of endogenous retinoid (see Sections 3.22.2 and 3.22.4). 6. Calculate % CV (see Note 56). 3.22.12. Inter-assay CV

1. Obtain sample (tissue, serum, cells, etc.) in sufficient quantity to perform multiple measurements (see Section 3.4). 2. Prepare multiple samples on several different days. For example, 3–10 samples each day on 3–5 days over the course of a week. Ideally samples will be identical, for example, prepared separately from a single minced mouse or rat liver. 3. Samples should be stored at −80◦ C until assay (see Section 3.4). 4. Homogenize, extract, and resuspend samples individually (see Sections 3.5, 3.6, 3.7, 3.8, and 3.9). 5. Collect data according to desired method (see Sections 3.12–3.21). 6. Quantify the amount of endogenous retinoid (see Sections 3.22.2 and 3.22.4). 7. Calculate % CV (see Note 56).

3.23. Identity Confirmation Strategies

3.23.1. Mass Signature

The identity of an analyte should be confirmed by multiple methods. This is especially important when new compounds are being identified. Identity confirmation is also important for new and existing assays to confirm that there are not any additional interfering species co-eluting during chromatography or during detection that could interfere with accurate identification and quantification. Interferences can be due to components of the biological matrix remaining in the prepared sample or other endogenous retinoids that are not sufficiently chromatographically resolved. 1. Use MS to determine characteristic mass of analyte. 2. Preferably, for more specificity, use MS/MS to determine characteristic precursor to product ion mass transition. 3. If analyte is unknown species, MS/MS can assist in providing structural information

3.23.2. Co-elution with Authentic Standards

1. Compare the retention time of analytes in a sample with the retention time of authentic standards. 2. Data for pure standards and sample analytes should be collected under identical conditions (separation method, detection, etc.).

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It is important to confirm that species have not been artifactually produced or converted into another analyte during sample preparation (e.g., by hydrolysis, isomerization). Stability of compounds during preparation can be evaluated by spiking in physiologically relevant levels of exogenous retinoid and comparing to a nonspiked control (see Notes 62 and 63). 1. Spike-in before homogenization. Exogenous retinoid is spiked into homogenization solution (e.g., saline) (see Section 3.2) and then samples are homogenized, extracted, and resuspended similar to control. The desired result is an increase only in the peak of analyte postulated to be of the same identity as the spiked-in retinoid (and observed augmentation is consistent with the concentration of added exogenous retinoid) (see Note 64, Fig. 1.12).

3.23.3. Addition of Authentic Standards During Sample Preparation

2. Spike-in after homogenization but before extraction. Spikein exogenous retinoid (as in step 1) but after homogenization to tissue. none

none

+9cROL

atROL

0.003 0.002 0.001

0.4 0.3 0.2 0.1 9cROL

0.0003

9cROL

0.0002 0.0001

26 27 28 29 30 31 Retention Time (min)

26 27 28 29 30 31 Retention Time (min)

0.10

0.0000

atROL

0.08 0.06 0.04 0.02 9cROL

0.00 26 27 28 29 30 31 Retention Time (min)

0.01 9cROL

atRAL 0.12 atROL

Absorbance (325 nm)

0.0002

Absorbance (325 nm)

9cROL

0.02

26 27 28 29 30 31 Retention Time (min)

0.0010

atROL

0.0004

0.03

none

0.0010

0.0006

0.04

26 27 28 29 30 31 Retention Time (min)

D

+9cRAL

0.0008

atROL

0.05

0.00

none

C Absorbance (325 nm)

0.0004

0.0000

0.0

0.000

Absorbance (325 nm)

9cROL

atROL

Absorbance (325 nm)

Absorbance (325 nm)

Absorbance (325 nm)

atROL

0.004

0.06

0.0005

0.5

0.005

+atROL

0.0008 0.0006

9cROL

0.0004 0.0002 0.0000

26 27 28 29 30 31 Retention Time (min)

Absorbance (325 nm)

A

B

atROL

0.10 0.08 0.06 0.04 0.02 9cROL

0.00 26 27 28 29 30 31 Retention Time (min)

26 27 28 29 30 31 Retention Time (min)

Fig. 1.12. Identity verification by spike in of exogenous retinoids. (a–d) Addition of exogenous retinoids to mouse liver before homogenization to show that 9c-retinol is not formed artifactually during sample preparation. The left panel of each pair is a magnified view of the right panel to show 9c-retinol more clearly. (a) The addition of 9c-retinol increases 9c-retinol only. (b) The addition of at-retinol increases at-retinol only. (c, d) The addition of either 9c-retinal or at-retinal does not increase either 9c-retinol or at-retinol. Reprinted from Ref. (51). Copyright © 2008, with permission from Elsevier.

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41

3. Spike-in after extraction to resuspended sample. Spike-in exogenous retinoid (as in step 1) but after extracted sample is resuspended. Here, sample can be injected and analyzed followed by the addition of exogenous retinoid, re-injection, and analysis. 4. Evaluate all three spike-in scenarios (steps 1, 2, and 3). 3.23.4. Use of Chromatography of Varied Selectivity

Altering the selectivity of the chromatographic separation should be done to confirm that no additional species are co-eluting with the analyte of interest. 1. Reverse phase, normal phase. Switching from reverse phase to normal phase or vice versa will significantly change the separation selectivity and analyte retention characteristics. Similar results should be obtained with both separation types. 2. Stationary phase. Switching column stationary phase chemistry can be effective if the stationary phases have sufficiently different selectivity. 3. Mobile phase. Similar to step 2. 4. 2D chromatography. The analyte of interest is separated with one column and separated again on a chromatographic column of different selectivity. Can be performed in tandem or a fraction containing the analyte of interest can be collected from the first separation and then re-chromatographed on the alternate selectivity system.

3.24. Application 3.24.1. Tissue

The assays described here have been applied to various tissues. A summary of retinoid values is provided (see Table 1.6). Retinoid levels vary according to some or all of the following: strain, age, sex, diet, genotype, and/or exogenous treatment. Ideally each experimental comparison has a cohort of control animals and a cohort of experimental animals side by side.

3.24.2. Cell Systems

Retinoids and retinoid assays can be quantified from cell systems, including isolated cells, primary culture cell systems, and established cell lines. If endogenous retinoids are to be quantified, handling precautious must be observed during isolation and culture. For enzyme assays, precautions must be observed during assay (see Sections 3.1, 3.2,3.3, and 3.4). 1. Cells should be switched to serum-free media before assay. Media should always also be measured for presence of retinoids (see Note 65). 2. Cells and/or media can be quantified for retinoid content either endogenous or as a result of treatment.

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Table 1.6 Select retinoid levels in adult mousea Tissue

atRA (pmol/g)

Total retinal (pmol/g)

Total retinol (nmol/g)

Total RE (nmol/g)

Serumb

2.7 ± 0.3 (21)

32.2 ± 6.2 (6)

0.81± 0.04 (70)

0.22 ± 0.02 (69)

Liverb

38.1 ± 3.4 (18)

160.9 ± 14.3 (26)

9.6 ± 0.9 (60)

562.6 ± 75.9 (55)

Kidneyb

15.2 ± 2.2 (30)

187.3 ± 31.2 (12)

0.60 ± 0.04 (37)

1.8 ± 0.2 (37)

Adiposeb

14.2 ± 2.4 (18)

63.5± 5.2 (12)

0.63 ± 0.03 (37)

0.59 ± 0.09 (29)

Muscleb

1.5 ± 0.2 (15)

–

0.15 ± 0.02 (38)

0.25 ± 0.03 (31)

Spleenb

7.3 ± 0.6 (14)

–

0.60 ± 0.06 (26)

1.2 ± 0.1 (26)

Testisb

8.9 ± 1.0 (14)

90.7 ± 10.1 (12)

0.08 ± 0.01 (27)

0.31 ± 0.02 (27)

Brainb

17.1 ± 3.7 (19)

–

0.32 ± 0.01 (5)

0.22 ± 0.02 (5)

Brainc

33.9 ± 3.9 (8)

–

0.68 ± 0.23 (19)

0.84 ± 0.16 (19)

Hippocampusc

45.3 ± 5.2 (8)

–

0.30 ± 0.03 (27)

0.70 ± 0.05 (27)

Cortexc

16.0 ± 1.3 (7)

–

0.08 ± 0.01 (18)

0.35 ± 0.04 (18)

Olfactory bulbc

76.5 ± 21.3 (4)

–

0.20 ± 0.02 (4)

0.63 ± 0.03 (4)

Thalamusc

80.9 ± 6.0 (4)

–

0.20 ± 0.04 (4)

0.47 ± 0.06 (4)

Cerebellumc

54.8 ± 3.6 (8)

–

0.42 ± 0.08 (8)

0.73 ± 0.10 (8)

Striatumc

78.0 ± 33.2 (3)

–

0.21 ± 0.05 (4)

0.41 ± 0.08 (4)

(epididymal)

a Only a partial list of recovered in vivo retinoid levels. For a full description see Kane et al. (49–51) b Data were obtained from 2- to 4-month-old male SV129 mice fed and bred from dams fed an AIN93G diet with

4 IU vitamin A/g c Data were obtained from 4-month-old male C57BL/6 mice fed an AIN93M with 4 IU vitamin A/g from weaning and bred from dams fed a stock diet (>30 IU vitamin A/g). Values are means ± SEM (n). Serum values are either pmol/ml or nmol/ml. –, not measured

3. Typical amount of cells and/or media analyzed depends on situation (see Note 66). 4. If retinoid production is to be monitored, retinoid precursors should be added to cells/samples via a calibrated glass syringe via a concentrated solution (freshly prepared), so that addition volume does not exceed 1–2% of sample volume (usually ∼5–10 μl). 5. Retinol as substrate should be purified before use (see Section 3.3). 3.25. Bio-analysis Limitations and Potential Pitfalls

Whereas some assays offer benefits over others, no method is devoid of limitations. Listed are some general potential pitfalls when assaying biological samples. 1. LC/MS/MS. LC/MS/MS can have interfering background contributions from the biomatrix (see Note 67). Contribution of biomatrix interference to overall retinoid

Quantification of Endogenous Retinoids

43

signal should be assessed and chromatography and/or MS/MS conditions adjusted to eliminate or minimize background contributions (see Sections 3.17 and 3.18.3). 2. LC/MS. LC/MS has less specificity and higher background than MS/MS. 3. LC/UV. UV detection is more effective with abundant retinoids (RE, retinol, retinal) than RA. Both single wavelength and DAD are less sensitive than MS/MS for RA detection and lack the positive mass identification provided by mass spectrometry. Chromatography and UV detection can also be optimized to eliminate or minimize background contributions and co-elution (see Sections 3.17 and 3.18.3). 4. LC/ECD. Retinoid quantification with ECD is potentially susceptible to interference from other species in the biomatrix and also lacks positive mass identification. 5. GC/MS. GC/MS presents challenges for isomer separation and derivatization is often necessary. 6. Isomer separations. Insufficient chromatographic resolution of isomers can potentially skew quantification due to coelution. This is especially problematic for RA detection (see Section 1). 7. ESI. ESI (electrospray ionization) is susceptible to matrix suppression effects that hinder reproducibility needed for accurate quantification. Retinoid signal can be suppressed to varying degrees by components of the biomatrix during ionization. These matrix suppression effects can fluctuate according to matrix components (sample type, preparation method, etc.) and also across a chromatographic gradient.

4. Notes 1. If you do not have a room with overhead yellow lights, a desk lamp outfitted with a yellow light bulb can be used in a darkened room. “Bug-light” type yellow bulbs (which block lower wavelength light that attracts bugs) work well and can be purchased at any hardware store. Do NOT use yellow “party bulb”-type lights. 2. Exposure to full spectrum (white) light (regular room lights) should be avoided, even for brief periods of time. (Noticeable degradation takes place in ∼10 min!) 3. All tissue harvest and dissections for retinoid analysis should be performed under yellow light. Sample collection

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under white lights will result in isomerization and degradation of retinoids. 4. Dissections should be performed with a yellow filter on the light source like a Volpi (Auburn, NY, USA) NCL 150 light source with a red or yellow filter. 5. Retinoids stick to many plastics and variable loss of up to 40% occurs when pipetting retinoid solutions with regular (plastic) pipette tips. 6. To flush, draw up clean solvent and expel to waste. 7. Retinoid residue within calibrated syringes can contaminate samples without proper cleaning. It is not an exaggeration that 15–20 flushes are required. 8. An evaporator with the capability to use disposable nitrogen delivery elements (such as glass pipettes) is highly desirable. Evaporators with permanent nitrogen delivery elements will cross-contaminate samples. 9. An acid bath can be used to periodically remove tissue (and/or retinoid) residue. Soak overnight in ∼1 M HCl or HNO3 followed by neutralization and flushing with copious amounts of water to remove all residue. 10. Use of a disposable instrument, like the tip of a Pasteur pipette, can help prevent contamination by retinoid residue remaining on reusable tools. 11. Cuvettes should be cleaned thoroughly before and after use with ethanol and/or acetone and then dried completely. If acetone is used, complete removal is particularly important as acetone will absorb significantly at low wavelengths. For a more stringent cleaning rinse cuvettes with concentrated nitric acid followed by water followed by 100% ethanol and complete drying. 12. Solutions with extra peaks, maxima at the wrong wavelength, or a large peak at low wavelength (200–300 nm) are either contaminated and/or degraded and should not be used. 13. Molar absorptivity (ε) is wavelength (λ) and solvent dependent. Measure absorbance according to the wavelength and solvent for a given ε as described in Table 1.2. 14. Acetone will interfere with the absorbance spectrum (absorbing highly at low wavelength) and must be removed. 15. 5% MeOH results in a faster reaction than THF alone. 16. Different matrices/different tissue types have different susceptibility to degradation during storage.

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17. Homogenized samples frozen immediately and stored at −80◦ C for 1 day gave comparable results (within 10%) with those of freshly analyzed samples; however, homogenized samples stored for 1 month gave values that were 50% less than samples analyzed immediately. Therefore, it should be possible to store tissue homogenates overnight in the freezer, but long-term storage of homogenized samples should be avoided, even at −80◦ C (49). 18. Alternate motion/position of pestle/homogenizer to avoid generating too much friction in a localized area. When homogenizing try not to pull any suction when moving the pestle up and down. 19. Polytron homogenization should only be used if other homogenization methods are ineffective. 20. Homogenization of skin will have a non-homogenizable portion left over which should be subtracted from the tissue weight to give a net extracted tissue amount. 21. Degradation and isomerization by the biological matrix will occur. Samples should be extracted within 30–60 min after homogenization. Homogenized samples will undergo significant degradation after 2 h at 4◦ C from matrix effects (48, 49). 22. For more information on Bradford dye-binding assay for total protein determination visit www.bio-rad.com. 23. Known amounts of BSA protein (or similar) should be used to generate a standard curve. 24. Protein should be diluted so absorbance readings from protein–dye are in the linear range (of the standard curve generated by known protein amounts). 25. Absorbance between 0.1 and 1.0 should fall in the linear range; however, this is dependent on standard curve (see Note 23). 26. Dedicated syringes to each internal standard can prevent inadvertent contamination by retinoid residue in a syringe. Syringes should be cleaned before each use. 27. Take care not to touch the tip of the syringe to the sample to avoid cross-contamination. 28. Hexane extracts are evaporated under nitrogen gas immediately or kept on ice until evaporation (as soon as possible). 29. There should be a crisp layer between phases. If a center layer is observed there may be inefficient extraction of the retinal O-ethyl oxime derivatives. Adjust reagents to eliminate.

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30. Samples resuspended in acetonitrile were more stable than suspensions in other solvents, including mixtures of methanol/acetonitrile/water, methanol/acetonitrile, and methanol. 31. Samples should not be resuspended or stored in mobile phases that contain acid. 32. Resuspension volume may depend on vitamin A status. For example, liver resuspension volumes can range from 120 to 1000 μl. 33. For adipose tissue (and other high lipid content tissues), there is often a sizable amount of lipid present after evaporation in the retinol/RE extract. Resuspend in a slightly larger volume (200 vs. 120 μl), vortex mix, and then transfer the sample avoiding the lipid drop. Lipid collection in the bottom of the low volume insert is problematic for injection and analysis. An internal standard can account for sample loss to the lipid drop (see Note 56). 34. The acid–base extraction described here (see Section 3.8) is preferable to analyses that use saponification. Saponification, an alkaline digestion that frees retinoids from the stabilizing matrix and lipids while hydrolyzing RE to retinol to yield a total retinol measurement, can be problematic. The elevated temperature and exposure to alkali often causes retinoid degradation and isomerization of 4–40% (2). This loss is illustrated by 30–65% lower total ROL values obtained after saponification compared with the sum of ROL and RE values obtained separately (16, 22). 35. Small samples include those ∼10–20 mg or less of tissue. Small samples can also include samples of low retinoid abundance such as VAD diet, genetically manipulated animals, or exogenous compound-treated animals. 36. Some stationary phases are susceptible to pH-dependant degradation. Manufacturer’s recommendations should be followed. 37. Confirm retention times and peak identity with authentic standards frequently. 38. Prepare mobile phase fresh as small changes in (normal phase) mobile phase composition (due to evaporation, etc.) will result in significant changes in retention time(s). 39. Although levels of RA in vivo were not detectable above background and/or were the same magnitude as random/interfering peaks, this method is useful for applications in which RA is high such as enzyme assays. Quantification of in vivo levels of RA is best accomplished with more sensitive detection methods (49, 50).

Quantification of Endogenous Retinoids

47

40. Note that the column needs to be flushed periodically when quantifying ester-rich tissue (e.g., liver) to reduce RE accumulation. A flush with the total retinol and RE separation described in Section 3.15.1 followed by equilibration with a blank total retinal separation as described in Section 3.14.1 is effective. 41. The amount of RE present can be significantly higher than retinol and require re-injection of a smaller volume to give an RE peak area within the linear range. 42. For Sections 3.14.1 and 3.15.1 reservoirs can be set up as follows: A: H2 O, B: H2 O with 10% formic acid, C: acetonitrile, D: 1,2-dichloroethane. For example, to achieve 40% H2 O/60% acetonitrile/0.1% formic acid HPLC flow can be set to 39% H2 O, 1% H2 O with 1% formic acid, and 60% acetonitrile. 43. The total ROL/RE reverse-phase method was modified from previous methods to use an acetonitrile/water/formic acid mobile phase that transitions to an acetonitrile/dichloroethane mobile phase (31, 99). The acetonitrile/water/formic acid mobile phase gave sharper ROL peaks than did the previous methanol/water-based ROL separations 44. Not all endogenous REs were resolved using this method; it was intended to quantify only total RE. Figure 1.3a shows retinyl palmitate only, which is up to 90% of the endogenous ester. Retinyl oleate co-eluted with retinyl palmitate, whereas other esters, such as retinyl linoleate, retinyl myristate, and retinyl stearate, eluted just before or after retinyl palmitate. 45. The sum of all ester peaks was used to calculate total RE. Retinyl palmitate was used as the calibrant to calculate total ester because retinyl palmitate and other RE have similar absorbance maxima (44). 46. Retinyl palmitate can be quantified using the retinol isomer method, but because of its minimal retention and the possible background contribution from minimally retained matrix components, the total ROL/RE method described here is preferable for RE quantification. 47. Optimum plate height and flow rate occur when contributions from eddy diffusion, longitudinal diffusion, and resistance to mass transfer are minimized (100). 48. Optimize parent ion/fragment ion transition for your conditions to maximize background and minimize background and interferences from biomatrix species.

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49. Source conditions listed are guidelines. Optimize conditions for your instrument/conditions. Tuning of instrumental conditions is essential to obtaining sufficient sensitivity to collect in vivo sample data. 50. ESI is more susceptible to ion suppression effects than APCI and is less desirable for quantification. 51. Always start with a more dilute solution (rather than a concentrated solution) when beginning tuning. 52. Some retinoids lose part of the parent molecule during ionization resulting in a molecular ion m/z of (parent-lost group). For example, retinol [MH – H2 O]+ and retinyl acetate [MH – AcOH]+ have been reported (60). 53. If multiple analytes are being detected using a singlewavelength detector, use a detection wavelength appropriate for all analytes. 54. r2 of 1 is a perfect correlation between peak area and concentration. r2 of 0.99 or greater is desirable. 55. The coefficient of variation (CV), also known as “relative variability,” equals the standard deviation divided by the mean. It can be expressed either as a fraction or a percent. 56. % CV of 5–10% or less is desirable. 57. Internal standard performance can vary by analyte and biological matrix. For example, retinyl acetate reflected RE recovery accurately for all tissues investigated using the acid–base extraction in Section 3.8. For adipose and other lipid-rich tissue tissues, retinyl acetate also accurately reflected the recovery of retinol. However, retinyl acetate did not always accurately reflect retinol recovery from liver and, thus, was not used to adjust liver retinol values (51). Retinyl acetate used as an internal standard represents the recovery of RE and retinol in adipose with reasonable accuracy. 58. If significant interference is observed with either the analyte of interest or the internal standard, analysis conditions must be altered. 59. For extraction described in Section 3.3.8.1, exogenous retinol spiked into liver homogenate before extraction was recovered 94 ± 4% (n = 3) and extraction losses were assumed to be negligible (No IS was used to adjust this value.) (51). 60. For extraction described in Section 3.3.8.4, recovery of the retinal O-ethyloximes routinely exceeded 95% and

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extraction losses were assumed to be negligible (No IS was used.) (51). 61. An example of an internal standard indicating analyte degradation is illustrated by 4,4-dimethyl-RA (see Figs. 1.2 and 1.5). In these panels the internal standard isomerization closely mirrors that of endogenous atRA under both mild and severe conditions. The ability of an RA internal standard to indicate handling-induced isomerization is a valuable characteristic, because it helps distinguish endogenous RA isomers from those formed during handling. Samples that have been handled properly have no isomers in the 4,4-dimethyl-RA chromatogram, indicating that observed isomers in the RA chromatogram are endogenous (see Fig. 1.5a). Samples with artifactual isomers also show isomers in the 4,4-dimethyl-RA chromatogram (see Fig. 1.5b). Note the difference in the decrease in atRA and increase in cis-RA isomers is concurrent with isomers occurring in the internal standard chromatogram. Samples with significant isomerization of the internal standard (>10–15%) should be discarded. Previous reports have concluded that the biological matrix can cause ∼7% isomerization of atRA into cis-isomers (48, 49). cisIsomers exceeding this proportion should be endogenous, provided that the internal standard shows no isomerization. 62. It important to not overwhelm endogenous levels or saturate analysis conditions to the point that augmentation of the retinoid of interest or changes to other retinoids cannot be observed. 63. Stability of analyte(s) during preparation should be tested for each analyte and each matrix type as various tissues have different potential for analyte hydrolysis, isomerization, etc. 64. Control and spike-in samples should be sufficiently similar to distinguish that changes in peak area are from exogenously added retinoid and not sample variation. 65. All serum-containing media contains retinol (and sometimes RE and RA) and many media formulations have supplements that include retinoids. 66. 0.5–1 ml media and/or a confluent amount of cells in a 6or 12-well plate has been successfully used in a variety of experiments in the Napoli Lab. 67. Some reports show significant background contributions that could be problematic during detection of in vivo levels of RA in a biomatrix (79).

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References 1. Wolf, G. (1984) Multiple functions of vitamin A. Physiol. Rev. 64, 873–937. 2. Sporn, M.B., Roberts, A.B., Goodman, D.S. (eds.). (1994) The Retinoids: Biology, Chemistry, and Medicine, 2nd ed., Raven, New York. 3. Wolf, G. (2001) Retinoic acid homeostasis: Retinoic acid regulates liver retinol esterification as well as its own catabolic oxidation in liver. Nutr. Rev. 59, 391–394. 4. Harrison, E.H. (2005) Mechanisms of digestion and absorption of dietary vitamin A. Annu. Rev. Nutr. 25, 87–103. 5. Blomhoff, R., Blomhoff, H.K. (2006) Overview of retinoid metabolism and function. J. Neurobiol. 66, 606–630. 6. Napoli, J.L. (1999) Interactions of retinoid binding proteins and enzymes in retinoid metabolism. Biochim. Biophys. Acta 1440, 139–162. 7. Napoli, J.L. (1999) Retinoic acid: its biosynthesis and metabolism. Prog. Nucleic Acid Res. Mol. Biol. 63, 139–188. 8. Chambon, P. (1996) A decade of molecular biology of retinoic acid receptors. FASEB J. 10, 940–954. 9. Shaw, N., Elholm, M., Noy, N. (2003) Retinoic acid is a high affinity selective ligand for the peroxisome proliferator-activated receptor beta/delta. J. Biol. Chem. 278, 41589–41592. 10. Schug, T.T., Berry, D.C., Shaw, N.S., Travis, S.N., Noy, N. (2007) Opposing effects of retinoic acid on cell growth result from alternate activation of two different nuclear receptors. Cell 129, 723–733. 11. Balmer, J.E., Blomhoff, R. (2002) Gene expression regulation by retinoic acid. J. Lipid Res. 43, 1773–1808. 12. Ruberte, E., Friederich, V., Chambon, P., Moriss-Kay, G. (1993) Retinoic acid receptors and cellular retinoid binding proteins. III. Their differential transcript distribution during mouse nervous system development. Development 118, 267–282. 13. Ong, D.E. (1994) Cellular transport and metabolism of vitamin A: Roles of the cellular retinoid-binding proteins. Nutr. Rev. 52, S24–S31. 14. Yamagata, T., Momoi, M.Y., Yanagisawa, M., Kumagai, H., Yamakado, M., Momoi, T. (1994) Changes of the expression and distribution of retinoic acid receptors during neurogenesis in mouse embryos. Brain Res. Dev. Brain Res. 77, 163–176. 15. Krezel, W., Kastner, P., Chambon, P. (1999) Differential expression of retinoid receptors

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

in the adult mouse central nervous system. Neuroscience 89, 1291–1300. Quadro, L., Blaner, W.S., Salchow, D.J., Vogel, S., Piantedosi, R., Gouras, P., Freeman, S., Cosma, M.P., Colantuoni, V., Gottesman, M.E. (1999) Impaired retinal function and vitamin A availability in mice lacking retinol-binding protein. EMBO J. 18, 4633–4644. Ghyselinck, N.B., Bavik, C., Sapin, V., Mark, M., Bonnier, D., Hindelang, C., Dierich, A., Nilsson, C.B., Hakansson, H., Sauvant, P., Azais-Braesco, V., Frasson, M., Picaud, S., Chambon, P. (1999) Cellular retinol-binding protein I is essential for vitamin A homeostasis. EMBO J. 18, 4903–4914. Zhang, X.E.L., Lu, J., Tso, P., Blaner, W.S., Levin, M.S., Li, E. (2002) Increased neonatal mortality in mice lacking cellular retinolbinding protein II. J. Biol. Chem. 277, 36617–36623. Vogel, S., Mendelsohn, C.L., Mertz, J.R., Piantedosi, R., Waldburger, C., Gottesman, M.E., Blaner, W.S. (2001) Characterization of a new member of the fatty acid binding protein family that binds all-trans-retinol. J. Biol. Chem. 276, 1353–1360. Gorry, P., Lufkin, T., Dierich, A., RochetteEgly, C., Decimo, D., Dolle, P., Mark, M., Durand, B., Chambon, P. (1994) The cellular retinoic acid binding protein I is dispensable. Proc. Natl. Acad. Sci. USA 91, 9032–9036. Lampron, C., Rochette-Egly, C., Gorry, P., Dolle, P., Mark, M., Lufkin, T., LeMeur, M., Chambon, P. (1995) Mice deficient in cellular retinoic acid binding protein II (CRABPII) or in both CRABPI and CRABPII are essentially normal. Development 121, 539–548. O’Byrne, S.M., Wongsiriroj, N., Libien, J., Vogel, S., Goldberg, I.J., Baerh, W., Palczewski, K., Blaner, W.S. (2005) Retinoid absorption and storage is impaired in mice lacking lecithin:retinol acyltransferase (LRAT). J. Biol. Chem. 280, 35647–35657. Liu, L., Gudas, L.J. (2005) Disruption of the lecithin:retinol acyltransferase gene makes mice more susceptible to vitamin A deficiency. J. Biol. Chem. 280, 40226–40234. Zhang, M., Hu, P., Krois, C.R., Kane, M.A., Napoli, J.L. (2007) Altered vitamin A homeostasis and increased size and adiposity in the rdh1-null mouse. FASEB J. 21, 2886–2896. Hu, P., Zhang, M., Napoli, J.L. (2007) Ontogeny of rdh9 (Crad3) expression: Ablation causes changes in retinoid and steroid

Quantification of Endogenous Retinoids

26.

27.

28.

29.

30.

31.

32.

33.

34.

metabolizing enzymes, but RXR and androgen signaling seem normal. Biochim. Biophys. Acta 1770, 694–705. Germain, P., Chambon, P., Eichele, G., Evans, R.M., Lazar, M.A., Leid, M., De Lera, A.R., Lotan, R., Mangelsdorf, D.J., Gronemeyer, H. (2006) International union of pharmacology: LXIII. Retinoid X receptors. Pharmacol. Rev. 58, 760–772. Kawaguchi, R., Yu, J., Honda, J., Hu, J., Whitelegge, J., Ping, P., Wiita, P., Bok, D., Sun, H. (2007) A membrane receptor for retinol binding protein mediates cellular uptake of vitamin A. Science 315, 820–825. Misner, D.L., Jacobs, S., Shimizu, Y., de Urquiza, A.M., Solomin, L., Perlmann, T., DeLuca, L.M., Stevens, C.F., Evans, R.M. (2001) Vitamin A deprivation results in reversible loss of hippocampal long-term synaptic plasticity. Proc. Natl. Acad. Sci. USA 98, 11714–11719. Deltour, L., Foglio, M.H., Duester, G. (1999) Metabolic deficiencies in alcohol dehydrogenase Adh1, Adh3, and Adh4 null mutant mice. J. Biol. Chem. 274, 16796–16801. Molotkov, A., Duester, G. (2002) Retinol/Ethanol drug interaction during acute alcohol intoxication in mice involves inhibition of retinol metabolism to retinoic acid by alcohol dehydrogenases. J. Biol. Chem. 277, 22553–22557. Lei, Z., Chen, W., Zhang, M., Napoli, J.L. (2003) Reduction of all- trans-retinal in the mouse liver peroxisome fraction by the short-chain dehydrogenase/reductase RRD: Induction by the PPARα ligand clofibrate. Biochemistry 42, 4190–4196. Fiorella, P.D., Olson, J.R., Napoli, J.L. (1995) 2,3,7,8-Tetrachlorodibenzo-p-dioxin induces diverse retinoic acid metabolites in multiple tissues of the Sprague-Dawley rat. Toxicol. Appl. Pharmacol. 134, 222–228. Kransler, K.M., Tonucci, D.A., McGarrigle, B.P., Napoli, J.L., Olson, J.R. (2007) Gestational exposure to 2,3,7,8tetrachlorodibenzo-p-dioxin alters retinoid homeostasis in maternal and perinatal tissues of the Holtzman rat. Toxicol. Appl. Pharmacol. 224, 29–38. Hoegberg, P., Schmidt, C.K., Fletcher, N., Nilsson, C.B., Trossvik, C., Schuur, A.G., Brouwer, A., Nau, H., Ghyselinck, N.B., Chambon, P., Hakansson, H. (2005) Retinoid status and responsiveness to 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) in mice lacking retinoid binding protein or retinoid receptor forms. Chem. Biol. Interact. 156, 25–39.

51

35. Staels, B. (2001) Regulation of lipid and lipoprotein metabolism by retinoids. J. Am. Acad. Dermatol. 45, S158–S167. 36. Yang, Q., Graham, T.E., Mody, N., Preitner, F., Peroni, Q.D., Zabolotny, J.M., Kotani, K., Quadro, L., Kahn, B.B. (2005) Serum retinol binding protein 4 contributes to insulin resistance in obesity and type 2 diabetes. Nature 436, 356–362. 37. Wolf, G. (2007) Serum retinol-binding protein: A link between obesity, insulin resistance, and type 2 diabetes. Nutr. Rev. 65, 251–256. 38. Ziouzenkova, O., Orasanu, G., Sharlach, M., Akiyama, T.E., Berger, J.P., Viereck, J., Hamilton, J.A., Tang, G., Dolnikowski, G.G., Volgel, S., Duester, G., Plutzky, J. (2007) Retinaldehyde represses adipogenesis and diet-induced obesity. Nat. Med. 13, 695–702. 39. Farias, E.F., Ong, D.E., Ghyselinck, N.B., Nakajo, S., Kuppumbatti, Y.S., Mira y Lopez, R. (2005) Cellular retinol-binding protein I, a regulator of breast epithelial retinoic acid receptor activity, cell differentiation, and tumorigenicity. J. Natl. Cancer Inst. 97, 21–29. 40. Lotan, R. (2005) A crucial role for cellular retinol-binding protein I in retinoid signaling. J. Natl. Cancer Inst. 97, 3–5. 41. Fields, A.L., Soprano, D.R., Soprano, K.J. (2007) Retinoids in biological control and cancer. J. Cell. Biochem. 102, 886–898. 42. Szeto, W., Jiang, W., Tice, D.A., Rubinfeld, B., Hollingshead, P.G., Fong, S.E., Dugger, D.L., Pham, T., Yansura, D.G., Wong, T.A., Grimaldi, J.C., Corpuz, R.T., Singh, J.S., Frantz, G.D., Devaux, B., Crowley, C.W., Schwall, R.H., Eberhard, D.A., Rastelli, L., Polakis, P., Pennica, D. (2001) Overexpression of the retinoic acid-responsive gene Stra6 in human cancers and its synergistic induction by Wnt-1 and retinoic acid. Cancer Res. 61, 4197–4205. 43. Goodman, A.B. (2006) Retinoid receptors, transporters, and metabolizers as therapeutic targets in late onset Alzheimer disease. J. Cell. Phys. 209, 598–603. 44. Barua, A.B., Furr, H.C. (1998) Properties of retinoids. Mol. Biotechnol. 10, 167–182. 45. Napoli, J.L. (1986) Quantification of physiological levels of retinoic acid. Method Enzymol. 123, 112–124 46. Barua, A.B., Furr, H.C. (1998) Properties of retinoids: Structure, handling, and preparation. Methods Mol. Biol. 89, 3–28. 47. Napoli, J.L., Horst, R.L. (1998) Quantitative analyses of naturally occurring retinoids. Methods Mol. Biol. 89, 29–40.

52

Kane and Napoli

48. Schmidt, C.K., Brouwer, A., Nau, H. (2003) Chromatographic analysis of endogenous retinoids in tissues and serum. Anal. Biochem. 315, 36–48. 49. Kane, M.A., Chen, N., Sparks, S., Napoli, J.L. (2005) Quantification of endogenous retinoic acid in limited biological samples by LC/MS/MS. Biochem. J. 388, 363–369. 50. Kane, M.A., Folias, A.E., Wang, C., Napoli, J.L. (2008) Quantitative profiling of endogenous retinoic acid in vivo and in vitro by tandem mass spectrometry. Anal. Chem. 80, 1702–1708. 51. Kane, M.A., Folias, A.E., Napoli, J.L. (2008) HPLC/UV quantitation of retinal, retinol, and retinyl esters in serum and tissues. Anal. Biochem. 378, 71–79. 52. Sucov, H.M., Evans, R.M. (1995) Retinoic acid and retinoic acid receptors in development. Mol. Neurobiol. 10, 169–184. 53. Bershad, S., Rubinstein, A., Paterniti, J.R., Le, N.A., Poliak, S.C., Heller, B., Ginsberg, H.N., Fleischmajer, R., Brown, W.V. (1985) Changes in plasma lipids and lipoproteins during isotretinoin therapy for acne. N. Engl. J. Med. 313, 981–985. 54. Rodondi, N., Darioli, R., Ramelet, A.-A., Hohl, D., Lenain, V., Perdrix, J., Wietlisbach, V., Riesen, W.F., Walther, T., Medinger, L., Nicod, P., Desvergne, B., Mooser, V. (2002) High risk for hyperlipidemia and the metabolic syndrome after an episode of hypertriglyceridemia during 13-cis retinoic acid therapy for acne: A pharmacogenetic study. Ann. Intern. Med. 136, 582–589. 55. Heliovaara, M. K., Remitz, A., Reitamo, S., Teppo, A.-M., Karonen, S.-L., Ebeling, P. (2007) 13-cis-Retinoic acid therapy induces insulin resistance, regulates inflammatory parameters, and paradoxically increases serum adiponectin concentration. Metabolism 56, 786–791. 56. McCormick, A.M., Kroll, K.D., Napoli, J.L. (1983) 13-cis-retinoic acid metabolism in vivo. The major tissue metabolites in the rat have the all-trans configuration. Biochemistry 22, 3933–3940. 57. Arnhold, T., Tzimas, G., Wittfoht, W., Plonait, S., Nau, H. (1996) Identification of 9-cis-retinoic acid, 9,13-di-cis-retinoic acid, and 14-hydroxy-4,14-retro-retinol in human plasma after liver consumption. Pharmacol. Lett. 59, 169–177. 58. Horst, R.L., Reinhardt, T.A., Goff, J.P., Nonnecke, B.J., Gambhir, V.K., Fiorella, P.D., Napoli, J.L. (1995) Identification of 9-cis,13-cis-retinoic acid as a major circulating retinoid in plasma. Biochemistry 34, 1203–1209.

59. Horst, R.L., Reinhardt, T.A., Goff, J.P., Koszewski, N.J., Napoli, J.L. (1995) 9,13Di-cis-retinoic acid is the major circulating geometric isomer of retinoic acid in the periparturient period. Arch. Biochem. Biophys. 322, 235–239. 60. Wang, Y., Chang, W.Y., Prins, G.S., van Breemen, R.B. (2001) Simultaneous determination of all-trans, 9-cis, 13-cis retinoic acid and retinol in rat prostate using liquid chromatography-mass spectrometry. J. Mass. Spectrom. 36, 882–888. 61. Gundersen, T.E., Bastani, N.E., Blomhoff, R. (2007) Quantitative high-throughput determination of endogenous retinoids in human plasma using triple-stage liquid chromatography/tandem mass spectrometry. Rapid Commun. Mass. Spectrom. 21, 1176–1186. 62. Luo, T., Wagner, E., Grun, F., Drager, U.C. (2004) Retinoic acid signaling in the brain marks the formation of optic projections, maturation of the dorsal telencephalon, and function of limbic sites. J. Comp. Neurol. 470, 297–316. 63. Wagner, M.A. (1997) Use of reporter cells to study endogenous retinoid sources in embryonic tissues. Methods Enzymol. 282, 98–107. 64. Zetterstrom, R.H., Lindqvist, E., Mata de Urquiza, A., Tomac, A., Eriksson, U., Perlmann, T., Olson, L. (1999) Role of retinoids in the CNS: Differential expression of retinoid binding proteins and receptors and evidence for presence of retinoic acid. Eur. J. Neurosci. 11, 407–416. 65. Wagner, M., Han, B., Jessell, T.M. (1992) Regional differences in retinoid release from embryonic neural tissue detected by an in vitro reporter assay. Development 116, 55–66. 66. National Research Council. (1995) Nutrient Requirements of Laboratory Animals, 4th ed., National Academy Press, Washington, D.C. 67. Biesalski, H.K. (1989) Comparative assessment of the toxicology of vitamin A and retinoids in man. Toxicology 57, 117–161. 68. Gundersen, T.E., Blomhoff, R. (2001) Qualitative and quantitative liquid chromatographic determination of natural retinoids in biological samples. J. Chromatogr. A 935, 13–43. 69. Wyss, R., Bucheli, F. (1997) Determination of endogenous levels of 13-cisretinoic acid (isotretinoin), all-trans-retinoic acid (tretinoin), and their 4-oxo metabolites in human and animal plasma by highperformance liquid chromatography with automated column switching and ultraviolet

Quantification of Endogenous Retinoids

70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

detection. J. Chromatogr. B Biomed. Sci. Appl. 700, 31–47. Hagen, J.J., Washco, K.A., Monning, C.A. (1996) Determination of retinoids by reversed-phase capillary liquid chromatography with amperometric electrochemical detection. J. Chromatogr. B Biomed. Sci. Appl. 677, 225–231. Sakhi, A.K., Gundersen, T.E., Ulven, S.M., Blomhoff, R., Laundanes, E. (1998) Quantitative determination of endogenous retinoids in mouse embryos by high-performance liquid chromatography with online solid-phase extraction, column switching, and electrochemical detection. J. Chromatogr. A 828, 451–460. Ulven, S.M., Gundersen, T.E., Weedon, M.S., Landaas, V.O., Sakhi, A.K., Fromm, S.H., Geronimo, B.A., Moskaug, J.O., Blomhoff, R. (2000) Identification of endogenous retinoids, enzymes, binding proteins, and receptors during early postimplantation development in the mouse: Important role of retinal dehydrogenase type 2 in synthesis of all-trans-retinoic acid. Dev. Biol. 220, 379–391. Toth, K., Stulik, K., Kutner, W., Feher, Z., Linder, E. (2004) Electrochemical detection in liquid flow techniques: Characterization and classification. Pure Appl. Chem. 76, 1119–1138. Napoli, J.L., Pramanik, B.C., Williams, J.B., Dawson, M.I., Hobbs, P.D. (1985) Quantification of retinoic acid by gas–liquid chromatography–mass spectrometry: Total versus all-trans retinoic acid in human plasma. J. Lipid Res. 26, 387–392. Napoli, J.L. (1990) Quantification and characteristics of retinoid synthesis from retinol and β-carotene in tissue fractions and established cell lines. Methods Enzymol. 189, 470–482. Schmidt, C.K., Volland, J., Hamscher, G., Nau, H. (2002) Characterization of a new endogenous vitamin A metabolite. Biochim. Biophys. Acta 1583, 237–251. McCaffery, P., Evans, J., Koul, O., Volpert, A., Reid, K., Ullman, M.D. (2002) Retinoid quantification by HPLC/MSn. J. Lipid Res. 43, 1143–1149. Chithalen, J.V., Luu, L., Petkovich, M., Jones, G. (2002) HPLC–MS/MS analysis of products generated from all-trans-retinoic acid using recombinant human CYP26A. J. Lipid Res. 43, 1133–1142. Ruhl, R. (2006) Method to determine 4-oxo-retinoic acids, retinoic acids, and retinol in serum and cell extracts by liquid chromatography/diode-array detection

80.

81.

82.

83.

84.

85.

86.

87. 88.

89.

53

atmospheric pressure chemical ionization tandem mass spectrometry. Rapid Commun. Mass Spectrom. 20, 2497–2504. David, D.C., Hoerndli, F., Götz, J. (2005) Functional Genomics meets neurodegenerative disorders Part I: Transcriptomic and proteomic technology. Prog. Neurobiol. 76, 153–168. Sadowski, P.G., Dunkley, T.P., Shadforth, I.P., Dupree, P., Bessant, C., Griffin, J.L., Lilley, K.S. (2006) Quantitative proteomic approach to study subcellular localization of membrane proteins. Nat. Protoc. 1, 1778–1789. Ferrara, C.T., Wang, P., Neto, E.C., Stevens, R.D., Bain, J.R., Wenner, B.R., Ilkayeva, O.R., Keller, M.P., Blasiole, D.A., Kendziorski, C., Yandell, B.S., Newgard, C.B., Attie, A.D. (2008) Genetic networks of liver metabolism revealed by integration of metabolic and transcriptional profiling. PLoS Genet. 4, e1000034. Furr, H.C., Barua, A.B., Olson, J.A. (1994) Analytical methods. In: Sporn, M.B., Roberts, A.B., Goodman, D.S. (eds.), The Retinoids: Biology, Chemistry, and Medicine, 2nd ed., Raven Press, New York, pp. 195–196. Hubbard, R. Brown, P.K., Bownds, D. (1971) Methodology of vitamin A and visual pigments. Methods Enzymol. 18C, 615–653. Robeson, C.D., Cawley, J.D., Weisler, L., Stern, M.H., Edinger, C.C., Checkak, A.J. (1955) Chemistry of vitamin A. XXIV. The synthesis of geometric isomers of vitamin A via methyl β-methylglutaconate. J. Am. Chem. Soc. 77, 4111–4119. Robeson, C.D., Blum, W.P., Dieterle, J.M., Cawley, J.D., Baxter, J.G. (1955) Chemistry of vitamin A. XXV. Geometrical isomers of vitamin A aldehyde and an isomer of its α-ionone analog. J. Am. Chem. Soc. 77, 4120–4125. Olson, J.A. (1990) Vitamin A. In: Machlin, L.J. (ed.), Handbook of Vitamins, MarcelDekker, New York, pp. 1–57. Schwieter, U., Isler, O. (1967) Vitamin A and carotene: Chemistry. In: Sebrell, W.H., Jr., Harris, R.H. (eds.), The Vitamins, 2nd ed., vol. 1, Academic Press, New York, pp. 5–101. von Planta, C., Schweiter, U. Choparddit-Jean, L., Ruegg, R., Kofler, M., Isler, O. (1962) Physikalische eigenschaften von isomeren vitamin-A- and vitamin -A2verbindungen [Physical properties of isomeric vitamin A and vitamin A2 compounds]. Helv. Chim. Acta. 45, 548–561.

54

Kane and Napoli

90. Rao, M.S.S., John, J., Cama, H.R. (1972) 96. Blumberg, B., Bolado, J., Jr., Derguini, F., Studies on vitamin A2 : Preparation, properCraig, A.G., Moreno, T.A., Chakravarti, D., Heyman, R.A., Buck, J., Evans, R.M. (1996) ties, metabolism, and biological activity of 4Novel retinoic acid receptor ligands in Xenooxoretinoic acid. Int. J. Vitamin. Nutr. Res. pus embryos. Proc. Natl. Acad. Sci. USA 93, 42, 368–370. 4873–4878. 91. Vahlquist, A., Torma, H., Rollman, O., Andersson, E. (1990) High-performance liq- 97. White, J.A., Ramshaw, H., Taimi, M., Stangle, W., Zhang, A., Everingham, uid chromatography of natural and synthetic S., Creighton, S., Tam, S.P., Jones, G., retinoids in human skin samples. Methods Petkovich, M. (2000). Identification of Enzymol. 190, 163–174. the human cytochrome P450, P450RAI92. Ross, A.C. (1981) Separation of long-chain 2, which is predominantly expressed in the fatty esters of retinol by high-performance adult cerebellum and is responsible for allliquid chromatography. Anal. Biochem. 115, trans-retinoic acid metabolism. Proc. Natl. 324–330. Acad. Sci. USA 97, 6403–6408. 93. Tsukida, K., Ito, M., Tanaka, T., Yagi, I. (1985) High-performance liquid chromato- 98. Taimi, M., Helvig, C., Wisniewski, J., Ramshaw, H., White, J., Amad, M., Korczak, graphic and spectroscopic characterization of B., Petkovich, M. (2004) A novel human stereoisomeric retinaloximes: Improvements cytochrome P450, CYP26C1, involved in in resolution and implications of the method. metabolism of 9-cis and all-trans isoJ. Chromatogr. 331, 265–272. mers of retinoic acid. J. Biol. Chem. 279, 94. Van Kuijk, F.J.G.M., Handleman, G.J., 77–85. Dratz, E.A. (1985) Rapid analysis of the major classes of retinoids by step gradi- 99. Williams, J.B., Pramanik, B.C., Napoli, J.L. (1984) Vitamin A metabolism: Analysis of ent reversed-phase high performance liqsteady-state neutral metabolites in rat tissues. uid chromatography using retinal (O J. Lipid. Res. 25, 638–645. ethyl) oxime derivatives. J. Chromatogr. 348, 100. Dorsey, J.G. (2000) Column theory 241–251. and resolution in liquid chromatography. 95. Landers, G.M., Olson, J.A. (1988) Rapid, In: Meyers, R.A. (ed.), Encyclopedia of simultaneous determination of isomers of Analytical Chemistry, Wiley, New York, retinal, retinal oxime, and retinol by highpp. 11334–11342. performance liquid chromatography. J. Chromatogr. 438, 383–392.

Chapter 2 Culture of Highly Differentiated Human Retinal Pigment Epithelium for Analysis of the Polarized Uptake, Processing, and Secretion of Retinoids Jane Hu and Dean Bok Abstract The retinal pigment epithelium (RPE) occupies a strategic position within the eye, given its location between the neurosensory retina and the vascular bed (choroid) that nourishes the photoreceptor cells (rods and cones). Among the many attributes of this versatile monolayer of cells is its unique ability to convert vitamin A (retinol) into the prosthetic group (11-cis-retinal) for the rod and cone opsins, the photopigments essential for vision. It does so by absorbing retinol via a receptor-mediated process that involves the interaction of a carrier protein secreted by the liver, retinol-binding protein (RBP), and a receptor/channel that is the gene product of STRA6 (stimulated by retinoic acid 6). Following its uptake through the basolateral plasma membrane of the RPE, retinol encounters a brigade of binding proteins, membrane-bound receptors, and enzymes that mediate its multi-step conversion to 11-cis-retinal and the transport of this visual chromophore to the light-sensitive photoreceptor cell outer segment, the portion of the cell that houses the phototransduction cascade. This process is iterative, repeating itself via the retinoid visual cycle. Most of the human genes that code for this cohort of proteins carry diseasecausing mutations in humans. The consequences of these mutations range in severity from relatively mild dysfunction such as congenital stationary night blindness to total blindness. The RPE, although post-mitotic in situ, is capable of proliferation when removed from its native milieu. This offers one the opportunity to study the retinoid visual cycle in modular form, providing insights into this intriguing process in health and disease. This chapter describes a cell culture method whereby the entire visual cycle can be created in vitro. Key words: Retinoid visual cycle, retinal pigment epithelium, 11-cis-retinal, all-trans-retinal, inherited retinal disease, Stargardt macular dystrophy, Leber congenital amaurosis, age-related macular degeneration.

H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_2, © Springer Science+Business Media, LLC 2010

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1. Introduction The retinal pigment epithelium (RPE) is a morphologically and functionally polarized monolayer of cells inserted between the neurosensory portion of the vertebrate retina and a highly pigmented vascular bed called the choroid. The light-sensitive components of the neurosensory retina’s rod and cone photoreceptor cells (outer segments) are in close association with the apical plasma membrane of the RPE. The photoreceptor cells (hereafter called the photoreceptors) are dependent upon the RPE for their function and survival. The RPE provides oxygen and nutrients to the photoreceptors, which it obtains from a large-bore, fenestrated capillary bed called the choriocapillaris, the innermost layer of the choroid (Fig. 2.1). The choriocapillaris is separated from the RPE by a pentalaminar extracellular matrix called Bruch’s membrane. The photoreceptors, RPE, Bruch’s membrane, and choriocapillaris are so intimately entwined functionally that two or more members of this complex are often collectively involved in disease processes. These diseases can be caused by gene mutations expressed either cell autonomously or systemically. Striking examples of this are observed in two monogenic diseases. The first example is recessive Stargardt macular dystrophy (STGT2) where a defective gene is expressed in photoreceptors (1, 2), with a bystander effect on the RPE. The second is a subset of Leber congenital amaurosis (LCA2) where the mutant gene is expressed in the RPE (3, 4), with a bystander effect on the photoreceptors. Finally, age-related macular degeneration (AMD), a complex trait disease, results from the interplay of genetic, environmental, behavioral (such as smoking), and dietary factors (5, 6). As a result, all of the components in the complex are affected. A common player in these three diseases is a class of lipid molecules collectively named retinoids because their parent compound is retinol (vitamin A). Before we describe the mechanisms for these diseases in more detail, we will briefly describe the retinoid visual cycle (Fig. 2.1). The retinoid visual cycle (7) requires a supply of retinol from the blood. This is accommodated through a receptor-mediated process that transpires at the basolateral plasma membrane of the RPE (8). The carrier protein for retinol in the blood is retinol-binding protein (RBP), whose main source is the liver (9), from which it is secreted in its ligand-bound form (holo-RBP). RBP (∼21 kDa) is secreted as a 1:1 complex with transthyretin (TTR), a homotetramer that carries thyroxin. The combined mass of the RBP/TTR complex is ∼75 kDa, which has a diameter sufficient to prevent kidney glomerular filtration into the urine. The RBP/TTR complex passes through fenestrations in

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Fig. 2.1. Diagram of the retinoid visual cycle showing the RBP receptor and most of the binding proteins and enzymes that mediate this complex process. All-trans-retinol (vitamin A) is extracted from retinol-binding protein (RBP) by an RBP receptor in the basolateral membrane of the RPE. RBP leaves large-bore capillaries in the choroid to gain access to the receptor. Following entry into the RPE, retinol is bound by cellular retinaldehyde binding-protein (CRBP), which then delivers it to lecithin retinol acyltransferase (LRAT). LRAT then adds a fatty acid to the vitamin. The resulting retinyl esters serve as substrates for the retinoid isomerase (also known as RPE 65), which hydrolyzes the ester bond and converts all-trans-retinol to 11-cis-retinol. 11-cis-retinol is bound by cellular retinaldehyde binding-protein (CRALBP), which serves as substrate carrier for 11-cis-retinol dehydrogenase during the oxidation of 11-cis-retinol to 11-cis-retinaldehyde, the visual chromophore for rod and cone photopigments. 11-cis-retinaldehyde exit across the apical plasma membrane of the RPE is mediated by interphotoreceptor retinoid-binding protein (IRBP) through an undetermined mechanism. Finally the chromophore is delivered to the rod and cone photoreceptor outer segments, the repositories of rhodopsin and the cone photopigments, respectively. Modified from Bok (1993) (21) J. Cell Sci. with permission.

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the endothelial lining of the choriocapillaris (10). These fenestrations face Bruch’s membrane, and the porosity of a healthy Bruch’s membrane is sufficient to allow diffusion of molecules of this mass. The RBP binds to a receptor (RBPR) in the basolateral plasma membrane of the RPE. The gene coding for this receptor has recently been identified (11) as STRA6 (stimulated by retinoic acid 6). The RBPR is a highly hydrophobic, putative nine-pass receptor/channel. The importance of this receptor for tissue survival in humans, not only in the eye, but systemically, has been emphasized through its recently documented involvement in Matthew-Wood syndrome (12, 13), which features severe developmental defects in multiple organ systems. Following entry into the RPE via the RBPR, retinol is processed by a cohort of binding proteins and enzymes. The final product of this process is 11-cis-retinaldehyde (11-cis-RAL), the chromophore of the rod and cone visual pigments. Key steps in this process include the fatty acyl esterification of retinol (14–16) by lecithin retinol acyltransferase (LRAT). These retinyl esters serve as substrates for an isomerohydrolase (17), which utilizes free energy stored in the ester bond to drive isomerization of retinol from the all-trans to the 11-cis configuration (18). RPE65 has recently been identified as the isomerhydrolase (19–22). Oxidation of 11-cis-retinol to 11-cis-retinal completes the process and transport out of the RPE across the extracellular space between RPE and photoreceptors (the subretinal space) and into the photoreceptors’ lightsensitive outer segments is completed by an additional complement of binding proteins (Fig. 2.1). Having briefly described retinoid uptake, processing and transport within the choroid and retina, we can now explain how failure of or deficiencies in these processes impact the inherited retinal diseases mentioned earlier. This will provide the rationale for this chapter, namely a reliable in vitro system for analysis of the visual cycle in health and disease. Recessive Stargardt macular dystrophy is caused by the malfunction of a phospholipid/all-transretinaldehyde translocator located in the light-sensitive outer segments of rod and cone photoreceptors (2, 23). This translocator, which is a member of the ABC protein family, normally translocates N-retinylidine phosphatidylethanolamine (N-RetPE), a byproduct of phototransduction, from the inner (lumen facing) leaflet of the outer segment disc membrane bilayer into the outer leaflet (cytosol facing), whereupon the aldimine linkage between phosphatidyl ethanolamine (PE) and all-trans-retinaldehyde is hydrolyzed. The aldehyde is then reduced by a dehydrogenase to reform retinol (2). Retinol leaves the photoreceptor and is returned to the RPE, thereby beginning the visual cycle anew. When this mechanism is retarded or absent, PE binds two alltrans-retinaldehyde molecules and this bis-retinoid–PE complex (A2PE) remains incarcerated in the discs until it is liberated by

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phagocytosis of the discs by the RPE (2). Within the acidic lysosomal compartment of the RPE, A2PE is converted to A2E (essentially removing the phospholipid head group), a toxic, detergent-like molecule that eventually poisons the RPE (24). The adjacent photoreceptors, which are dependent upon the RPE for their nurture, die collaterally. For Leber congenital amaurosis, the process is simpler. In this case, the isomerohydrolase (RPE-65) is absent or defective and the RPE cannot supply the 11-cis-retinaldehyde prosthetic group to the rod and cone apo-opsins. Consequently, light sensitivity is severely impaired or absent (4). Recent attempts at replacement gene therapy with the aid of an adeno-associated virus vector in patients with this recessive disease have met with promising results (25–27). The role of defective retinoid metabolism in the context of age-related macular degeneration is probably, in part, A2E related as well. RPE tissue culture experiments demonstrate that, when the RPE is experimentally loaded with A2E and illuminated with light in the blue portion of the visible spectrum, A2E is oxidized into epoxides and furanoids, which are apparently able to escape from the RPE cell (28). These compounds are capable of activating complement C3, which lies at the crossroads of three complement cascades (classic, lectin, and alternative pathways). This local, inappropriate activation of C3 probably places the RPE under inappropriate complement attack if it occurs in the context of predisposing variations in genes such as complement factor H, which serves as a fluid phase, negative regulator of complement (29–32). The RPE in situ is post-mitotic, functioning in a healthy individual from birth through the full life span without dividing. However, when placed in the appropriate environment, the RPE can re-enter the mitotic cycle. This capability, which can be detrimental during retinal injury such as a tear of the neurosensory retina, can be used to our advantage in several ways. Among these is the opportunity to collect RPE cells from animals or human donor tissue and to culture these cells to a high level of differentiation, whereby components of the retinoid visual cycle and other aspects of RPE function can be studied in isolation from the choroid and neurosensory retina (33, 34). The purpose of this chapter is to provide a detailed description of the conditions required for a high state of differentiation of the RPE in culture, including all of the features required for the vectorial uptake, processing, and release of retinoids. The scientific literature contains many reports on the culture and use of human and animal RPE for a variety of experimental procedures. However, the quality of the cultures used in these reports varies considerably. Moreover some of these studies utilize immortalized cell lines that have been derived by the introduction

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of agents such as viral large T antigen or cell lines that have arisen spontaneously in culture. The quality of results obtained from these experiments depends upon the specific process under investigation and whether or not the RPE cells have achieved a proper level of differentiation. If one is interested in transepithelial transport, an important criterion is the integrity of the barrier properties of the cultured monolayer. This is typically measured by the transepithelial resistance (TER), which is determined in part by the quality and number of tight junctions between adjacent cells. Moreover, the polarized expression of key ion transporters such as the Na, K ATPase is important, because this transporter sets up the ion gradients that drive other transporters in the plasma membrane. Most reports have used RPE cultures that do not display proper expression of Na, K ATPase, which in the RPE and choroid plexus epithelium is uniquely polarized to the apical plasma membrane. Studies involving the retinoid visual cycle as discussed in this chapter require expression of all key membrane receptors, binding proteins, and enzymes that serve the visual cycle. Among the transformed cell lines, none express the entire cohort of components sufficient to carry out the entire visual cycle or the vectorial uptake of retinol and secretion of 11-cis-retinal. Conversion of retinol into 11-cis-retinal and apical expression of Na, K ATPase are two features of the RPE that represent the gold standard in RPE culture. Obviously, other attributes are required as well but, in our experience, if the cells exhibit these two properties, the prognosis for success is high. Figure 2.2 illustrates a welldifferentiated culture in which the Na, K ATPase is largely apical. Figure 2.3 shows a culture in which the membrane receptor for RBP is largely polarized to the basolateral membrane. Finally, Fig. 2.4 is an electron micrograph of highly differentiated RPE. It

Fig. 2.2. Expression of Na, K ATPase by cultured human fetal RPE. The RPE cells were cultured on Millicell wells with a polycarbonate substrate for 2 months until they reached full differentiation. The upper portion of the image is a cross section of the RPE monolayer using the Phi-Z mode of a Zeiss 210 laser scanning confocal microscope. The lower portion of the image shows a single optical section taken in the X–Y axis. Na, K ATPase is largely polarized to the apical plasma membrane in properly differentiated RPE cells, a feature that it shares uniquely with the choroid plexus epithelium. Modified from Hu and Bok (2000) Molecular Vision with permission.

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Fig. 2.3. Expression of the RBP membrane receptor (STRA6) in cultured human RPE. The image in the upper panel, taken in the X–Y axis, shows a bright field image of the antibody-stained cells illustrated in the lower panel. RPE cell nuclei in the upper panel are shown as white oblate spheres. The lower panel is an optical confocal section of the same cells in the X–Y axis. Outlines of cell boundaries are evident due to basolateral labeling of the RBP receptor. Numerous membrane-associated and cytoplasmic vesicles are a typical feature of well-differentiated cultured RPE cells and of RPE cells stained in intact tissue (see (11), Figure 5A and B).

Fig. 2.4. Electron micrograph of cultured human RPE grown to full differentiation in a Millicell with a nitrocellulose substrate (NC). The cells show heavy melanin pigmentation and a rich array of apical microvilli.

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features the morphological hallmarks on this cell type, including a rich population of microvilli on its apical surface and abundant melanin granules. RPE cells, appropriately differentiated in culture, are fully capable of carrying out the entire visual cycle in chambers whose porous support for the cell monolayer are made of nitrocellulose or polycarbonate (33, 34). To provide an appropriate surface for cell adhesion, the support material is pre-coated with extracellular matrix components. The first evidence regarding the suitability of this culture system for the study of the visual cycle in vitro was provided by Carlson and Bok (33), who tested this process in RPE cultured from fetal calf eyes. Figure 2.5 shows the results of an experiment utilizing fetal bovine RPE grown to confluent monolayers in Millicell chambers. Holo-retinol-binding protein (RBP) saturated with 3 H-retinol was applied at a concentration of 3 μM in the culture medium bathing the basal surface of the monolayer. The apical culture medium contained various retinoid-binding proteins: either 3 μM interphotoreceptor retinoid-binding protein (IRBP), 3 μM apo-cellular retinaldehyde-binding protein (CRALBP), 3 μM apo-retinol-binding protein (RBP), 90 μM

Fig. 2.5. Release of 3 H-labeled 11-cis-retinaldehyde from bovine RPE cells into the apical culture medium as a function of various retinoid carrier proteins in the apical medium. Holo-RBP carrying 3 H retinol was added to the medium bathing the basal surface of the cultured monolayer. After 16 h, the apical medium was analyzed by HPLC and an in-line liquid 3H 11-cis-retinaldehyde in apical medium; 3H all-trans-retinal in apical medium, 3H allscintillation counter. trans-retinol in basal medium, 3H retinyl palmitate intracellular. Modified from Carlson and Bok (1992) Biochemistry with permission.

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bovine serum albumin (BSA), or no binding protein. After 16 h of incubation retinoids were extracted from the basal medium, from the RPE cells and from the apical medium and analyzed by HPLC and in-line liquid scintillation counting. Interestingly, only IRBP efficiently promoted the release of 3 H-11-cis-retinal, the chromophore of the rod and cone visual pigments, even though CRALBP is the natural, intracellular-binding protein for 11-cis-retinaldehyde. It is thought that the apical membrane of the RPE carries a specific receptor for IRBP that mediates this binding protein-specific process. We have also performed these experiments on highly differentiated human RPE derived from fetuses (34) or young adults, analyzing the release of 11-cisretinaldehyde by identifying and monitoring the retinoids via their specific absorption spectra (Fig. 2.6). Again, the presence of IRBP was essential for the efficient release of 11-cis-retinal into the apical culture medium. Recently, Maminishkis et al. (35) also published a paper in which human fetal RPE cells were used to produce highly differentiated RPE monolayers. We have tested this method and found that it is capable of producing high quality monolayers. One advantage of this method is that it utilizes culture medium ingredients that are all available commercially. However, we have not tested these cells in terms of their ability to carry out all steps of the visual cycle. The following is a detailed description of human RPE cultured in laminin-coated Millicell chambers and suitable for the in

Fig. 2.6. Release of 11-cis-retinaldehyde from human RPE cells into the apical culture medium. Holo-RBP (5 μM) was added to the medium bathing the basal surface of the cultured monolayer. Either 1% bovine serum albumin (BSA) or 10 μM IRBP was present in the apical medium.

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vitro analysis of the retinoid visual cycle. In order to expand the RPE cells that are initially collected from donor eyes, we grow them in a culture medium that is low in calcium (0.05 mM, see Section 2.1.2). The RPE cells proliferate and reach about 60% confluence at 10 days and then, because of the low calcium concentration, the lightly attached cells begin to release into the medium. Batches of non-attached cells are collected and cryopreserved for future use. This process of amplification is continued until the cells exhibit signs of reduced size and viability. Cells for experiments are thawed and cultured in medium containing normal calcium levels (see Section 2.1.3).

2. Materials 2.1. Primary Human RPE Culture 2.1.1. Human Donor Eyes

Human fetal eyes ranging from 18 to 21 weeks gestation are obtained from Advanced Bioscience Resources (Alameda, CA). The donor eyes are placed in transport medium in 50 ml tubes containing Eagle’s minimal essential medium (MEM; SigmaAldrich, St. Louis, MO) with 5% heat-inactivated calf serum and 1% penicillin–streptomycin. The tubes are packed on ice and shipped overnight.

2.1.2. Culture Medium for Cell Expansion

RPE cells are first grown in expansion culture medium in “low” calcium (0.05 mM) to prevent attachment and to allow for proliferation. Spherical cells produced in this manner become suspended in the culture medium and they can be collected readily and frozen for future use. For each ingredient below, we indicate the strength of the stock solution from which 1 l of culture medium is prepared. For example, “1000× aqueous solution” means that one would add 1 ml of a stock solution that is 1000 times more concentrated than the final solution. 1. MEM culture medium with Joklik modification (M8028, Sigma-Aldrich, St. Louis, MO). 2. CaCl2 ·H2 O (7.92 mg/l; 5.4 × 10−5 M). 3. TAPSO-free acid (5.0 g/l; 1.9 × 10–2 M). 4. ZnSO4 ·7H2 O (0.1438 mg/l; 5.0 × 10−7 M, 1000× aqueous solution). 5. CuSO4 5H2 O (0.2496 μg/l; 1.0 × 10−9 M, 1000× aqueous solution).

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6. MnCl2 4H2 O (9.9 × 10−5 mg/l; 5.0 × 10−10 M, 1000× aqueous solution). 7. Selenious acid, sodium salt (2.6 μg/l; 1.5 × 10−8 M, 1000× aqueous solution). 8. Hydrocortisone (10.0 μg/l; 2.8 ×10−8 M, 200,000× stock in absolute ethanol). 9. Calf serum, heat inactivated (1.0% by volume). 10. Linoleic acid with albumin [0.0842 mg/l; 3.0 × 10−7 M, 1000× stock solution in Hank’s balanced salt solution (HBSS)]. 11. Insulin (5.0 mg/l; ∼8.3 × 10−7 M, 100× stock solution in HBSS). 12. Transferrin (5.0 mg/l; ∼6.3 × 10−8 M, 100× stock in HBSS). 13. Putrescine.2HCl (0.3 mg/l; 1.86 × 10−6 M, 1000× in HBSS). 14. L-Ascorbic acid (45.0 mg/l; 2.6 × 10−4 M, 1000× aqueous solution). 15. L-Glutamine (292.0 mg/l; 2.0 mM, 100× aqueous solution). 16. Triiodothyronine (6.5 × 10−6 mg/l; 1.0 × 10−11 M, 1000× in HBSS). 17. Bovine retinal extract (1% by volume, see Section 3.1.4). 18. Alanine (0.02 g/l; 2.2 × 10−4 M, 200× aqueous solution). 19. Asparagine (0.025 g/l; 1.7 × 10−4 M, 100× aqueous solution). 20. Aspartic acid (0.02 g/l; 1.5 × 10−4 M, 100× aqueous solution). 21. Glutamic acid (0.03 g/l; 2.0 × 10−4 M, 100× aqueous solution). 22. Glycine (0.025 g/l; 3.3 × 10−4 M, 200× aqueous solution). 23. Proline (0.02 g/l; 1.7 × 10−4 M, 200× aqueous solution). 24. Serine (0.015 g/l; 1.4 × 10−4 M, 200× aqueous solution). 25. Biotin (0.1 mg/l; 4.1 × 10−7 M, 2000× aqueous solution). 26. Oxaloacetic (0.15 g/l; 1.1 × 10−3 M). 27. Thymidine (0.3 mg/l; 1.2 × 10−6 M, 2000× aqueous solution). 28. Ferric nitrate (0.1 mg/l; 2.5 × 10−7 M, 2000× aqueous solution freshly made).

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2.1.3. Culture Medium and Chambers for Cell Differentiation

Culture medium for cell differentiation is composed of MEM with Earle’s salts (M 2279; Sigma Chemical) with the addition of above components 3–28. 1. Millicell-HA culture inserts (Millipore, Bedford, MA). 2. Mouse laminin (100 μg/ml in MEM; BD Bioscience, Bedford, MA).

2.1.4. Bovine Retinal Extract

2.2. Interphotoreceptor Retinol-Binding Protein (IRBP) Purification

We have determined that bovine retinal extract enhances the development of a robust transepithelial resistance (TER) across RPE monolayers, thereby making them highly suitable for transport studies. 1. Amicon ultra concentrator (100,000 MWCO, Millipore, MA). 2. Affinity column containing concanavalin A–Sepharose 4B matrix (Amersham Biosciences). 3. Binding buffer (20 mM Tris–HCl, 0.5 M NaCl, pH 7.4). 4. Elution buffer [200 mM of methyl-α-D-mannopyranoside in phosphate-buffered saline (PBS)].

2.3. Recombinant Retinol-Binding Protein (RBP) and 6Histidine-Tagged Transthyretin (6His-TTR) Production in Escherichia coli

2.4. Recombinant Retinol-Binding Protein (RBP) Refolding and Purification

1. RBP or 6His-TTR recombinant plasmids [Dr. Wayne Hubbell (TTR) or Dr. Dean Bok (RBP), University of California, Los Angeles]. 2. E. coli strain BL21 Star (DE3) (Invtritrogen, CA). 3. Luria broth (LB) (100 μg/ml).

medium

with

added

ampicillin

4. Isopropyl β-D-1-thiogalactopyranoside (IPTG; 4 mM for RBP and 1 mM for TTR) for induction. 1. Lysis buffer [1 mM dithiothreitol (DTT), 50 mM Tris, 2 mM ethylenediaminetetraacetic acid (EDTA), 0.1% triton X-100, pH 7.5]. 2. Lysis buffer with 8 M urea and 5 mM DDT. 3. Dim red light (Kodak Safelight Filter No. 2). 4. Refolding buffer (20 mM Tris, 20 mM NaCl, 0.1 mM EDTA, 5 mM reduced glutathione, and 1 mM of oxidized glutathione, pH 8.6). 5. PBS.

2.5. TTR-Affinity Column

1. Ni-NTA resin (Qiagen, Valencia, CA) for purification of 6His-TTR. 2. Coupling buffer (0.1 M NaHCO3 , 0.5 M NaCl, pH 8.3).

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3. Imidazol (150 mM). 4. Cyanogen bromide (CNBr)-activated Sepharose 4B with 1 mM HCl. 5. Blocking buffer (0.1 M Tris, pH 8) for 2 h at room temperature. 6. Acetate buffer (0.1 M Na acetate, 0.5 M NaCl, pH 4). 2.6. Incubation Conditions and Sample Collection for Retinoid Processing and Secretion

1. Dulbecco’s minimal essential medium (DMEM; SigmaAldrich, St. Louis, MO).

2.7. Retinoid Extraction and Analysis

1. Normal-phase high performance liquid chromatography (HPLC) column (Zorbax RX-SIL, 5 μm, and Model1100 HPLC system equipped with ultraviolet photodiode array detector; Agilent, Wilmington, DE).

2. Hydroxylamine (100 mM).

2. Gradient solution (0.2–10% dioxane in hexane).

3. Methods 3.1. Primary Human RPE Culture 3.1.1. Human Donor Eyes

1. Trim away excess tissue from eyes. 2. Irrigate briefly with normal saline. 3. Disinfect by immersing in detergent-free betadine for 10 s. 4. Gripping eyes firmly with hemostat, irrigate copiously with normal saline. 5. After transferring eyes to fresh gauze, open globes with sharp blade, entering slightly posterior to limbus. 6. Use curved iris scissors to trim away cornea and iris with lens. 7. Cut away optic nerve by approaching posteriorly and cutting through the full thickness of the eye wall to facilitate removal of the retina. 8. Transfer eyecups to 60-mm dishes with Hanks BSS (Caand Mg-free HBSS used throughout). 9. Remove retinas. 10. Cut each eyecup into quadrants. 11. Using two pairs of fine forceps, dissect pieces of choroidRPE free from the sclera. 12. Transfer pieces of choroid-RPE to fresh HBSS (2–4 pieces per 60-mm dish).

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13. Under dissecting microscope, use fine forceps to peel sheets of RPE away from the choroid. Cut away and discard any choroidal contaminants that adhere to the RPE (see Note 1). 14. Collect RPE explants in iced 15-ml tube containing HBSS. 15. When all explants have been collected, carefully remove HBSS by pipetting. Replace with fresh HBSS, cap tube securely, and agitate gently to rinse explants well. 16. Concentrate explants by allowing them to settle or by centrifuging and rewash in fresh HBSS as before. 17. Repeat HBSS wash one more time. 3.1.2. Culture Medium for Cell Expansion

1. Using 10-ml pipette, add low calcium growth medium to tube, resuspend explants, and transfer to culture dish (two eyes per 100-mm dish). 2. Place the explants in a 37◦ C incubator with 5% CO2 until new cells release into the medium in about 8–10 days and the cultures reach 60–80% confluence. 3. Collect the non-attached cells (floaters) for cryopreservation and for cell differentiation.

3.1.3. Culture Medium and Chambers for Cell Differentiation

1. Pre-wet Millicell-HA culture inserts (Millipore, Bedford, MA) by adding sterile water to the inserts and then withdrawing it. 2. Coat the culture inserts with 100 μl of mouse laminin (100 μg/ml in MEM; BD Bioscience, Bedford, MA). 3. Let the inserts dry under sterile conditions overnight. 4. Resuspend the RPE cells from the explanted culture in culture medium for cell differentiation, which contains 10% heat-inactivated calf serum. 5. Place 200,000 cells into each insert and switch to 1% heatinactivated calf serum after 1 week. Continue the cultures until they become confluent and differentiated in about 2 months. A good index of full differentiation is a TER of >500  cm2 and melanization. The ultimate proof for full differentiation is vectorial uptake of retinol from RBP applied to the basal side of the monolayer and secretion of 11-cis-retinal into the apical medium in the presence of IRBP.

3.1.4. Bovine Retinal Extract

On ice 1. use a sonicator equipped with a micro-tip and sonicate 12 fresh bovine retinas in 100 ml calcium- and magnesium-free Ringer’s buffer applying six, 10 s bursts at 75–100 W with 10 s of cooling between each burst;

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2. stir in cold, protected from light, for 2 h to overnight; 3. centrifuge at 4◦ C for 20 min at 17,300×g or until clear and collect the supernatant; 4. freeze aliquots and store at −80◦ C. 3.2. Interphotoreceptor Retinol-Binding Protein (IRBP) Purification

1. The retinas are removed from bovine eyes and the eyecups are washed three times with PBS. 2. The bovine retina interphotoreceptor matrix (IPM) wash is collected from the eyecups and concentrated using an Amicon ultra concentrator. 3. The buffer is changed to binding buffer for a concanavalin A affinity column. 4. The pre-washed Con A–Sepharose 4B matrix is mixed with the concentrated IPM wash overnight on a rotator at 4◦ C. 5. The column is packed and the flow-through is collected, the column is washed three times with wash buffer. 6. The IRBP is then eluted with 200 mM of methyl-α-Dmannopyranoside in wash buffer.

3.3. Recombinant Retinol-Binding Protein (RBP) and 6His-TTR Production in E. coli

1. Perform a fresh E. coli transformation with an RBP or 6HisTTR recombinant plasmid using E. coli strain BL21 Star (DE3). 2. Prepare overnight culture in 10 ml of LB medium with added ampicillin (100 μg/ml) and one colony of the E. coli cells. 3. Add 5 ml of overnight culture to 500 ml LB medium containing ampicillin and incubate at 37◦ C until the culture density reaches mid-log (OD ∼0.7, 3–4 h). 4. The culture is induced with IPTG (4 mM for RBP and 1 mM for TTR) and continued overnight at 30◦ C in a shaker. 5. The expression of RBP or TTR is determined by western blot.

3.4. Recombinant Retinol-Binding Protein (RBP) Refolding and Purification

1. Centrifuge overnight cultures at 5000 rpm. 2. Discard the supernatant and resuspend cells in 35 ml lysis buffer. 3. Let lysis reaction proceed at room temperature for 10 min. 4. Sonicate briefly and spin the lysate at 10,000×g for 10 min. 5. Discard the supernatant; the protein is in the white pellet. 6. Resuspend the pellet in 4 ml of lysis buffer containing 8 M urea and 5 mM DDT. 7. Deoxygenate the refolding buffer with nitrogen for 10 min.

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8. All procedures are performed under dim red light from this point. 9. Add all-trans-retinol to refolding buffer one drop at a time to the center of the solution while stirring (see Note 2). 10. Once the pellet has dissolved (it takes about 1 h), using a pipette, slowly add the solubilized protein into 50 ml of refolding buffer while stirring. 11. Continue to stir for 15–30 min. 12. Load the refolding buffer with the RBP protein to the TTR column. 13. Reload the flow-through once again onto the TTR column. 14. Wash the column with PBS until no unbound protein comes out of the wash buffer (or when the absorbance of 280 nm peak reaches baseline). 15. Elute the sample with distilled H2 O and collect 1 ml fractions. 16. Check the fractions with spectrophotometer to insure that the retinol-binding protein is saturated with retinol (1:1 ratio of 330 nm peak due to retinol and protein, 280 nm peak due to protein – see Note 4). 3.5. TTR-Affinity Column

1. Purify recombinant 6His-TTR with Ni-NTA resin following manufacturer’s protocol for E. coli lysates under native conditions. 6His-TTR is eluted with 150 mM imidazol. 2. Dialyze 6His-TTR overnight at 4◦ C with coupling buffer (see Note 3). 3. Wash 1 g of CNBr-activated Sepharose 4B with 1 mM HCl. 4. Mix 6His-TTR in coupling buffer with the gel suspension in an end-to end mixer overnight at 4◦ C. 5. Transfer gel to blocking buffer for 2 h at room temperature. 6. Wash away excess adsorbed protein with coupling buffer containing 0.5 M NaCl followed by acetate buffer. 7. Store column in PBS containing 1 mM sodium azide (NaH3).

3.6. Incubation Conditions and Sample Collection for Retinoid Processing and Secretion

The following procedures are performed under dim red light in the dark: 1. Wash cultured RPE cells grown on filters three times with DMEM. 2. Add 400 μl of 5 μM holo-retinol-binding protein in DMEM to the basal side (see Note 4). 3. Add 400 μl of the mixture of 10 μM holo-IRBP with 5 μM of all-trans-retinol in DMEM to the apical side (see Section 3.3).

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4. Incubate overnight at 37◦ C in 5% CO2 in incubator. 5. Collect the medium (400 μl) from the apical and basal compartments into tube containing hydroxylamine (100 mM final concentration). 3.7. Retinoid Extraction and Analysis

1. Add 200 μl of methanol, then extract the retinoids twice into 2 ml of hexane. 2. Dry the retinoid extract under a stream of argon and redissolve in 100 μl of hexane. 3. Load the hexane solutions onto a normal-phase HPLC column. 4. Analyze the contents using a gradient elution (0.2–10% dioxane in hexane at 2 ml/min) on an Agilent Zorbax silica column (see Section 2.7). 5. Confirm the retinoid identity by on-line spectral analysis and co-elution with authentic retinoid standards.

4. Notes 1. If the eyes are too small or the RPE does not separate well from the choroid, good cultures may also be obtained by using RPE-choroid explants rather than RPE alone. 2. Dry down all-trans-retinol (3 mM ethanol stock solution) under a stream of nitrogen and resuspend in 0.1% DMSO. The amount of all-trans-retinol is twofold molar excess over the total protein in the refolding buffer. 3. We also use Amicon Ultra concentrator to eliminate imidazol in the eluting buffer and change to coupling buffer. 4. We have used both recombinant RBP and commercial RBP (Sigma Chemical). However, it is more economical to prepare rather than purchase this reagent. The amount of 11-cis-retinal production by the RPE cells is about the same. Since RBP from Sigma-Aldrich contains both holo- and apoRBP, we complex it with 5 μM of all-trans-retinol before adding to the incubation.

References 1. Allikmets, R., Singh, N., Sun, H., et al. (1997) A photoreceptor cell-specific ATPbinding transporter gene (ABCR) is mutated in recessive Stargardt macular dystrophy. Nat. Genet. 15, 236–246. 2. Weng, J., Mata, N.L., Azarian, S.M., Tzekov, R.T. Birch, D.G., Travis, G.H.

(1999) Insights into the function of Rim protein in photoreceptors and etiology of Stargardt’s disease from the phenotype in abcr knockout mice. Cell 98, 13–23. 3. Marlhens, F., Bareil, D., Griffoin, J-M, et al. (1997) Mutations in RPE65 cause Leber’s

72

4.

5. 6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

Hu and Bok congenital Amaurosis. Nat. Genet. 17, 139–141. Redmond, T.M., Yu, S., Lee, E., et al. (1998) Rpe65 is necessary for production of 11-cisvitamin A in the retinal visual cycle. Nat. Genet. 20, 344–351. Allikmets, R., Dean, M. (2008) Bringing age-related macular degeneration into focus. Nat. Genet. 40, 820–821. Gehrs, K.M., Jackson, J.R., Brown, E.N., Allikmets, M., Hageman, G. (2009) Complement, age-related macular degeneration and a vision of the future. Arch. Ophthalmol. 128, 349–358. Travis, G.H., Golczak, M., Moise, A.R., Palczewski, K. (2007) Diseases caused by defects in the visual cycle: Retinoids as potential therapeutic agents. Annu. Rev. Pharmacol. Toxicol. 47, 469–512. Bok, D., Heller, J. (1976) Transport of retinol from the blood to the retina: An autoradiographic study of the pigment epithelial cell surface receptor for plasma retinol-binding protein. Exp. Eye Res. 22, 395–402. Kanai, M., Raz, A., Goodman, D.S. (1968) Retinol-binding protein: The transport protein for vitamin A in human plasma. J. Clin. Invest. 47, 2025–2044. Bok, D., Heller, J. (1980). Autoradiographic localization of serum retinol-binding protein receptors on the pigment epithelium of dystrophic rat retinas. Invest. Ophthalmol. Vis. Sci. 19, 1405–1414. Kawaguchi, R., Yu, J., Honda, J., et al. (2007) A membrane receptor for retinol binding protein mediates cellular uptake of Vitamin A. Science 315, 820–825. Golzio, C., Martinovic-Bouriel, J., Thomas, S., et al. (2007) Matthew-Wood syndrome is caused by truncating mutations in the retinol-binding protein receptor gene STRA6. Am. M. Hum. Genet. 80, 1179– 1187. Pasutto, F., Sticht, H., Hammersen, G., et al. (2007) Mutations in STRA6 cause a broad spectrum of malformations including anophthalmia, congenital heart defects, diaphragmatic hernia, alveolar capillary dysplasia, lung hypoplasia and mental retardation. Am. J. Hum. Genet. 80, 550–560. MacDonald, P.N., Ong, D.E. (1988) Evidence for a lecithin-retinol acyltransferase activity in the rat small intestine. J. Biol. Chem. 263, 12478–12482. Saari, J., Bredberg, L. (1989) Lecithin: Retinolacyltransferase in retinal pigment epithelial microsomes. J. Biol. Chem. 264, 8636–8648.

16. Ruiz, A., Winston, A., Rando, R., Bok, D. (1999) Molecular and biochemical characterization of lecithin retinol acyltransferase. J. Biol. Chem. 274, 3834–3841. 17. Bernstein, P., Law, W., Rando, R. (1987) Biochemical characterization of the retinoid isomerase4 system of the eye. J. Biol. Chem. 262, 16848–16857. 18. Deigner, P., Law, W., Canada, F., Rando, R. (1989) Membranes as the energy source in the undergone transformation of vitamin A to 11-cis-retinol. Science 244, 968–971. 19. Jin, M., Moghrabi, W.N., Sun, H., Travis, G.H. (2005) Rpe65 is the retinoid isomerase in bovine retinal pigment epithelium. Cell 122, 449–459. 20. Moiseyev, G., Chen, Y., Takahashi, Y., Wu, B.X., Ma, J.X. (2005) RPE65 is the isomerhydrolase in the retinoid visual cycle. Proc. Nat. Acad. Sci. USA 102, 12413–12418. 21. Bok, D. (1994) The retinal pigment epithelium; a versatile partner in vision. J. Cell Sci. 17(Suppl), 189–195. 22. Redmond, R.M., Poliakov, E., Yu, S., Tsai, J.Y., Lu, Z., Gentlemen, S. (2005) Mutation of key residues of ROE65 abolishes its enzymatic role as isomerohydrolase in the visual cycle. Proc. Natl. Acad. Sci. USA 102, 13658–13663. 23. Molday, L.L., Rabin, A.R., Molday, R.S. (2000) ABCR expression in foveal cone photoreceptors and its role in Stargardt macular dystrophy. Nat. Genet. 25, 67–73. 24. Eldred, G.E., Lasky, M.R. (1993) Retinal age pigments generated by self-assembling lysosomotropic detergents. Nature 361, 724–726. 25. Maguire, A.M., Simonelli, F., Pierce, E.A. et al. (2008) Safety and efficacy of gene transfer for Leber’s congenital Amaurosis. N. Engl. J. Med. 358, 2240–2248. 26. Bainbridge, J.W., Smith, A.J., Barker, S.S., et al. (2008) Effect of gene therapy on visual function in Leber’s congenital Amaurosis. N. Engl. J. Med. 358, 2231–2239. 27. Hauswirth, W.W., Aleman, T.S., Causal, S., et al. (2008) Treatment of Leber congenital amaurosis due to RPE65 mutations by ocular subretinal injection of adenoassociated virus gene vector: Short-term results of a phase I trial. Hum. Gene Ther. 19, 979–990. 28. Zhou, J., Jang, P., Kim, S.R., Sparrow, J.R. (2006) Complement activation by photooxidation products of A2E, a lipofuscin constituent of the retinal pigment epithelium. Proc. Natl. Acad. Sci. USA 103, 16182–16187.

Culture of Highly Differentiated Human Retinal Pigment Epithelium 29. Hageman, G.S., Anderson, D.H. Johnson, L.V., et al. (2005) A common haplotype in the complement regulatory gene factor H (HF1/CFH) predisposes individuals to age-related macular degeneration. Proc. Natl. Acad. Sci. USA 102, 7227–7232. 30. Haines, J.L., Hauser, M.A., Schmidt, S., et al. (2005) Complement factor H variant increases the risk of age-related macular degeneration. Science 308, 419–421. 31. Klein, R.J., Zeiss, D., Chew, E.Y., et al. (2005) Complement factor H polymorphism in age-related macular degeneration. Science 308, 385–389. 32. Edwards, A.O., Ritter, R., 3rd, Abel, K.J., Manning, A., Panhuysen, C. Farrer, L.A. (2005) Complement factor H polymorphism and age-related macular degeneration. Science 308, 421–424.

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33. Carlson, A., Bok, D. Promotion of the release of 11-cis-retinal from cultured retinal pigment epithelium by interphotoreceptor retinoid-binding protein. Biochemistry 31, 9056–9062. 34. Radu, R.A., Hu, J., Peng, J., Bok, D., Mata, N., Travis, G.H. (2008) Retinal pigment epithelium-retinal G protein receptor-opsin mediates light-dependent translocation of all-trans-retinyl esters for synthesis of visual chromophore in retinal pigment epithelial cells. J. Biol. Chem. 283, 19730–19738. 35. Maminishkis, A., Chen, S., Jalickee, S., et al. (2006) Confluent monolayers of cultured human fetal retinal pigment epithelium exhibit morphology and physiology of native tissue. Invest. Ophthalmol. Vis. Sci. 47, 3612–3624.

Chapter 3 Feeder-Independent Culture of Mouse Embryonic Stem Cells Using Vitamin A/Retinol Jaspal S. Khillan and Liguo Chen Abstract Embryonic stem (ES) cells derived from the inner cell mass of a mammalian blastocyst represent unlimited source of all types of cells for regenerative medicine and for drug discovery. Mouse and human ES cells require mouse embryonic fibroblast feeder cells to maintain their undifferentiated state which involve additional time-consuming and labor-intensive steps. Recently we reported a novel function of retinol, the alcohol form of vitamin A, in preventing the differentiation of mouse ES cells. Retinol/vitamin A induces the overexpression of Nanog, a key transcription factor that is important for maintaining the pluripotency of mouse and human ES cells. Further, retinol/vitamin A also supports feeder-independent culture of ES cells in long-term cultures. The cells continue to maintain the expression of pluripotent cellspecific markers such as Nanog, Oct4, and Sox2 and form chimeric animals after injection into blastocysts. In this chapter, we describe feeder-independent cultures of mouse ES cells in the medium supplemented with retinol. The ES cells are cultured over plates coated with gelatin in ES medium with leukemia inhibitory factor (LIF) which is supplemented with 0.5 μM retinol/vitamin A. The cells are passaged every 3–5 days by trypsinization. The pluripotency of the cells is tested by different undifferentiated ES cell-specific markers. Key words: Self-renewal of ES cells, feeder-independent cultures, vitamin A/retinol, Nanog regulation, Adh1, Adh4, RALDH2.

1. Introduction Embryonic stem (ES) cells derived from mammalian blastocysts (1, 2) have indefinite potential for self-renewal and represent a powerful source of all types of cells for regenerative medicine and drug discovery. ES cells also represent an excellent model system to study mammalian development via gene targeting by H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_3, © Springer Science+Business Media, LLC 2010

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homologous recombination followed by preparation of chimeric animals by microinjection into blastocysts. The cells maintain pluripotency through a complex interplay of different signaling pathways including LIF-Jak-Stat3, BMP 2/4, and Wnt/β-catenin pathway and intrinsic factors such as Nanog, Sox2, and Oct3/4 of which Nanog plays a crucial role in maintaining the pluripotency of ES cells (3). Although LIF plays a critical role in mouse ES cells, it is dispensable for human ES cells (4). Mouse and human ES cells maintain pluripotency when cocultured with mouse embryonic fibroblasts (MEFs) as feeders (5). The culture of ES cells with feeders involves several cumbersome and time-consuming steps such as generation of MEFs from mouse embryos, inactivation of cells by mitomycin or by γ-irradiation, and finally removal of feeder cells before using for subsequent studies. Novel recipes have been developed for feederindependent culture of human ES cells (6). It will be highly beneficial if the cells can be cultured without the feeder cells. Recently, we have demonstrated that retinol, the alcohol form of vitamin A, suppresses the differentiation of mouse ES cells by the overexpression of Nanog (7). Vitamin A/retinol also supports ES cells in the absence of feeder cells. The cells maintain undifferentiated characteristics and express pluripotent-specific genes such as Nanog, Oct4, and Sox2 in long-term cultures (8). Retinol prevents differentiation of ES cells independent of strain background such as 129Sv (Fig. 3.1), FVB/N (Fig. 3.2), and C57BL6 strains of mice (8). The cells maintained pluripotency over several passages (passage 7; Fig. 3.3 and passage 15; Fig. 3.4) and formed chimeric animals after microinjection into blastocysts (8). Retinol is usually associated with differentiation via its metabolite retinoic acid (9). Contrary to this, however, in ES cells retinol prevents differentiation and supports their self-renewal. The cells cultured in medium supplemented with retinol maintain complete potential to differentiate into all the primary germ layers in embryoid bodies and therefore provide a protocol for preparation of pure population of ES cells. Industrial-scale pure population of ES cells may be achieved in bioreactors in suspension cultures. Retinol plays an important role in a range of essential biological functions including reproduction, differentiation, immunology (10). It is stored as retinyl ester in the liver from where it is mobilized into blood plasma. In the plasma it forms a complex with 21 kDa retinol-binding protein (RBP) and thyroxine binding-protein transthyretin (TTR) for delivery to the target cells. Retinol is then transported to the cytoplasm of the target cell through RBP receptor STRA6 (stimulated by retinoic acid 6), a multitransmembrane domain protein (11) where it binds to a 15-kDa cellular retinol-binding protein (CRBP). In the cytoplasm, retinol is metabolized by two families of alcohol

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Fig. 3.1. Feeder-independent culture of R1 ES cells from 129Sv strain of mice. ES cells were cultured over gelatin-coated plates. The cells were treated with 0.5 μM retinol for 5 days followed by staining for alkaline phosphatase. (A) Normal ES cells cultured in ES medium without LIF. The cells are completely differentiated as noticed by the complete absence of staining for alkaline phosphates (AP). (B) The cells cultured in the medium with LIF. Although some cells stain positive for AP, overall the colonies exhibit flat morphology of differentiating cells. (C) ES cells cultured in the medium containing 0.5 μM retinol in the absence of LIF. Most of the cells exhibit undifferentiated morphology and stained strongly positive for AP. Only a few cells in the center were flat cells. (D) ES cells cultured in medium with LIF and 0.5 μM retinol. Almost all the colonies exhibit undifferentiated morphology with strong staining for AP (40× magnification).

Fig. 3.2. Feeder-independent culture of ES cells from FVB/N strain of mice. ES cells were cultured over gelatin-coated plates. The cells were treated with 0.5 μM retinol for 5 days followed by staining for alkaline phosphatase. (A) Normal ES cells cultured in ES medium supplemented with LIF. The cells are completely differentiated as noticed by the complete absence of staining for AP. (B) ES cells cultured in medium with LIF and 0.5 μM retinol. Almost all the colonies exhibit undifferentiated morphology with strong staining for AP (20× magnification).

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Fig. 3.3. Retinol-treated ES cells at passage 7. ES cells were cultured in ES medium supplemented with 0.5 μM retinol on gelatin-coated plates without feeders. The cells were passaged every 4–5 days to fresh plates coated with gelatin (20× magnification).

Fig. 3.4. Retinol-treated ES cells at passage 15. ES cells were cultured in ES medium supplemented with 0.5 μM retinol on gelatin-coated plates without feeders. The cells were passaged every 4–5 days to fresh plates coated with gelatin. Almost all the colonies maintained undifferentiated morphology (20× magnification).

dehydrogenases (Adh1 and Ad4) into retinaldehyde which is then converted into retinoic acid by retinaldehyde dehydrogenase 2 (RALDH2) that binds to specific cellular retinoic acid-binding proteins (CRABP1 and CRABP2) that shuttle retinoic acid to the nucleus. In the nucleus, retinoic acid binds to specific retinoic acid receptors (RAR) and retinoic X receptors (RXR) to bind to the promoter elements of retinoic acid-responsive genes (RARE) to activate transcription of >500 genes (12). Interestingly, our studies have shown that ES cells do not express Adh1, Adh4, and RALDH2 that metabolize retinol into retinoic acid. The vitamin A/retinol-mediated ES cells

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self-renewal, therefore, is independent of retinoic acid (7, 8). In this chapter, we describe the protocol for feeder-independent culture mouse ES cells using vitamin A/retinol as supplement.

2. Materials 2.1. Plasticware and Chemicals

1. Tissue culture plates 35, 60, and 100 mm. 2. 1, 5, and 10 ml pipettes. 3. 0.1% gelatin solution. 4. Retinol from Sigma-Aldrich (St. Louis, MO). A 100 μM stock solution is prepared in 100% ethanol (see Note 2).

2.2. Culture and Propagation of ES Cells

1. Dulbecco’s modified Eagle’s medium (DMEM) (Invitrogen). 2. Fetal bovine serum, L-glutamine, non-essential amino acids, β-mercaptoethanol, trypsin-EDTA 0.25%.100X Pen-strep (all purchased from Invitrogen Corporation). 3. Leukemia inhibitory factor (LIF) (from Chemicon International).

2.3. Preparation of ES Culture Medium

1. ES medium is prepared using Dulbecco’s modified Eagle’s medium (DMEM) (5) with 15% fetal bovine serum, 1 mM L-glutamine, 1% non-essential amino acids, 0.1 mM β-mercaptoethanol, and Pen-Strep solution (see Note 1). 2. The medium is supplemented with 1000 IU of LIF. 3. The cells are cultured on appropriate size of Petri dish in incubators at 37◦ C in a humid atmosphere containing 5% CO2 .

2.4. Alkaline Phosphatase Detection of Cells

1. Alkaline phosphatase staining kit from Chemicon International (Cat#SCR004). 2. 4% paraformaldehyde. 3. 1X phosphate-buffered saline.

2.5. Isolation of RNA

1. 100 mm dishes with ES cells as described earlier. 2. STAT 60 solution (TEL-TEST, Friendswood, TX).

2.6. Western Blot Analysis

1. Anti-β-actin, anti-Oct4, and anti-Sox2 antibodies are purchased from Santa Cruz Biotechnology (Santa Cruz, CA). 2. Nanog antibody from Chemicon International. 3. Protein extraction RIPA buffer (Sigma Cat. #R0278).

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2.7. RT-PCR Primer Sequences

HPRT F 5 -GTAATGATCAGTCAACGGGGGAC-3 , R 5 -CCAGCAAGCTTGCAACCTTAACCA-3 , annealing temperature 55◦ C; Oct3/4 F 5 -GGCGTTCTCTTTGGAAAGGTGTTC-3 , R 5 -CTCGAACCACATCCTTCTCT-3 , annealing temperature 55◦ C; Stat3 F 5 -ATGAAGAGTGCCTTCGTGGTGG-3 , R 5 -GGATTGATGCCCAAGCATTTGG-3 , annealing temperature 55◦ C; Nanog F 5 -AGGGTCTGCTACTGAGATGCTCTG-3 , R 5 -CAACCACTGGTTTTTCTGCCACCG-3 , annealing temperature 55◦ C; Rex1 F5 -ATCCGGGATGAAAGTGAGATTAGC-3 , R 5 -∗ CTTCAGCATTTCTTCCCTGCCTTTGC-3 , annealing temperature 61◦ C; Sox2 F 5 -GAGAGCAAGTACTGGCAAGACCG-3 , R 5 -TATACATGGATTCTCGCCAGCC-3 , annealing temperature 64◦ C.

3. Methods 3.1. Culture and Propagation of Embryonic Stem Cells 3.1.1. Culturing of ES Cells

1. Prepare gelatin-coated plates by treating with 0.1% gelatin for 1–2 h. 2. Aspirate gelatin solution and transfer ES cells onto the plates (1 × 103 cells/cm2 ). 3. Add ES medium and allow cells to settle for 10–12 h. 4. Replace ES medium with fresh medium supplemented with 0.5 μM retinol (stock solution 100 μM dissolved in 100% ethanol as 2 μl/ml of medium, see Note 2). 5. Replace medium everyday adding fresh retinol solution taking care not to disturb ES colonies (see Note 3).

3.1.2. Propagation of Cells

1. Aspirate medium taking care not to suck off the cells (Note 3). 2. Wash cells two times with 1X PBS and add 0.25% trypsinEDTA (1 ml/60 mm, 2 ml/100 mm plate). 3. Transfer cells into CO2 incubator for 4 min followed by addition of 5 ml ES medium.

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4. Break colonies into lumps of 5–10 cells. Do not break colonies to single-cell suspension (see Note 4). 5. Add fresh medium supplemented with 0.5 μM retinol and change medium everyday using fresh retinol each time. 6. Monitor colonies everyday and as soon as the colonies are about 300–500 cells, the culture should be trypsinized. 7. Usually it takes about 4–5 days for the cells to become confluent. 8. Do not allow colonies to grow very large, otherwise the cells may differentiate. 9. More than 90% of the colonies maintain sharp phase bright morphology and are undifferentiated. 10. Duplicate plates may be created for staining for alkaline phosphatase to check undifferentiated cells. Alkaline phosphatase is a marker for undifferentiated ES cells. 11. The cells should be frozen at every passage in 10% DMSO and 90% FBS for future usage. 3.2. Alkaline Phosphatase Assay

1. Transfer approximately 2.0 × 104 cells over six-well tissue culture plates coated with gelatin in ES medium. 2. Replace with ES medium containing 0.5 μM retinol after 10–14 h. 3. Allow cells to grow 3–5 days to form colonies with the change of medium everyday using fresh retinol. 4. Aspirate medium carefully without disturbing the colonies and also add fresh medium so as not to dislodge the colonies. 5. Fix cells with 4% paraformaldehyde for 2 min at room temperature. 6. Stain cells for alkaline phosphatase using a kit from Chemicon (Temecula, CA) following protocols provided by the manufacturer.

3.3. RT-PCR Analysis

1. Culture approximately 1.0 × 106 cells over 60 mm tissue culture plates coated with gelatin using ES medium. 2. Replace medium with ES medium containing 0.5 μM retinol after 10–14 h. 3. Allow cells to grow 3–5 days with the change of medium everyday using fresh retinol. 4. Aspirate medium carefully and add 2 ml STAT 60 solution (TEL-TEST, Friendswood, TX) and pass the cells through pipette several times. 5. Store cells at room temperature for 5 min and add 0.2 ml chloroform/ml of STAT-60; shake vigorously and centrifuge at 12,000×g for 15 min at 4◦ C.

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6. Collect the aqueous phase and add 0.5 ml isopropanol/ml of STAT-60 used for cell lysis. 7. Spin down for 10 min at 12,000×g for 10 min to pellet total RNA. 8. Wash RNA with 75% ethanol and dissolve in RNase-free water. 9. Total RNA is converted into cDNA using oligo-dT and avian myeloblastosis virus (AMV) reverse transcriptase using kit purchased from Invitrogen (Carlsbad, CA). 10. Run RT-PCR in a total reaction volume of 50 μl using specific primers using PCR conditions; denaturation at 94◦ C for 45 s; extension at 72◦ C 2 for min; annealing at temperature as specified for each primer pair for 30 cycles. 11. The PCR products are resolved by agarose gel electrophoresis. 12. Visualize the amplified DNA by ethidium bromide staining using HPRT primers as control. 3.4. Western Blot Analysis

1. Culture ES cells as explained in Section 3.3. 2. Extract total protein with RIPA buffer (Sigma Cat. #R0278). 3. Load 50 μg of the protein onto 12% SDS-PAGE followed by transfer onto nylon membrane (Bio-Rad). 4. The membranes are incubated with antibodies to specific protein followed by incubation with HRP-conjugated goat antibody to mouse IgG or rabbit antibody to goat IgG (Santa Cruz Biotechnology). 5. The membranes are developed with chemiluminescence reagent (Pierce, Rockford, IL) and exposed to X-ray film.

4. Notes 1. ES cells are sensitive to different lots of serum. Only the ES cell-tested serum for optimal cell growth should be used. Usually several batches of serum are tested to select a batch that shows robust growth of cells and no toxicity at higher concentrations such as 30%. Alternatively, ES celltested serum can be purchased directly from the commercial suppliers. 2. All-trans-retinol is sensitive to light; therefore, all the preparations of retinol solution and the operations with retinol solution must be carried out in the dark. A 100 μM stock solution is prepared in 100% ethanol and the stock solution can be stored in Eppendorf tubes covered with aluminum

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foil at −80◦ C. The stock solution is diluted 1:40 with 100% ethanol and stored at −20◦ C wrapped in aluminum foil. To prepare medium with 0.5 μM retinol, 2 μl 1:40 dilution is added to 1 ml of ES medium. 3. The ES cells cultured over gelatin-coated plates are less firmly attached as compared to cells cultured over feeder cells; therefore, utmost care should be taken to prevent accidental dislodging of the colonies. The medium should be aspirated by tilting the plates on one side using low suction. Caution must be observed not to aspirate the medium completely to prevent drying of the cells. Similar caution must be observed while adding the fresh medium. The medium should be delivered very slowly from the sides of the Petri dish. 4. The cells should be trypsinized to break colonies to lumps of 5–10 cells and not to disperse as single cells. The individual cells take longer to form colonies and the survival of cells is also compromised. The cells should be trypsinized as soon as the colonies grow to about 300–500 cells. Although vitamin A/retinol-treated colonies exhibit undifferentiated morphology for longer periods, the cells must be trypsinized every 4–5 days. References 1. Martin, G. (1981) Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc. Natl. Acad. Sci. USA 78, 7634–7638. 2. Evans, M.J., Kaufman, M.H. (1981) Establishment in culture of pluripotential cells from mouse embryos. Nature 292, 154–156. 3. Boiani, M., Scholer, H.R. (2005) Regulatory networks in embryo-derived pluripotent stem cells. Nat. Rev. Mol. Cell. Biol. 6, 872–881. 4. Dahéron, L., Opitz, S.L., Zaehres, H., Lensch, M.W., Andrews, P.W., ItskovitzEldor, J., Daley, G.Q. (2004) LIF/STAT3 signaling fails to maintain self-renewal of human embryonic stem cells. Stem Cells 22, 770–778. 5. Robertson, E.J. (1987) Embryo derived cell lines. In: Teratocarcinoma and Embryonic Stem Cells: A Practical Approach, IRL Press, Oxford, pp. 71–112. 6. Ludwig, T., Thomson, J.A. (2007) Defined, feeder-independent medium for human embryonic stem cell culture. Curr. Protoc. Stem Cell Biol. Chapter 1, Unit 1C.2. 7. Chen, LG., Yang, M., Dawes, J., Khillan, J.S. (2007) Suppression of ES cell differentiation

8.

9.

10.

11.

12.

by retinol (vitamin A) via the over expression of Nanog. Differentiation 75, 682–693. Chen, L., Khillan, J.S. (2008) Promotion of feeder independent self-renewal of embryonic stem cells by retinol (vitamin A). Stem Cells 26, 1858–1864. Clagett-Dame, M., De Luca, H.F. (2002) The role of vitamin A in mammalian reproduction and embryonic development. Annu. Rev. Nutr. 22, 347–381. Mark, M., Ghyselinck, N.B., Chambon, P. (2006) Function of retinoid nuclear receptors: Lessons from genetic and pharmacological dissections of the retinoic acid signaling pathway during mouse embryogenesis. Annu. Rev. Pharmacol. Toxicol. 46, 451–480. Kawaguchi, R., Yu, J., Honda, J., Hu, J., Whitelegge, J., Ping, P., et al. (2007) Membrane receptor for retinol binding protein mediates cellular uptake of vitamin A. Science 315, 820–825. Lane, M.A., Xu, J., Wilen, E.W., Sylvester, R., Derguini, F., Gudas, L.J. (2008) LIF removal increases CRABPI and CRABPII transcripts in embryonic stem cells cultured in retinol or 4-oxoretinol. Mol. Cell Endocrinol. 280, 63–67.

Chapter 4 In Vitro Assays of Rod and Cone Opsin Activity: Retinoid Analogs as Agonists and Inverse Agonists Masahiro Kono and Rosalie K. Crouch Abstract Upon absorption of a photon, the bound 11-cis-retinoid isomerizes to the all-trans form resulting in a protein conformational change that enables it to activate its G protein, transducin, to begin the visual signal transduction cascade. The native ligand, 11-cis-retinal, acts as an inverse agonist to both the apoproteins of rod and cone visual pigments (opsins); all-trans-retinal is an agonist. Truncated analogs of retinal have been used to characterize structure–function relationships with rod opsins, but little has been done with cone opsins. Activation of transducin by an opsin is one method to characterize the conformational state of the opsin. This chapter describes an in vitro transducin activation assay that can be used with cone opsins to determine the degree to which different ligands can act as an agonist or an inverse agonist to gain insight into the ligand-binding pocket of cone opsins and differences between the different classes of opsins. The understanding of the effects of ligands on cone opsin activity can potentially be applied to future therapeutic agents targeting opsins. Key words: Retinal analog, cone opsin, G protein-coupled receptor, transducin, cone pigment, rhodopsin.

1. Introduction There are two types of photoreceptors in vertebrate retina – rods and cones. They have distinct physiological roles, the rods operating under dim light conditions and being exquisitely sensitive and the cones requiring more light and discerning colors. The cones are required for normal human vision. The light-detecting components in the photoreceptor cells are visual pigments. Visual pigments are comprised of proteins (opsins) and chromophore (11-cis-isomer of vitamin A aldehyde (retinal) or 3,4-dehydroretinal). Because there is no variation in the H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_4, © Springer Science+Business Media, LLC 2010

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chromophore, the ability to detect light across the color spectrum depends on the influence of the different opsins on the absorption properties of the chromophore. Retinal analogs have been useful in the past to probe spectral tuning and the binding site restraints of visual pigments (1). The protein moieties of rod and cone pigments (opsins) are highly homologous to each other and also belong to the superfamily of G protein-coupled receptors (GPCRs). The G protein activated by these visual pigments in initiating the visual signal transduction cascade is transducin. An opsin is referred to as being active when it is able to activate this G protein. Unlike other GPCRs, the ligand of visual pigments is covalently bound to a strictly conserved lysine in the seventh transmembrane helix of the opsins through a Schiff base linkage. 11-cis-Retinal (or the 3,4-dehydro form) acts as an inverse agonist with all the vertebrate visual opsins tested, maintaining the receptor in an inactive state. On absorption of a photon, bound 11-cis-retinal isomerizes to the more stable all-trans form and the protein receptor is transformed into an active conformation. Thus, it is the light that converts the inverse agonist into an agonist via photoisomerization. In the eye, the Schiff base between the all-trans-retinal and the protein is subsequently hydrolyzed and the retinal is reduced to all-trans-retinol leaving the opsin as the apoprotein. The opsins themselves are weakly constitutively active (2–6) and all regenerate in the presence of 11-cis-retinal, reforming the inactive, photosensitive pigments. Although the native ligand is covalently bound to the opsins in the inactive and photoactivated states, a covalently attached ligand is not absolutely required to deactivate or activate the opsins. Several truncated analogs of retinal have been demonstrated to activate the rod opsin (7–9). Furthermore, a highly constitutively active rhodopsin mutant where the conserved lysine that normally forms the Schiff base with the chromophore has been mutated has been shown to be deactivated and made light-sensitive with the 11-cis-retinyl Schiff base, where 11-cis-retinal has been coupled to n-propylamine (10). To date, there has been a dearth of information of liganddependent activation and deactivation of cone opsins. A major reason for this is the lack of methods and sources for pure cone opsins; whereas, rod opsins in good purity are easily isolated. Another reason is the perceived instability of the protein (11). Cone pigments have been shown to lose its chromophore or, in the presence of analogs, exchange chromophores in the dark (12– 14) unlike the rod opsin where the pigment (rhodopsin) is quite stable even to hydroxylamine. Heterologously expressed opsins are a convenient tool for probing opsin–ligand interactions as there is no question of retinoid photoproducts remaining attached to the membranes,

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there is no mixture of different opsins, and mutants can be readily constructed and tested. We have shown that different ligands can affect the cone opsins and pigment activation differently depending on opsin type and ligand (4, 15, 16). All-trans-retinal and alltrans-retinol are both agonists with all four opsin groups (unpublished results, 2009). The other physiologically relevant ligand is 11-cis-retinol. Surprisingly, this ligand has quite different activities with the various opsins which have important physiological consequences (15, 17, 18). β-Ionone, representing a fragment of retinal, has been shown to be both an inverse agonist and agonist depending on cone opsin type (4), illustrating that cone opsins do not all interact with ligands in the same manner. In Fig. 4.1, we illustrate the modulation of transducin activation by expressed human red cone opsin as a function of retinal

Fig. 4.1. Transducin activation by expressed human red cone opsin. (a) Time-dependent activation of transducin by human red cone opsin without (open circles) and with (triangles) 11-cis-retinal. At 5.5 min, the opsin with 11-cis-retinal was exposed to >530 nm light for 12 s demonstrating that pigment had formed and light-dependent activation occurred due to photoisomerization of the bound 11-cis form of the chromophore to the all-trans form. Note the reduction in transduction activation after 11-cis-retinal was added. The pH of the reaction was 6.4. (b) Relative transducin activation by expressed human red cone opsin after addition of 200 μM retinal analogs [AT-RAL (all-trans-retinal); C17-RAL (17 carbon all-trans-retinal analog); and β-ionone]. Activity was normalized to the activation by opsin alone.

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analog length. This methodology provides a convenient in vitro tool for studying the interactions of opsins with various compounds that are potential ligands for these opsins. For the human red cone opsin, as the polyene chain decreases in length, the ligand converts from an agonist activating the G protein to an inverse agonist, decreasing the opsin’s ability to activate this G protein (Fig. 4.1b). We describe here an in vitro assay for determining the ability of a retinal analog to act as an agonist or an inverse agonist with various opsins. The use of this assay can serve to provide insight into structural and functional similarities and differences among cone opsins.

2. Materials

2.1. Material Sources

2.2. Stock Solutions

GTPγS-35: catalog number NEG030H250UC, PerkinElmer Life and Analytical Sciences, Waltham, MA; 1D4 antibody – available through a number of vendors including catalog number MA1-722 from Affinity BioReagents/Thermo Fisher Scientific, Rockford, IL. 1. Membrane prep buffer: 150 mM NaCl, 1 mM MgCl2 , 1 mM CaCl2 , 0.1 mM EDTA, 10 mM Tris–HCl (pH 7.4). The opsin concentration is typically 10–50 nM (see Note 1). 2. Transducin buffer (2×): 20 mM Tris (pH 7.4), 4 mM MgCl2 , and 2 mM DTT. 3. Assay buffer (10×): 100 mM MES buffer, 1 M NaCl, 50 mM MgCl2 at pH 6.5. 4. DTT solution: 50 mM in Milli-Q water. 5. Analog solution: 20 mM in ethanol (see Note 2). 6. GTPγS solution: 150 μM cold GTPγS with GTPγS35 at ∼0.25 mCi/ml; 100 μl of a 150 μM solution of GTPγS from ∼3 mM stock solution and add 2 μl (25 μCi) GTPγS-35. 7. Assay rinse buffer: 10 mM Tris, pH 6.4, 100 mM NaCl, 5 mM MgCl2 .

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3. Methods

3.1. Membrane Preparation Containing Opsins

1. Transiently expressed cone opsins in COS cells (19) with opsin genes with the codons for at least the last eight amino acid residues of bovine rhodopsin, the 1D4 epitope (see Note 3). 2. COS cell membranes containing the opsins isolated using a discontinuous sucrose gradient (5, 20, 21). 3. Membrane suspensions of 25 μM aliquots in membrane prep buffer stored at −80◦ C. 4. The amount of opsin in the membrane preparations are determined by slot blot analysis (20) using known amounts of bovine rhodopsin as reference and probed with the rhodopsin 1D4 antibody.

3.2. Transducin Preparation

1. Transducin purified from bovine retinae (W.L. Lawson, Lincoln, NE) (22–24) (see Note 4). 2. This sample is then applied to a 3 ml DEAE-cellulose anion exchange column (22), which is washed with 10 column volumes of 1× transducin buffer and then 20 column volumes of the same buffer with 100 mM NaCl. 3. Transducin is eluted with the transducin buffer containing 500 mM NaCl and fractions monitored by absorbance at 280 nm. Pooled fractions containing transducin are dialyzed three times against a 1:1 mixture of glycerol and 2× transducin buffer, diluted to 50 μM and stored at −20◦ C.

3.3. Activity Assay

The ability of the specific opsin to activate bovine rod transducin is determined using a radioactive filter-binding assay with membrane preparations of opsin expressed in COS cells essentially as described previously with a few modifications (5, 21) (see Note 5). 1. Wet filter membranes with water in a tray; place filters onto the vacuum manifold; assemble the manifold. We use a Millipore 1225 sampling vacuum manifold (Millipore, Billerica, MA) with 25 mm diameter Millipore mixed cellulose ester membranes (HAWP 02500; Millipore, Billerica, MA) attached. 2. Prepare the reaction mixture without retinoid and GTPγS in a 1.5 ml microcentrifuge tube:

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Volume (µl) 70

Deionized water

10

10× assay buffer (see Note 6)

2

50 mM DTT

5

Transducin (50 μM stock)

10

Opsin/visual pigment (typically, 10–50 nM stock concentration) (see Note 1)

3. Add 1 μl retinal/retinal analog/ethanol (for opsin control). 4. Start reaction by adding 2 μl of 150 μM GTPγS solution and start the clock. 5. At each time point, remove 10 μl aliquots and pipet onto filter. Wash filters three times with 4 ml rinse buffer with a repeating pipettor. 6. Continue with each time point (usually 1 min intervals). 7. Transfer filter membranes into scintillation vials. 8. Add 10 μl of reaction mixture directly into scintillation vials. These counts will be used to convert counts per minute (cpm) to GTPγS amounts in pmol because these scintillation vials contain 30 pmol GTPγS since none was washed away. 9. Add 10 ml Amersham BCS scintillation cocktail (catalog number: NBCS104, GE Healthcare, Piscataway, NJ). 10. The vials are shaken for at least 1 h and often overnight for convenience and measured in a scintillation counter (usually 1–5 min counts). The counts per minute can be converted to pmol GTPγS bound, which reflects the amount of transducin being activated with time (see Notes 5, 7, and 8). 11. Other visual pigment protein preparations can be used (see Notes 9–11), and if a light-sensitive pigment is generated, dark/light differences can be determined (see Note 12).

4. Notes 1. Opsin concentrations should be kept as low as possible, typically nanomolar range, to allow for multiple turnovers. 2. As a starting point to quickly assay a ligand, we have been using 200 μM of the ligand (20 mM stock solution in

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ethanol, if possible). For most ligands we have tested, this is more than sufficient. However, for completeness, the ligand concentration dependence ought to be determined. 3. We express opsins in COS cells. Other cells such as HEK293 (25, 26) and Sf 9 (27) cells have been used to successfully express rod and cone opsins and can certainly be used for these assays. We prefer to use COS cells because of our experience with them and relative low cost of maintenance and transfection. The DEAE-dextran transient transfection method is quite harsh but tolerated by confluent COS cells, and the reagents are relatively inexpensive and readily prepared in the lab rather than purchased from a kit. Furthermore, we passage the cells with media supplemented with bovine serum (19) rather than fetal bovine serum, which results in considerable cost savings. 4. This is essentially a protocol for rod outer segments prepared in the light using bleached rhodopsin to anchor the transducin to the membrane and releasing transducin from the membrane with GTP. 5. The assay is based on the finding that as the receptor (opsin) activates the G protein (transducin), a bound GDP is released and free GTPγS binds to the G protein. Proteins including transducin and transducin bound with radioactive GTPγS adhere to the filter membrane, and unbound GTPγS flows through with the wash buffer. In this manner, the rate at which GTPγS is taken up by the G protein can be determined by plotting cpm per unit time (or more appropriately picomole-bound GTP per unit time). The cpm can be converted to mol GTPγS because the amount of GTPγS in the scintillation vial(s) from step 8 is 30 pmol. 6. The constitutive activity of the opsins is measured at an acidic pH (the final pH is 6.4 in our assays, but the 10× stock buffer is made at pH 6.5), which enhances the activation by the apoprotein such that the lower activity in the presence of inverse agonist such as 11-cis-retinal is clearly distinguishable (2, 4, 5). 7. Kinetics are generally linear as we assume pseudo-firstorder kinetics. This requires the substrates to be in excess and opsin to be limiting. Deviations from linearity can occur if the photoactive intermediate is decaying rapidly compared to the timescale of the assay such as with cone pigments (16) or if other substrates are being depleted. 8. We generally report our activities as a mean of three or more measurements ± standard error of the mean. 9. We have described methods for purified membranes of opsins transiently expressed in COS cells. However, opsins

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can be purified using the 1D4 antibody coupled to Sepharose 4B to immunopurify opsins solubilized and purified with CHAPS and lipid. For example, rhodopsin mutants have been purified with CHAPS/asolectin (28). Because of the high critical micelle concentration of CHAPS, this detergent is easily removed by a number of methods to leave behind opsins in asolectin vesicles. 10. If the analog to be tested results in generation of a stable pigment, then the pigments can be detergent solubilized and immunopurified allowing for spectroscopic and activity measurements on the same samples. Such was done with 9-demethylretinal and salamander rod and cone opsins (16). In these situations, the detergent choice and concentration can greatly affect transducin activation assays. CHAPS is not a good detergent for transducin activation assays. Dodecylmaltoside, if the final concentration is kept at or below 0.1%, is a suitable detergent (29). 11. Native opsins can be measured. Rod outer segments are the most easily obtained from a sucrose float. However, cone pigments from cone-dominant retinae have been purified from native sources such as chicken (30) and geckos (31) and could be used. However, opsins from native sources ideally need to have the native chromophore removed to ensure that effects are due to retinal analogs and not the native chromophore. 12. If rod and/or cone pigments are to be assayed for lightdependent activation, then light conditions must be considered. If membrane preparations are used with an excess of chromophore, the assays should be conducted in the dark (dim red light conditions) and a pulse of light used to bleach the pigment but the assay continued in the dark. Continuous light can result in photoreactions of the light-activated product and photoactivation of new pigment regenerated after hydrolysis of the chromophore. The latter is especially a concern with cone pigments as their chromophore is released in the timescale of seconds, whereas the release is several minutes with rhodopsin (16). We bleach our samples with a slide projector containing a 300-W bulb with a longpass filter attached to minimize bleaching the active intermediate. The main consideration for the filter is to overlap with the absorption spectrum of the pigment band and to minimize light hitting the nearUV spectrum. Thus the type of optical filter depends on the absorption spectrum of the pigment of interest. While there are different sources and types of optical filters available, we have purchased a number of longpass filters from Edmund Optics (Barrington, NJ) as they are quite inexpensive and

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available in a variety of colors and sizes. For example, the 2-in. square OG-530 longpass glass filter is convenient for rhodopsin and green and red cone pigments. Appropriate band-pass filters can also be used.

References 1. Lou, J., Tan, Q., Karnaukhova, E., Berova, N., Nakanishi, K., Crouch, R.K. (2000) Synthetic retinals: Convenient probes of rhodopsin and visual transduction process. Methods Enzymol. 315, 219–237. 2. Cohen, G.B., Yang, T., Robinson, P.R., Oprian, D.D. (1993) Constitutive activation of opsin: Influence of charge at position 134 and size at position 296. Biochemistry 32, 6111–6115. 3. Surya, A., Foster, K.W., Knox, B.E. (1995) Transducin activation by the bovine opsin apoprotein. J. Biol. Chem. 270, 5024–5031. 4. Isayama, T., Chen, Y., Kono, M., DeGrip, W.J., Ma, J.-X., Crouch, R.K., Makino, C.L. (2006) Differences in the pharmacological activation of visual opsins. Vis. Neurosci. 23, 899–908. 5. Kono, M. (2006) Constitutive activity of a UV cone opsin. FEBS Lett. 580, 229–232. 6. Melia, T.J., Jr., Cowan, C.W., Angleson, J.K., Wensel, T.G. (1997) A comparison of the efficiency of G protein activation by ligandfree and light-activated forms of rhodopsin. Biophys. J. 73, 3182–3191. 7. Bartl, F.J., Fritze, O., Ritter, E., Herrmann, R., Kuksa, V., Palczewski, K., Hofmann, K.P., Ernst, O.P. (2005) Partial agonism in a G protein-coupled receptor. Role of the retinal ring structure in rhodopsin activation. J. Biol. Chem. 280, 34259–34267. 8. Han, M., Groesbeek, M., Sakmar, T.P., Smith, S.O. (1997) The C9 methyl group of retinal interacts with glycine-121 in rhodopsin. Proc. Natl. Acad. Sci. USA 94, 13442–13447. 9. Buczylko, J., Saari, J.C., Crouch, R.K., Palczewski, K. (1996) Mechanisms of opsin activation. J. Biol. Chem. 271, 20621–20630. 10. Zhukovsky, E.A., Robinson, P.R., Oprian, D.D. (1991) Transducin activation by rhodopsin without a covalent bond to the 11-cis-retinal chromophore. Science 251, 558–560. 11. Ramon, E., Mao, X., Ridge, K.D. (2009) Studies on the stability of the human cone visual pigments. Photochem. Photobiol. 85, 509–516.

12. Crescitelli, F. (1984) The gecko visual pigment: The dark exchange of chromophore. Vision Res. 24, 1551–1553. 13. Kefalov, V.J., Estevez, M.E., Kono, M., Goletz, P.W., Crouch, R.K., Cornwall, M.C., Yau, K.-W. (2005) Breaking the covalent bond – a pigment property that contributes to desensitization in cones. Neuron 46, 879–890. 14. Matsumoto, H., Tokunaga, F., Yoshizawa, T. (1975) Accessibility of the iodopsin chromophore. Biochim. Biochem. Acta 404, 300–308. 15. Ala-Laurila, P., Cornwall, M.C., Crouch, R.K., Kono, M. (2009) The action of 11cis-retinol on cone opsins and intact cone photoreceptors. J. Biol. Chem. 284, 16492– 16500. 16. Das, J., Crouch, R.K., Ma, J.-X., Oprian, D.D., Kono, M. (2004) Role of the 9-methyl group of retinal in cone visual pigments. Biochemistry 43, 5532–5538. 17. Jones, G.J., Crouch, R.K., Wiggert, B., Cornwall, M.C., Chader, G.J. (1989) Retinoid requirements for recovery of sensitivity after visual-pigment bleaching in isolated photoreceptors. Proc. Natl. Acad. Sci. USA 86, 9606–9610. 18. Kono, M., Goletz, P.W., Crouch, R.K. (2008) 11-cis and all-trans retinols can activate rod opsin: Rational design of the visual cycle. Biochemistry 47, 7567–7571. 19. Oprian, D.D. (1993) Expression of opsin genes in COS cells. Methods Neurosci. 15, 301–306. 20. Kono, M., Crouch, R.K., Oprian, D.D. (2005) A dark and constitutively active mutant of the tiger salamander UV pigment. Biochemistry 44, 799–804. 21. Robinson, P.R. (2000) Assays for the detection of constitutively active opsins. Methods Enzymol. 315, 207–218. 22. Baehr, W., Morita, E.A., Swanson, R.J., Applebury, M.L. (1982) Characterization of bovine rod outer segment G-protein. J. Biol. Chem. 257, 6452–6460. 23. Wessling-Resnick, M., Johnson, G.L. (1987) Allosteric behavior in transducin activation

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24.

25.

26.

27.

Kono and Crouch mediated by rhodopsin. J. Biol. Chem. 262, 3697–3705. Yu, H., Kono, M., McKee, T.D., Oprian, D.D. (1995) A general method for mapping tertiary contacts between amino acid residues in membrane-embedded proteins. Biochemistry 34, 14963–14969. Nathans, J. (1990) Determinants of visual pigment absorbance: Identification of the retinylidene Schiff’s base counterion in bovine rhodopsin. Biochemistry 29, 9746–9752. Tsutsui, K., Imai, H., Shichida, Y. (2007) Photoisomerization efficiency in UVabsorbing visual pigments: Protein-directed isomerization of an unprotonated retinal Schiff base. Biochemistry 46, 6437–6445. Vissers, P.M.A.M., Bovee-Geurts, P.H.M., Portier, M.D., Klaassen, C.H.W., DeGrip, W.J. (1998) Large-scale production and purification of the human green

28.

29.

30.

31.

cone pigment: Characterization of late photo-intermediates. Biochem. J. 330, 1201–1208. Rim, J., Oprian, D.D. (1995) Constitutive activation of opsin: Interaction of mutants with rhodopsin kinase and arrestin. Biochemistry 34, 11938–11945. Han, M., Groesbeek, M., Smith, S.O., Sakmar, T.P. (1998) Role of the C9 methyl group in rhodopsin activation: Characterization of mutant opsins with the artificial chromophore 11-cis-9-demethylretinal. Biochemistry 37, 538–545. Okano, T., Fukada, Y., Artamonov, I.D., Yoshizawa, T. (1989) Purification of cone visual pigments from chicken retina. Biochemistry 28, 8848–8856. Liang, J., Govindjee, R., Ebrey, T.G. (1993) Metarhodopsin intermediates of the gecko cone pigment P521. Biochemistry 32, 14187–14193.

Chapter 5 Physiological Studies of the Interaction Between Opsin and Chromophore in Rod and Cone Visual Pigments Vladimir J. Kefalov, M. Carter Cornwall, and Gordon L. Fain Abstract The visual pigment in vertebrate photoreceptors is a G protein-coupled receptor that consists of a protein, opsin, covalently attached to a chromophore, 11-cis-retinal. Activation of the visual pigment by light triggers a transduction cascade that produces experimentally measurable electrical responses in photoreceptors. The interactions between opsin and chromophore can be investigated with electrophysiologial recordings in intact amphibian and mouse rod and cone photoreceptor cells. Here we describe methods for substituting the native chromophore with various chromophore analogs to investigate how specific parts of the chromophore affect the signaling properties of the visual pigment and the function of photoreceptors. We also describe methods for genetically substituting the native rod opsin gene with cone opsins or with mutant rod opsins to investigate and compare their signaling properties. These methods are useful not only for understanding the relation between the properties of visual pigments and the function of photoreceptors but also for understanding the mechanisms by which mutations in rod opsin produce night blindness and other visual disorders. Key words: Opsin, chromophore, visual pigment, photoreceptor, phototransducion, dark adaptation, transgenic pigment, rhodopsin mutation.

1. Introduction Visual pigments are photon-absorbing molecules that enable rod and cone photoreceptors to produce electrical signals in response to light. They consist of a protein called opsin and a chromophore, in vertebrates usually 11-cis-retinal. In contrast to other G protein-coupled receptors, the ligand (retinal) in the visual pigment is covalently attached to the protein and functions both as reverse agonist in the dark (11-cis-configuration) and as an H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_5, © Springer Science+Business Media, LLC 2010

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agonist upon absorption of a photon (all-trans-configuration). The relative ease of delivering light stimuli of known strength and duration allows the detailed characterization of the function of visual pigments in intact photoreceptors. As a result, it is possible to use physiological measurements from single rod and cone photoreceptors to investigate the interaction between opsin and chromophore, as well as the relation between the properties of visual pigments and the function of photoreceptors. There are two complementary approaches to the study of visual pigments under physiological conditions. The first involves the modification of the chromophore by substituting the native form with a retinoid analog. The second is the transgenic expression of exogenous opsin in place of, or in addition to, the native opsin. The first approach is based on the light-induced decay of the photoactivated (bleached) visual pigment. Following photon absorption, the activated complex decays into free opsin and all-trans-retinal (1). All-trans-retinal is then reduced to all-transretinol by a retinol dehydrogenase and is translocated from the photoreceptors to the retinal pigment epithelium (RPE), where it is converted back into 11-cis-retinal. The recycled 11-cis-retinal is then sent back to the photoreceptors where it recombines covalently with opsin to form the ground-state visual pigment molecule (2). However, experimental detachment of the retina from the RPE interrupts this visual cycle and prevents the recycling of chromophore and regeneration of the bleached visual pigment. As a result, after exposure of the isolated retina or isolated photoreceptors to bright light, most of the visual pigment is converted to free opsin which is now available for pigment regeneration (3, 4). Application of exogenous retinoid analogs to such bleached photoreceptors allows investigating the noncovalent and covalent binding properties of chromophore to opsin by physiological techniques. The second approach is based on the tremendous progress in the techniques of molecular biology that has occurred during the last two decades, which has made possible the expression of mutant or foreign opsins in photoreceptors. These techniques allow studies of the effects of opsin mutations on the signaling properties of the visual pigments. Furthermore, as rods and cones use distinct forms of opsin but share the same chromophore (11-cis-retinal) (5), transgenic expression of rod and cone opsins allows examination of the differences in sensitivity and response kinetics that derive from differences in opsin structure. 1.1. Animal Models

Salamander (Ambystoma tigrinum) has been the animal of choice for single-cell recordings from rod and cone photoreceptors to investigate different aspects of the interaction between retinoid and opsin using retinoid analogs (6, 7). This well-established

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Fig. 5.1. Suction electrode configuration for recording from single photoreceptors. (a) Dissociated salamander rod with its inner segment drawn in the suction electrode and the outer segment protruding out of it. (b) Mouse rod with its outer segment drawn in the suction electrode and the inner segment still attached to a piece of retina.

preparation offers large and abundant rods (Fig. 5.1a) and cones that can be easily dissociated and maintained in culture. Physiological recordings from salamander photoreceptors are stable over hours and allow extended and rigorous experimental protocols. This greatly facilitates experimental approaches involving replacement of the native chromophore, because the decay of photoactivated pigments and their subsequent regeneration with exogenous chromophore can take as much as 1–2 h. The animals of choice for studies of transgenic opsins have been Xenopus laevis and mouse. Xenopus photoreceptors are relatively large and provide stable and reproducible recordings (8). In addition, the high yield of transgenic Xenopus animals produced by oocyte injection makes unnecessary the breeding and maintenance of transgenic lines for extended periods of time. However, the native chromophore in Xenopus (11-cis-3,4-dehydroretinal or A2) is slightly different from the native chromophore in most mammals, including mouse and human (11-cis-retinal or A1). As a result, pigment properties that depend on the chromophore are likely to differ in Xenopus and mammalian photoreceptors. Another drawback of this preparation has been the lack of developed tools for deleting endogenous genes. This has limited Xenopus studies to the transgenic expression of exogenous opsin genes. Mice, on the other hand, are amenable to both transgenic and gene knockout manipulations. In addition, mouse studies can take advantage of the wide and continuously increasing number of genetically modified lines including transgenic and knockout

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animals. Finally, mouse rod recordings (see Fig. 5.1b) have been used routinely for over a decade to investigate rod phototransduction (9). Although the methods developed for regenerating salamander visual pigments with exogenous retinoid analogs are yet to be widely used in mouse photoreceptors, recent studies indicate that the same methods might be applicable there (10–12). Finally, although mouse cone recordings have been challenging, the recent development of genetically modified mice and the creative use of single-cell and electroretinographic (ERG) recording techniques have proven successful (13, 14). This indicates the feasibility of physiological studies of mouse cone pigment properties in their native environment. 1.2. Chromophore Analog Studies

Pigment regeneration takes place in two steps. Initially, the chromophore binds noncovalently in a hydrophobic pocket in the core of opsin (15). Then the aldehyde group located at the end of its polyene chain forms a Schiff-base covalent bond with a lysine residue of opsin (16). As the amount of free 11-cisretinal in the retina (17) and specifically in photoreceptors (18) is minimal, bright light exposure can remove a large fraction of the chromophore from opsin, provided the photoreceptors have been removed from the retina and isolated from other cells and RPE. This allows the application of various exogenous retinoids to photoreceptors to investigate the interactions between opsin and chromophore and the role of specific parts of the retinoid molecule in the function of visual pigment. With this approach, the noncovalent interaction between opsin and chromophore can be investigated with analogs of 11-cis-retinal having a shortened or modified side chain that are capable of binding in the chromophore pocket of opsin but not of forming a covalent bond with it (19). Studies with such retinoids have revealed that in rods, the noncovalent binding of retinoid to opsin upregulates its activity and results in activation of the rod phototransduction cascade, producing desensitization of the rod and acceleration of its flash response (7, 20–22). In contrast, in red cones the binding of retinoid inactivates opsin and relieves adaptation, thus increasing cone sensitivity and slowing the time course of its flash response (4, 21, 23). As a result, in rods the noncovalent binding of retinoid, including the native 11-cisretinal, to opsin slows down dark adaptation. In red cones, however, the noncovalent binding of retinoid to cone opsin reverses the effects of bleaching adaptation and accelerates cone recovery from a bleach even before the pigment can be regenerated (24). This noncovalent interaction between cone opsin and its chromophore is believed to be one of the mechanisms contributing to the faster dark adaptation of cones compared to rods. A similar experimental approach has been used successfully to investigate the role of the methyl group at position 9 on the

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polyene side chain of retinal. Removal of that group produces 9-demethyl retinal which can form a 9-demethyl visual pigment in both rods and cones (25, 26). In rods, 9-demethyl visual pigment produces a quantal response that is about 30 times smaller and decays 5 times slower than that of the native pigment (27). These results reveal that the 9-methyl group is critical for controlling the activation of the G protein transducin by the rod visual pigment as well as for the inactivation of the rod visual pigment by rhodopsin kinase and arrestin. In cones, on the other hand, 9-demethyl visual pigment produces a quantal response with an amplitude and kinetics that are identical to those of the native pigment (28). However, for flashes activating more than ∼0.2% of the 9-demethyl cone pigment, response inactivation with increasing flash intensity is progressively slower than that of the native cone pigment (Fig. 5.2). These results are consistent with the slower decay of the physiologically active meta II state of

Fig. 5.2. Comparison of response termination in red salamander cones with pigment containing 11-cis-retinal (black traces) and 9-demethyl retinal (gray traces). Responses produced by the two pigments were identical for low flash strengths (<0.1% bleach). For brighter flashes, termination of 9-demethyl retinal cone pigment responses became progressively slower than the termination of 11-cis-retinal cone pigment responses. Reprinted with permission from (28).

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9-demethyl visual pigment (25) and indicate that, in bright light, the inactivation of cone photoresponses may be rate limited by cone meta II decay. Other physiological studies using the technique of chromophore replacement in salamander rods have been performed to examine the role of the chromophore in modulating the spontaneous thermal activation of the visual pigment. Vertebrate visual pigments incorporate either 11-cis-retinal (A1, e.g., mouse, bovine, and human) or 11-cis-3,4-dehydroretinal (A2, e.g., Xenopus, bullfrog) as their chromophore. The salamander presents a curious case because its pigment undergoes a shift from the A2 to A1 chromophore type as the animal metamorphoses from the larval to adult stage (29, 30). Larval salamander rods with predominantly A2 as their native chromophore can be bleached and then regenerated with A1 chromophore. Studies of this kind have revealed that A1 rod pigment is at least 36 times more stable than the corresponding A2 pigment (31). As a result, spontaneous activation of salamander rod A2 pigment occurs more frequently than that of the corresponding A1 pigment and may produce adaptation of the rod. Although the rate of thermal activation for cone pigments is significantly higher than that of rods, the stability of human L-cone A1 pigment has also been estimated to be 40fold higher than that of the corresponding A2 pigment (32). The effect of cone pigment thermal activation on cone physiology is discussed below. Finally, physiological recordings have also allowed the investigation of the stability of the covalent bond between opsin and 11-cis-retinal in rod and cone visual pigments. Biochemical studies had suggested that the formation of the covalent bond between 11-cis-retinal and opsin is reversible in cones but irreversible in rods (33, 34). These results predicted the presence of a dynamic equilibrium between chromophore-bound and chromophore-free opsin in dark-adapted cones, which should be shifted in opposite directions by applying excess chromophore or chromophore-binding proteins. In addition, excess exogenous chromophore would be expected to displace gradually the native chromophore in cone pigment even in darkness. Indeed, application of exogenous 9-cis-retinal to dark-adapted isolated salamander red cones results in a gradual blue spectral shift consistent with the gradual replacement of the native 11-cis-retinal with 9-cis-retinal as the cone chromophore (18). Furthermore, treatment of isolated cones with the 11-cis-specific chromophorebinding protein CRALBP results in their gradual desensitization which can be reversed by application of 11-cis-retinal (18). Pigment dissociation into opsin and 11-cis-retinal has been observed in salamander red and blue cones as well as in green rods, but not in red rods. Notably, the equilibrium between chromophorebound and chromophore-free opsin in dark-adapted cones is such

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that ∼10% of opsin is in the apo-protein state. Despite the very weak catalytic activity of opsin, this surprisingly large fraction of free opsin in dark-adapted cells produces substantial activation of the cone phototransduction cascade. Hence, in salamander the reversibility of the covalent bond formation in cone, but not rod, visual pigment contributes to the difference in sensitivity between rods and cones. In red cones, the resulting desensitization is equivalent to that produced by background light activating ∼500 pigment molecules per second (Fig. 5.3) (18). Notably, this level of activity is comparable to the effect of thermal activation by red cone pigment (35, 36). Thus, in salamander red cones the dissociation of cone pigment and its thermal activation produce comparable effects on cone sensitivity. However, unlike thermal activation, the opsin desensitization can be removed by applying excess 11-cis-retinal to the dark-adapted salamander cones (18). So far, chromophore analog studies as those listed above have been done exclusively in amphibian photoreceptors. However, recent work showing that exogenous retinal can be successfully incorporated into mouse rods (12) indicates the feasibility of such studies in mammalian photoreceptors as well. A useful preparation for studies of this kind is the Rpe65 –/– mouse (37), which lacks the gene for the retinal isomerase, the enzyme in the RPE that converts all-trans-retinyl ester to 11-cis-retinol (38). Rods from Rpe65–/– mice have very little chromophore, and their outer segments contain mostly bare opsin. Since rod opsin is noisy in dark adapted

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Fig. 5.3. Desensitization of salamander red cones by free opsin produced by pigment dissociation and by pigment activation. The desensitization produced by free opsin ( opsin, filled circle) was measured by applying 100 μM 11-cis-retinal to dark-adapted salamander red cones. To obtain the equivalent level of pigment activation, the cells were next exposed to a series of background lights of increasing intensity and the corresponding desensitization was measured (filled squares). By interpolation, the desensitization produced by free opsin was equivalent to that produced by the photoactivation of ∼500 cone pigments per second. This level of pigment activation is comparable to that produced by spontaneous thermal activation of salamander red cone pigment in darkness. Reprinted with permission from (18).

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darkness (3), the rods are desensitized (see Fig. 5.4) and behave as if in the presence of a continuous background light (39), with a reduced circulating current and greatly accelerated response decay to a brief stimulus. When lipid vesicles containing 11-cisretinal are added to the rods, the time course of response recovery (Fig. 5.4a, b) and the sensitivity (Fig. 5.4c) recover to values indistinguishable from those of dark-adapted wild-type rods. The dark adaptation of Rpe65 –/– rods with exogenous 11-cis-retinal demonstrates that retinoids can be successfully added to mouse rods lacking chromophore so as to completely regenerate the pigment and relieve the adaptation produced by the constitutively active opsin. The question still remains whether pigment regeneration with exogenous chromophore is possible for wild-type and other mouse strains in experimental conditions where the native chromophore has been removed by bleaching light.

Fig. 5.4. Addition of vesicles containing 11-cis-retinal to Rpe65−/− rods. (a) Responses of Rpe65−/− rods before (grey) and after (black) 30–60 min incubation with vesicles containing 11-cis-retinal. Responses in both cases have been averaged from 17 rods. Intensities of flashes were 4.9 × 104 photons μm−2 (grey) and 43 photons μm−2 (black). (b) Traces in (a) were normalized cell by cell to the maximum response amplitude of the flash and then averaged. Black lines are fitted to single exponential decay function with time constants of 75 ms (grey) and 164 ms (black). (c) Response–intensity curves of Rpe65−/− rods before (open triangles) and after (open squares) addition of vesicles containing 11-cis-retinal. Current densities were calculated by dividing by OS surface area. Intensities in rhodopsins bleached were calculated from light intensities in photons per square cm and collecting areas, estimated for Rpe65−/− rods to be 0.10 before addition of vesicles and 0.31 after vesicle addition. Difference in collecting area before and after is result of different absorption peak and quantum efficiency of 9-cis-based isorhodopsin pigment. Filled squares and curves are for WT rods shown for comparison. Reproduced with permission from (12).

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The small number of cones as a percentage of the total photoreceptor population (3–5%) in most mammalian species, including mouse and human, has limited the biochemical studies of cone phototransduction proteins, including cone visual pigments. As a result, the properties of cone pigments are not as well understood as those of rods. The use of transgenic animals has allowed the expression of cone opsins in rods in order to investigate their signaling properties. In addition, the use of genetically modified mice has allowed the investigation of the mechanisms by which mutations in opsins lead to retinal diseases. It has been long known that cones are 30- to 100-fold less sensitive than rods (40, 41), although the mechanisms for this difference in sensitivity had been poorly understood. One possible explanation for this lower sensitivity of cones is the faster decay of the physiologically active meta II state of cone pigments compared to rod pigments (42–44). The generation of transgenic animals co-expressing rod and cone opsins has made the testing of this hypothesis possible. With transgenic animals, rod and cone opsins can be co-expressed in the same photoreceptor allowing direct comparison of their signaling properties. Such studies from transgenic Xenopus (35) and mice (32, 45–47) expressing various cone opsins have revealed that, surprisingly, cone pigments produce rod-like responses when expressed in rods. For instance, the expression of mouse cone S-opsin in mouse rods that lack their native rod pigment (rho−/− ) results in a dramatic spectral shift in rod sensitivity and renders the rods most sensitive to ultraviolet light (Fig. 5.5). Nevertheless, the flash responses produced by cone S-opsin are comparable to those produced by the native rod pigment (Fig. 5.5a, b). Similarly, rod pigment produces conelike responses when expressed in transgenic Xenopus cones (35). These results indicate that rod and cone visual pigments have similar signaling properties. Thus, rod and cone pigments activate rod transducin and are inactivated by rod rhodopsin kinase and arrestin with similar efficiencies. These results also demonstrate that meta II decay is not the rate-limiting step for the inactivation of the photoresponse under dark-adapted conditions. The studies of transgenic Xenopus and mouse rods expressing cone opsins have also revealed an interesting dichotomy between the function of cone pigments in these two species. In Xenopus rods, the thermal activation of transgenic red cone pigment is some 10,000-fold higher than that of the rod pigment (35). The high molecular rate of thermal activation of red cone pigment is consistent with the estimate of noise from thermal pigment activation in salamander red cones (36, 48) and indicates that in animals using A2 11-cis-retinal, thermal activation of red cone pigment contributes significantly to the low sensitivity of cones compared to rods. In contrast, in transgenic mouse rods,

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the same red cone pigment, while still 600 times more thermally active than rod pigment (32), has a substantially lower rate of thermal activation than would be expected from the Xenopus and salamander studies. This result indicates that in animals using A1 11-cis-retinal, such as mouse and human, thermal activation of cone pigment is not significant enough to affect cone sensitivity. The transgenic expression of cone opsins in rods has also allowed the measurement of the rate of decay of the meta II state of cone pigments under physiological conditions. Whereas in vitro meta II decays spontaneously, in intact photoreceptors the photoactivated visual pigment is rapidly inactivated, initially by phosphorylation by rhodopsin kinase and then by the binding of arrestin (49). In arrestin-deficient (arr1–/–) mouse rods, the response displays a biphasic shutoff, with normal initial inactivation by rhodopsin kinase, followed by a slow tail that represents the decay of rod meta II (Fig. 5.6a) (50). The same method

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Fig. 5.6. Measurement of the physiologically active meta II state of rod and cone pigments expressed in rods lacking arrestin. (a) Normalized dim flash response of mouse rod lacking arrestin (rho+/+ arr−/− ). The slow tail of the response inactivation is fit by a single exponential decay function with τ = 51 s which represents the decay of rod pigment meta II. (b) Normalized dim flash response of mouse rod that expresses exclusively S-opsin and lacks arrestin (S-opsin+/− rho−/− arr−/− ) showing that τ = 1.2 s for S-opsin meta II. (c) Normalized dim flash response of mouse rod that expresses exclusively L-opsin and lacks arrestin (L-opsin+/− rho−/− arr−/− ) showing that τ = 0.56 s for L-opsin meta II. Reprinted with permission from (47) and (32).

can be used to observe the decay of cone meta II. The deletion of arrestin in transgenic rods expressing cone opsin removes the inactivation of phosphorylated cone pigment. This renders the decay of the active cone meta II state the rate-limiting step for photoresponse inactivation, which can be observed experimentally. Such experiments reveal that whereas the time constant of the rod meta II decay is ∼50 s (Fig. 5.6a), mouse S-cone meta II decays in ∼1.3 s (Fig. 5.6b) (47), and human L-cone meta II in only 0.6 s (Fig. 5.6c) (32). These measurements were done from intact photoreceptors at 37◦ C and most likely reflect accurately the in vivo decay rates of S- and L-cone meta II. 1.4. Rhodopsin Mutation Studies

Transgenic expression of mutant rod opsin has also been used to investigate the mechanism by which certain rhodopsin mutations lead to congenital night blindness. Patients with the rhodopsin G90D mutation and several other rhodopsin mutations have been shown to have a persistent loss of rod sensitivity, produced by a mechanism that is similar if not identical to that produced by steady background light (51, 52). Two possible explanations have been given for why the G90D mutation of opsin might result in constitutive activation of the rods. According to one model, the G90D mutation produces opsin without chromophore with a high constitutive activity (53). Several studies have shown that G90D opsin in vitro is very effective in stimulating the G protein transducin (54). According to the second model, it is G90D rhodopsin rather than G90D opsin that produces spontaneous activity of the cascade, whether by a high rate of spontaneous conversion of G90D rhodopsin to meta II (51, 52) or by some other mechanism (12).

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The validity of these two hypotheses has been tested by introducing the mutant G90D rhodopsin into rods of two species, Xenopus and mouse. In transgenic Xenopus rods (55), expression of the mutant G90D opsin produces constitutive activity and results in desensitization and acceleration of the decay of the flash response (55). Normal sensitivity (Fig. 5.7a) and response kinetics (Fig. 5.7b) can be restored with the application of exogenous 11-cis-retinal. This result shows that in dark-adapted Xenopus rods the spontaneous activation responsible for the reduction in sensitivity is produced by the high constitutive activity of G90D opsin, which can be removed upon the covalent attachment of the reverse agonist 11-cis-retinal. A

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Fig. 5.7. Desensitization in night blindness mutant G90D and phenotypic rescue with 11-cis-retinal in Xenopus rod photoreceptors. Intensity response curves (a) and dim flash responses (b) were measured in single cells isolated from retinae of wild-type frogs and transgenic frogs containing the rhodopsin EGFP gene with the G90D mutation. Normalized curves plotted in (a) were constructed from averages of peak flash response amplitude elicited from different isolated rods at different flash intensities: wild type, n = 6; G90D, n = 7; G90D+11-cis-retinal, n = 8. Error bars indicate standard deviation. Dim flash responses plotted in (b) are averaged from rods from different animals: wild type, n = 6; G90D, n = 7; G90D + 11-cis-retinal, n = 6. Reprinted with permission from (55).

In transgenic mouse rods, expression of the mutant G90D opsin also produces constitutive activity, which results in decreased circulating current and lower sensitivity, as well as an accelerated decay of the flash response (12, 51, 52). However, the desensitization produced by mammalian G90D pigment expressed in a mammal is 200–3000 times smaller than that produced when the pigment is expressed at the same concentration in Xenopus (12). In addition, unlike in Xenopus, normal sensitivity (Fig. 5.8a) and response kinetics (Fig. 5.8a, inset) in a mouse rod cannot be restored by the addition of exogenous 11cis-retinal (12). If 11-cis-retinal is added to an Rpe65 –/– mouse containing the normal rhodopsin gene, the sensitivity of the rod recovers to that in a wild-type animal, as we have seen (Fig. 5.4). But if 11-cis-retinal is added to rods from an Rpe65 –/– mouse with G90D pigment and no wild-type pigment, the sensitivity

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Fig. 5.8. Addition of 11-cis-retinal to G90D mouse rods. (a) Response–intensity curves for 30 D+/+ rods homozygous for the G90D mutation before treatment with retinal (black squares) and 15 D+/+ rods after 1–3 h of vesicle incubation (open circles). Measurements were made from rods from the same retinas. Current densities were calculated by dividing by outer segment area of 99 μm2 ; collecting area was 0.40. Small shift in curves was not statistically significant. Filled circles and curves are for WT rods shown for comparison. Inset: mean unnormalized responses to 20 ms flash of 1.5 × 105 photons μm−2 from D+/+ rods homozygous for the G90D mutation, averaged from 24 rods before treatment (black trace) and 20 rods after 70–140 min treatment with phospholipid vesicles containing 11-cis-retinal (grey trace). (b) Addition of vesicles to D+/− ; Rh−/− rods with or without the RPE65 isomerase. Black squares and curve are mean responses from 24 D+/− ; Rh−/− ; Rpe65−/− rods before addition of vesicles. Current density was calculated from outer segment area of 40.5 μm2 ; collecting area was 0.06 [see (13)]. Open circles and curves are mean responses from 24 D+/− ; Rh−/− ; Rpe65−/− rods after addition of vesicles containing 11-cis-retinal. Outer segment area was 69 μm2 and collecting area was 0.28.

recovers only to that of a G90D rod with its normal complement of chromophore (Fig. 5.8b). Thus, the lack of recovery of G90D-expressing mouse rods cannot be attributed to the failure of exogenous chromophore to promote G90D pigment regeneration. Instead, these experiments show that in a dark-adapted mouse the spontaneous activation responsible for the elevation of threshold is produced by G90D rhodopsin bound to chromophore and not by G90D opsin. Because the two studies in Xenopus and mouse used identical recording methods, the most likely explanation for the discrepancy is that mammalian G90D pigment behaves differently in Xenopus and in mouse rods. Possible factors for this difference include differences in the interaction of G90D opsin with A2 (Xenopus) and A1 (mouse) chromophore, the different body

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temperature of amphibians and mammals, differences in the lipid composition of the outer segment membrane, or different modification of the opsin molecule in Xenopus and mouse rods. Although unlikely, it is also possible that the bovine opsin, used in the Xenopus study, and the mouse opsin, used in the mouse study, are affected differently by the G90D mutation. Regardless of the exact mechanism, the discrepancy between these two studies emphasizes the need for caution when extrapolating animal studies to the function of human visual pigments. These studies also indicate that mammalian pigments may respond differently when expressed in a mammal than when expressed in an amphibian. 1.5. Epilogue

Studies of amphibian rods and cones and transgenic Xenopus and mouse rods have yielded a wealth of information about the interactions between chromophore and opsin, the signaling properties of visual pigments, and the modulation of photoreceptor function by visual pigments. Future studies are likely to focus on two areas. First, the methods of chromophore replacement by bleaching and regeneration developed in salamander photoreceptors need to be extended to mouse photoreceptors. Previous work has shown that chromophore can be successfully introduced into rods lacking their normal complement of photopigment, as for example in Rpe65–/– mice (see Fig. 5.4), but it would be useful to be able to bleach and replace pigment in photoreceptors from wild-type or other genetically modified animals. This would allow the combination of chromophore replacement and transgenic opsin modifications to provide a powerful set of tools to study the interactions between retinoid and opsin in mammalian visual pigments. Second, the recent development of methods for recording from mouse cone photoreceptors will allow investigating the properties of cone visual pigments in their native environment and how these properties modulate the function of mammalian cones. It will also make possible experiments investigating the role of cone pigment properties for cone function not only in darkness but also in bright light or during dark adaptation, both critical for the photoreceptors that mediate our daytime, acute, and color vision.

2. Materials 2.1. Perfusion Solutions

1. Salamander perfusion solution: 110 mM NaCl, 2.5 mM KCl, 1.6 mM MgCl2 , 1.0 mM CaCl2 , 10 mM dextrose, 10 mM HEPES, pH 7.8, and bovine serum albumin (100 mg/l) (3).

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2. Mouse perfusion solution: 112.5 mM NaCl, 3.6 mM KCl, 2.4 mM MgCl2 , 1.2 mM CaCl2 , 10 mM HEPES (pH 7.4), 20 mM NaHCO3 , 3 mM Na succinate, 0.5 mM Na glutamate, 0.02 mM EDTA, and 10 mM glucose. The solution is bubbled with 95% O2 /5% CO2 and warmed to 36–38◦ C (see Note 1). 3. Mouse recording electrode solution: 140 mM NaCl, 3.6 mM KCl, 2.4 mM MgCl2 , 1.2 mM CaCl2 , 3 mM HEPES (pH 7.4), 0.02 mM EDTA, and 10 mM glucose (47). 2.2. Preparation and Application of Retinoid Solution

The application of retinoids to photoreceptors is complicated by their high hydrophobicity. The two most practical approaches to preparing retinoid solution include suspending the retinoid in lipid vesicles or dissolving it in ethanol and diluting that solution in Ringer (6, 12). 1. Aliquots containing 300 μg dry retinoid are prepared by dissolving the retinoid in ethanol to 10 mg/ml, aliquoting 30 μl solution in individual small conical vials, drying under gentle stream of nitrogen (see Note 2), and then storing in darkness at −80◦ C until use. 2. Phospholipid vesicles are prepared by placing 25 mg L-αphosphatidylcholine (type V-E; Sigma) in a glass scintillation vial and evaporating the solvent to dryness under a stream of nitrogen. Ringer solution (15 ml) is then added to the vial and sonicated for 20 min in an ice bath (1 s on/ 1 s off) at 45 W using a 1.0-cm probe. Fresh vesicle solution is prepared daily. One hour before the beginning of the experiment, 1.5 ml vesicle solution is added to a vial of dry retinoid in darkness and sonicated for 60 s on low power using a tapered microtip. The retinoid content in the solution is determined from the absorption of a small aliquot, diluted 10-fold in ethanol, using the molar extinction coefficient for the particular retinoid. 3. An alternative method for dissolving the retinoid in Ringer solution involves using ethanol. Initially, 4 μl ethanol is added to the bottom of the vial containing the dry retinoid. The solution is stirred gently using the pipet tip and then Ringer is gradually added to the vial to a final volume of 4 ml. The resulting 0.1% ethanol solution can then be applied to the photoreceptor preparation. Although not as efficient as the lipid vesicle method, the ethanol solution offers the advantage that it can be prepared very quickly, typically after the photoreceptors have been bleached. With the perfusion off, a small volume of retinoid solution (∼0.5 ml) can be added to the recording chamber, and the photoreceptors can be incubated for ∼2 min before turning the

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perfusion on again. The chamber should be rinsed with ethanol between experiments to ensure that it is clean of retinoid.

3. Methods The single-cell suction electrode technique allows recordings from individual rod and cone photoreceptors without penetrating their plasma membrane. The method, developed by Yau, Lamb, and Baylor (56), is based on the polar nature of photoreceptors with inward current flowing at their outer segment, matched by outward current at their inner segment. Drawing the photoreceptor outer segment (see Note 3) into a tightly fitting glass electrode (Fig. 5.1) allows collection of that current, which can then be amplified, digitized, and stored on a computer for analysis. Exposure of the photoreceptor to a test flash activates the phototransduction cascade and results in closure of the cGMP transduction channels in the outer segment which can be observed as a decrease in the current recorded from the outer segment. Inversed, but otherwise identical, photoresponses can be recorded if the inner segment of the photoreceptor is drawn into the suction electrode instead. Experiments are typically performed after animals are dark adapted overnight. 1. The animal is killed and the eyes are removed and hemisected under dim red light illumination (see Note 4). All subsequent manipulations are performed in infrared light with the help of infrared image converters. 2. The retina is torn free of the pigment epithelium and chopped into small pieces with a razor blade. Overdoing this step can result in a preparation with many broken outer segments but few intact cells. Gently drawing and releasing the solution containing retinal pieces through a transfer pipette also helps in dissociating photoreceptors. 3. A fraction of the resulting suspension is transferred to a recording chamber fit on the stage of an inverted microscope (see Note 5). 4. Recordings can be performed either from dissociated photoreceptors or from the outer segments of cells protruding from a piece of retina. As salamander photoreceptors are rather sturdy and continue to produce robust photoresponses for hours after dissociation from the retina, suction recordings are typically done from dissociated cells. As dissociated mouse rods do not usually yield robust and last-

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ing responses, mouse rod recordings are typically performed from the outer segments of cells protruding from a piece of retina. 5. Test flashes as well as bleaching and background lights are provided from a calibrated dual-beam optical stimulator (57). Flash intensity and wavelength are controlled by a set of neutral density filters and narrow band interference filters, respectively. Test flash duration, typically 10–20 ms, is controlled by computer-driven shutters. 6. The fraction of bleached pigment can be estimated from the relation F = 1 – exp(–IPt), where F is the fraction of bleached pigment, I is the light intensity in photons per square micrometer per second, and t is the duration of light exposure in seconds. The value used for the photosensitivity of the cell P is 6.2 × 10−9 μm2 for rods (58) and 6.0 × 10−9 μm2 for cones (59).

4. Notes 1. The perfusion solution can be brought to 36–38◦ C by running it through a ceramic resistor (e.g., SBCHE633RJ; Tyco Electronics) under small DC voltage via stainless steel tubing. The heater should be located as close to the recording chamber as possible, ideally at the stage of the microscope. To prevent bubbles from forming in the tubing, the solution should be pre-heated to 36–38◦ C when bubbling with 95% O2 /5% CO2 . 2. When preparing retinoid aliquots by drying retinoid ethanol solution, the gas stream should be low enough to prevent the solution from splashing along the walls of the vial. This makes resuspending the retinoid easier and more efficient. 3. A simple suction device can be constructed by connecting tubing partially filled with mineral oil to the side port of the electrode holder. The other end of the tubing is connected to a small reservoir of mineral oil (1 cm3 syringe works well). Raising or lowering the reservoir applies positive or negative pressure to the tip of the electrode. Suction for drawing the cell in the electrode can be applied by mouth through tubing and a needle drawn through the piston of the syringe. 4. An easy method to achieve clean and fast eye hemisection involves the use of two matching plastic blocks. The bottom one has a row of round semi-spherical wells of different diameters, matching the size of eyeballs of different species

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(e.g., fish, mouse, salamander). The eyeball is placed in the matching well, iris facing up, and then clamped with the second plastic block that has holes matching the wells in the bottom block. A clean cut of the eye can be achieved by slicing a razor blade between the two blocks. Usually, the lens is pushed out through the hole in the top block, leaving a clean-cut eyecup in the well of the bottom block. 5. Stopping the perfusion for ∼2 min right after adding the cell suspension to the recording chamber allows most cells to settle on the bottom of the chamber and not get washed away by the solution flow. If necessary, a bypass tubing can be added to prevent the chamber from drying while the perfusion is off. References 1. Ebrey, T., Koutalos, Y. (2001) Vertebrate photoreceptors. Prog. Retin. Eye Res. 20, 49–94. 2. Saari, J.C. (2000) Biochemistry of visual pigment regeneration: The Friedenwald lecture. Invest. Ophthalmol. Vis. Sci. 41, 337–348. 3. Cornwall, M.C., Fain, G.L. (1994) Bleached pigment activates transduction in isolated rods of the salamander retina. J. Physiol. 480(Pt 2), 261–279. 4. Cornwall, M.C., Matthews, H.R., Crouch, R.K., Fain, G.L. (1995) Bleached pigment activates transduction in salamander cones. J. Gen. Physiol. 106, 543–557. 5. Yokoyama, S. (2000) Molecular evolution of vertebrate visual pigments. Prog. Retin. Eye Res. 19, 385–419. 6. Cornwall, M.C., Jones, G.J., Kefalov, V.J., Fain, G.L., Matthews, H.R. (2000) Electrophysiological methods for measurement of activation of phototransduction by bleached visual pigment in salamander photoreceptors. Methods Enzymol. 316, 224–252. 7. Crouch, R.K., Kefalov, V., Gartner, W., Cornwall, M.C. (2002) Use of retinal analogues for the study of visual pigment function. Methods Enzymol. 343, 29–48. 8. Xiong, W.H., Yau, K.W. (2002) Rod sensitivity during Xenopus development. J. Gen. Physiol. 120, 817–827. 9. Lem, J., Makino, C.L. (1996) Phototransduction in transgenic mice. Curr. Opin. Neurobiol. 6, 453–458. 10. Fan, J., Woodruff, M.L., Cilluffo, M.C., Crouch, R.K., Fain, G.L. (2005) Opsin activation of transduction in the rods of darkreared Rpe65 knockout mice. J. Physiol. 568, 83–95.

11. Luo, D.G., Yau, K.W. (2005) Rod sensitivity of neonatal mouse and rat. J. Gen. Physiol. 126, 263–269. 12. Dizhoor, A.M., Woodruff, M.L., Olshevskaya, E.V., Cilluffo, M.C., Cornwall, M.C., Sieving, P.A., Fain, G.L. (2008) Night blindness and the mechanism of constitutive signaling of mutant G90D rhodopsin. J. Neurosci. 28, 11662–11672. 13. Nikonov, S.S., Kholodenko, R., Lem, J., Pugh, E.N., Jr. (2006) Physiological features of the S- and M-cone photoreceptors of wildtype mice from single-cell recordings. J. Gen. Physiol. 127, 359–374. 14. Heikkinen, H., Nymark, S., Koskelainen, A. (2008) Mouse cone photoresponses obtained with electroretinogram from the isolated retina. Vision Res. 48, 264–272. 15. Matsumoto, H., Yoshizawa, T. (1975) Existence of a beta-ionone ring-binding site in the rhodopsin molecule. Nature 258, 523–526. 16. Bownds, D. (1967) Site of attachment of retinal in rhodopsin. Nature 216, 1178–1181. 17. Lyubarsky, A.L., Pugh, E.N., Jr. (2007) Over 98% of 11-cis retinal in the dark-adapted mouse eye is bound to rod and cone opsins. Invest. Ophthalmol. Vis. Sci. 48, 3246. 18. Kefalov, V.J., Estevez, M.E., Kono, M., Goletz, P.W., Crouch, R.K., Cornwall, M.C., Yau, K.W. (2005) Breaking the covalent bond – a pigment property that contributes to desensitization in cones. Neuron 46, 879–890. 19. Crouch, R.K. (1986) Studies of rhodopsin and bacteriorhodopsin using modified retinals. Photochem. Photobiol. 44, 803–807.

Physiological Studies of the Interaction Between Opsin and Chromophore 20. Kefalov, V.J., Carter Cornwall, M., Crouch, R.K. (1999) Occupancy of the chromophore binding site of opsin activates visual transduction in rod photoreceptors. J. Gen. Physiol. 113, 491–503. 21. Corson, D.W., Kefalov, V.J., Cornwall, M.C., Crouch, R.K. (2000) Effect of 11-cis 13demethylretinal on phototransduction in bleach-adapted rod and cone photoreceptors. J. Gen. Physiol. 116, 283–297. 22. Isayama, T., Chen, Y., Kono, M., Degrip, W.J., Ma, J.X., Crouch, R.K., Makino, C.L. (2006) Differences in the pharmacological activation of visual opsins. Vis. Neurosci. 23, 899–908. 23. Jin, J., Crouch, R.K., Corson, D.W., Katz, B.M., MacNichol, E.F., Cornwall, M.C. (1993) Noncovalent occupancy of the retinal-binding pocket of opsin diminishes bleaching adaptation of retinal cones. Neuron 11, 513–522. 24. Kefalov, V.J., Crouch, R.K., Cornwall, M.C. (2001) Role of noncovalent binding of 11-cis-retinal to opsin in dark adaptation of rod and cone photoreceptors. Neuron 29, 749–755. 25. Das, J., Crouch, R.K., Ma, J.X., Oprian, D.D., Kono, M. (2004) Role of the 9-methyl group of retinal in cone visual pigments. Biochemistry 43, 5532–5538. 26. Ganter, U.M., Schmid, E.D., Perez-Sala, D., Rando, R.R., Siebert, F. (1989) Removal of the 9-methyl group of retinal inhibits signal transduction in the visual process. A Fourier transform infrared and biochemical investigation. Biochemistry 28, 5954–5962. 27. Corson, D.W., Cornwall, M.C., MacNichol, E.F., Tsang, S., Derguini, F., Crouch, R.K., Nakanishi, K. (1994) Relief of opsin desensitization and prolonged excitation of rod photoreceptors by 9-desmethylretinal. Proc. Natl. Acad. Sci. USA 91, 6958–6962. 28. Estevez, M.E., Ala-Laurila, P., Crouch, R.K., Cornwall, M.C. (2006) Turning cones off: The role of the 9-methyl group of retinal in red cones. J. Gen. Physiol. 128, 671–685. 29. Harosi, F.I. (1975) Absorption spectra and linear dichroism of some amphibian photoreceptors. J. Gen. Physiol. 66, 357–382. 30. Makino, C.L., Dodd, R.L. (1996) Multiple visual pigments in a photoreceptor of the salamander retina. J. Gen. Physiol. 108, 27–34. 31. Ala-Laurila, P., Donner, K., Crouch, R.K., Cornwall, M.C. (2007) Chromophore switch from 11-cis-dehydroretinal (A2) to 11-cis-retinal (A1) decreases dark noise in salamander red rods. J. Physiol. 585, 57–74.

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32. Fu, Y., Kefalov, V., Luo, D.G., Xue, T., Yau, K.W. (2008) Quantal noise from human red cone pigment. Nat. Neurosci. 11, 565–571. 33. Crescitelli, F. (1984) The gecko visual pigment: The dark exchange of chromophore. Vision Res. 24, 1551–1553. 34. Matsumoto, H., Tokunaga, F., Yoshizawa, T. (1975) Accessibility of the iodopsin chromophore. Biochim. Biophys. Acta 404, 300–308. 35. Kefalov, V., Fu, Y., Marsh-Armstrong, N., Yau, K.W. (2003) Role of visual pigment properties in rod and cone phototransduction. Nature 425, 526–531. 36. Rieke, F., Baylor, D.A. (2000) Origin and functional impact of dark noise in retinal cones. Neuron 26, 181–186. 37. Redmond, T.M., Yu, S., Lee, E., Bok, D., Hamasaki, D., Chen, N., Goletz, P., Ma, J.X., Crouch, R.K., Pfeifer, K. (1998) Rpe65 is necessary for production of 11-cis-vitamin A in the retinal visual cycle. Nat. Genet. 20, 344–351. 38. Jin, M., Li, S., Moghrabi, W.N., Sun, H., Travis, G.H. (2005) Rpe65 is the retinoid isomerase in bovine retinal pigment epithelium. Cell 122, 449–459. 39. Woodruff, M.L., Wang, Z., Chung, H.Y., Redmond, T.M., Fain, G.L., Lem, J. (2003) Spontaneous activity of opsin apoprotein is a cause of Leber congenital amaurosis. Nat. Genet. 35, 158–164. 40. Fain, G.L., Dowling, J.E. (1973) Intracellular recordings from single rods and cones in the mudpuppy retina. Science 180, 1178–1181. 41. Schnapf, J.L., Baylor, D.A. (1987) How photoreceptor cells respond to light. Sci. Am. 256, 40–47. 42. Imai, H., Imamoto, Y., Yoshizawa, T., Shichida, Y. (1995) Difference in molecular properties between chicken green and rhodopsin as related to the functional difference between cone and rod photoreceptor cells. Biochemistry 34, 10525–10531. 43. Okada, T., Matsuda, T., Kandori, H., Fukada, Y., Yoshizawa, T., Shichida, Y. (1994) Circular dichroism of metaiodopsin II and its binding to transducin: A comparative study between meta II intermediates of iodopsin and rhodopsin. Biochemistry 33, 4940–4946. 44. Starace, D.M., Knox, B.E. (1997) Activation of transducin by a Xenopus short wavelength visual pigment. J. Biol. Chem. 272, 1095–1100. 45. Imai, H., Kefalov, V., Sakurai, K., Chisaka, O., Ueda, Y., Onishi, A., Morizumi, T.,

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46.

47.

48.

49.

50.

51.

Kefalov, Cornwall, and Fain Fu, Y., Ichikawa, K., Nakatani, K., Honda, Y., Chen, J., Yau, K. W., Shichida, Y. (2007) Molecular properties of rhodopsin and rod function. J. Biol. Chem. 282, 6677–6684. Sakurai, K., Onishi, A., Imai, H., Chisaka, O., Ueda, Y., Usukura, J., Nakatani, K., Shichida, Y. (2007) Physiological properties of rod photoreceptor cells in green-sensitive cone pigment knock-in mice. J. Gen. Physiol. 130, 21–40. Shi, G., Yau, K.W., Chen, J., Kefalov, V.J. (2007) Signaling properties of a short-wave cone visual pigment and its role in phototransduction. J. Neurosci. 27, 10084–10093. Sampath, A.P., Baylor, D.A. (2002) Molecular mechanism of spontaneous pigment activation in retinal cones. Biophys. J. 83, 184–193. Makino, C.L., Wen, X.H., Lem, J. (2003) Piecing together the timetable for visual transduction with transgenic animals. Curr. Opin. Neurobiol. 13, 404–412. Xu, J., Dodd, R.L., Makino, C.L., Simon, M.I., Baylor, D.A., Chen, J. (1997) Prolonged photoresponses in transgenic mouse rods lacking arrestin. Nature 389, 505–509. Sieving, P.A., Richards, J.E., Naarendorp, F., Bingham, E.L., Scott, K., Alpern, M. (1995) Dark-light: Model for nightblindness from the human rhodopsin Gly-90–>Asp mutation. Proc. Natl. Acad. Sci. USA 92, 880–884.

52. Sieving, P.A., Fowler, M.L., Bush, R.A., Machida, S., Calvert, P.D., Green, D.G., Makino, C.L., McHenry, C.L. (2001) Constitutive “light” adaptation in rods from G90D rhodopsin: A mechanism for human congenital nightblindness without rod cell loss. J. Neurosci. 21, 5449–5460. 53. Rao, V.R., Cohen, G.B., Oprian, D.D. (1994) Rhodopsin mutation G90D and a molecular mechanism for congenital night blindness. Nature 367, 639–642. 54. Rao, V.R., Oprian, D.D. (1996) Activating mutations of rhodopsin and other G protein-coupled receptors. Annu. Rev. Biophys. Biomol. Struct. 25, 287–314. 55. Jin, S., Cornwall, M.C., Oprian, D.D. (2003) Opsin activation as a cause of congenital night blindness. Nat. Neurosci. 6, 731–735. 56. Yau, K.W., Lamb, T.D., Baylor, D.A. (1977) Light-induced fluctuations in membrane current of single toad rod outer segments. Nature 269, 78–80. 57. Cornwall, M.C., Fein, A., MacNichol, E.F., Jr. (1990) Cellular mechanisms that underlie bleaching and background adaptation. J. Gen. Physiol. 96, 345–372. 58. Jones, G.J. (1995) Light adaptation and the rising phase of the flash photocurrent of salamander retinal rods. J. Physiol. 487(Pt 2), 441–451. 59. Jones, G.J., Fein, A., MacNichol, E.F., Jr., Cornwall, M.C. (1993) Visual pigment bleaching in isolated salamander retinal cones. Microspectrophotometry and light adaptation. J. Gen. Physiol. 102, 483–502.

Chapter 6 Measurement of the Mobility of All-Trans-Retinol with Two-Photon Fluorescence Recovery After Photobleaching Yiannis Koutalos Abstract The mobility of all-trans-retinol makes a crucial contribution to the rate of the reactions in which it participates. This is even more so because of its low aqueous solubility, which makes the presence of carrier proteins and the spatial arrangement of cellular membranes especially relevant. In rod photoreceptor outer segments, all-trans-retinol is generated after light exposure from the reduction of all-trans-retinal that is released from bleached rhodopsin. The mobility of all-trans-retinol in rod outer segments was measured with fluorescence recovery after photobleaching (FRAP), using two-photon excitation of its fluorescence. The values of the lateral and axial diffusion coefficients indicate that most of the all-transretinol in rod outer segments move unrestricted and without being aided by carriers. Key words: Photoreceptors, rod outer segment, retina, visual cycle, rhodopsin, diffusion.

1. Introduction All-trans-retinol is formed in rod photoreceptor outer segments after light excitation from the reduction of all-trans-retinal released by photoactivated rhodopsin. It is then transferred to the adjacent pigment epithelial cells where it is converted to retinyl ester by lecithin retinol acyltransferase (1–3). The formation of all-trans-retinol removes all-trans-retinal, and its transport out of the rod outer segment further improves through mass action the clearance of all-trans-retinal. In addition, it feeds the retinoid into a recycling pathway that converts it back to 11-cis-retinal used to regenerate rhodopsin. In the rest of the chapter, the terms H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_6, © Springer Science+Business Media, LLC 2010

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retinal and retinol without qualification refer to the all-trans isomers. The mobility of retinol makes a significant contribution to the rate of reactions, such as the removal of retinal or the recycling of the chromophore of rhodopsin, in which it participates. Retinol is highly insoluble in aqueous solutions (4) and its transfer across intracellular and extracellular space is typically aided by specialized carrier proteins (5). Thus, in serum, retinol is carried by retinol-binding protein (RBP), in the interphotoreceptor matrix by interphotoreceptor retinoid-binding protein (IRBP), while within retinal pigment epithelial cells it is transported by cellular retinol-binding protein (CRBP-I). The concentration of retinol can be monitored from its fluorescence (6–9), and its mobility can be measured with fluorescence recovery after photobleaching (FRAP) (10). A FRAP measurement begins with the photobleaching of the fluorophore within a defined volume and then monitors the redistribution of fluorescence as unbleached fluorophore molecules move into that volume. The time course of the recovery of fluorescence reflects the mobility of the fluorophore and can be analyzed to obtain its diffusion coefficient. Today, laser scanning confocal microscopes usually have all the necessary optical components, including a software module, and can routinely be used for FRAP measurements. Retinol absorbs maximally ∼325 nm, which would require the use of an ultraviolet laser line for fluorescence excitation and photobleaching. Another option is two-photon excitation of retinol fluorescence with 700–720 nm light (10, 11), and using the same light for photobleaching. Non-linear optical confocal microscopes incorporate infrared lasers with sufficient power to reach the intensities needed for two-photon excitation of fluorescence (1); they typically have all the necessary hardware and software components to carry out FRAP measurements. An important feature of two-photon excitation and photobleaching is that they take place only at the plane of focus. This has the important advantage of minimizing phototoxicity and fluorophore bleaching out of the focal plane, but it can complicate the analysis of FRAP measurements. In order to obtain the diffusion coefficient from the time course of the fluorescence recovery of a FRAP experiment, it is necessary to have an analytical expression linking the two. Such an expression is typically obtained by solving the diffusion equation for the movement of the fluorophore. In this study, we have used two-photon fluorescence excitation and photobleaching to examine the mobility of retinol in frog rod photoreceptor outer segments. The cylindrical symmetry of the outer segment simplifies the procedure for solving the diffusion equation and obtaining the requisite analytical expressions (10). A more general approach for the analysis of multiphoton FRP experiments that is independent of the geometry of the system has been

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presented (12) but requires detailed characterization and knowledge of the bleaching volume. Because of its low aqueous solubility, virtually all of the retinol in the rod outer segment will be in the membrane compartments, and specifically in the disks, which comprise ∼99% of the total membrane. The movement of retinol within the outer segment will therefore consist of diffusion in the plane of the disk membrane and of transfer between the disks. At the cellular level, diffusion in the plane of the disk membrane will be manifested as a lateral movement, perpendicular to the outer segment axis. On the other hand, transfer between the disks will appear as a longitudinal movement, parallel to the outer segment axis. Because of the cylindrical symmetry of the outer segment, the movement of retinol in the lateral and longitudinal dimensions can be measured separately. The lateral diffusion coefficient of retinol measured with multiphoton FRAP was found to be 2.5 ± 0.3 μm2 s−1 , in close agreement with the diffusion of lipid molecules, suggesting that the bulk of retinol moves freely in the disk membrane. The proper way to measure the lateral diffusion coefficient is through the decline of fluorescence in the unbleached area (Fig. 6.1). Multiphoton FRP measurements also demonstrate that retinol moves along the length of the outer segment

Fig. 6.1. Simulation of the FRAP experiment used for determining lateral diffusion coefficient for a rod outer segment with radius R = 3 μm. The fluorophore (shown as oval-shaped) is assumed to diffuse in the plane of the disk membrane with coefficient D = 2 μm2 s−1 . The scheme on the right shows a transverse cross-section of the outer segment (bleached areas are lightly shaded and are devoid of fluorophore), with the rod lying on the bottom of the chamber and being scanned from the top. The graph on the left shows the corresponding kinetics of fluorescence recovery in bleached and unbleached regions. Fluorescence recovered in the bleached disk area (•) faster than in the unbleached (). The lines are single exponential fits with rate constants of 1.1 and 0.6 s−1 for the bleached and unbleached areas, respectively. Used in conjunction with Eq. [1], these rate constants would result in apparent diffusion coefficients of 4 and 2.2 μm2 s−1 , respectively. Used with Eq. [1], the rate of fluorescence decline in the unbleached area provides a good estimate of the lateral diffusion coefficient. Reprinted from (10) with permission.

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and with a diffusion coefficient of 0.07 ± 0.01 μm2 s−1 . Because of its limited aqueous solubility, this longitudinal movement of retinol is expected to be via the plasma membrane of the outer segment. This interpretation is consistent with the relative values of the lateral and longitudinal diffusion coefficients and the relative areas of the disk and plasma membranes (10).

2. Materials A dark room is necessary for dark-adapting animals and for dissection. An area of ∼50 sq ft is sufficient. A revolving door for entering is convenient, but a thick black curtain is also adequate. 2.1. Photoreceptor Cell Preparation

1. Red lights for the dark room. They are obtained from photographic equipment stores, nowadays through the internet. One choice is adjustable Kodak safelights with filters number 2. For individual red bulbs, an appropriate choice is the Delta 1 Jr. Safelight. It is best to keep the red lights as dim as possible. 2. Frogs (Rana pipiens) are obtained from approved vendors (The Sullivan Company, Nashville, TN; NASCO, Fort Atkinson, WI and Modesto, CA; or Carolina Biological, Burlington, NC). Check with the supplier well in advance for seasonal availability. 3. Amphibian Ringer’s with composition: 110 mM NaCl, 2.5 mM KCl, 1.6 mM MgCl2 , 1 mM CaCl2 , 5 mM HEPES, pH 7.55. The pH should be adjusted to the final value with NaOH. The Ringer’s solution can be kept well sealed at room temperature for months (see Note 1). 4. Stock glucose solution (1 M). This solution has to be kept at −20◦ C to avoid bacterial growth. 5. Dissecting microscope. 6. Infrared light source. It can be a homemade infrared-LEDbased system. Alternatively, an infrared safelight (FJW Optical Systems, Inc.) can be used. 7. Two infrared image viewers attached to the dissecting microscope eyepieces. The viewers are the FINDR-SCOPE Infrared Viewer Model 84499A (FJW Optical Systems, Inc.). The same company also provides the components necessary for attaching the viewers to the eyepieces.

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8. An infrared viewer with illuminator: An option is FIND-RSCOPE Infrared Viewer with Illuminator Model 85100A (FJW Optical Systems, Inc.). 9. Petri dishes: 35 mm Falcon (Fisher Scientific). 10. Plastic transfer pipettes: 5 ml (Fisher Scientific). 11. Dissecting tools (Fine Science Tools or Roboz Surgical Instruments): One pair of delicate iris scissors, straight, 11.5 cm long. One pair of extra fine Bonn scissors, curved, 8.5 cm long. At least two pairs of fine Dumont forceps, numbers 5 or 7. A couple of pairs of inexpensive student Dumont forceps (number 5) are also useful during the dissection. One blade holder/breaker. Pithing needles. 12. Sylgard 184 elastomer kit (Essex, Charlotte, NC). 13. Metal cutter (local hardware store or Fisher Scientific). 14. Double-edge razor blades, “Personna Double Edge Platinum Chrome” (Wal-Mart). 15. Experimental chambers: These can be 35-mm culture dishes with a 12-mm chamber (Warner Instruments, Hamden, CT). 16. Coating solution for chambers: 0.01% solution of poly-Lornithine or 0.1% solution of poly-L-lysine (Sigma-Aldrich Chemical Company, St. Louis, MO). The poly-L-lysine solution is diluted with distilled water to a final 0.01% concentration. 17. Three light-tight boxes that can accommodate 2–3 of 35-mm Petri dishes each. 18. Fiber optic illuminator and longpass (>530 nm) filter for bleaching the cells (Edmund Optics, Barrington, NJ).

2.2. FRAP Measurement

1. Non-linear optical confocal microscope with a Ti–sapphire tunable infrared laser. The software running the system typically includes a program for setting up the parameters for the FRAP experiment. There are several such systems available, for example, the one based on the Zeiss LSM 510 (Carl Zeiss, Thornwood, NY). Make sure that the experimental chambers fit on the microscope stage. A special stage accessory that is usually available from the microscope manufacturer might be necessary. 2. High numerical aperture objective lens. For a system based on an upright microscope, use the 63× water immersion lens (NA = 0.9). For a system with an inverted microscope, you could use either the 40× or the 63× oil-immersion lenses.

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3. Methods 3.1. Photoreceptor Cell Preparation 3.1.1. Dishes, Chambers, and Razor Blades

1. Coat the bottoms of 35-mm Falcon Petri dishes with Sylgard elastomer. Prepare the elastomer according to the instructions on the box and pour a small amount in each dish to cover its bottom with a thick layer. Replace the covers on the dishes and store them. The elastomer will harden over a period of few days and the dishes will be ready. 2. Coat the bottoms of the experimental chambers with 0.01% poly-L-lysine or poly-L-ornithine; 200 μl of solution per chamber is enough. Cover the chambers with a paper towel to protect them from dust and let them sit until dry. Wash them with distilled water and keep them upside down to dry. Store in a closed box and use within 2 weeks. 3. The chambers can be re-used. After an experiment, wash the chamber with 100% ethanol to remove oil on the outside (from the oil-immersion lens) and the cell debris on the inside. Use cotton-tipped applicators to gently scrub the bottom of the chamber to remove the debris. Wash with distilled water and let dry. 4. Prepare several small razor blades by cutting each doubleedged blade into eight pieces with the metal cutter.

3.1.2. Isolated Retinas

1. Keep the animals healthy and clean, feed them, and provide veterinary care (see Note 2). 2. Dark adapt an animal in a ventilated container (for example, a suitably modified bucket) in the dark room for at least 2–3 h before beginning experiments. 3. Immediately before the experiment, add 0.5 ml of the glucose stock to 100 ml of the Ringer’s (final glucose concentration of 5 mM). Use this Ringer’s for experiments. Discard the leftover solution at the end of the day, as it might grow bacteria. 4. Pour some of the Ringer’s solution to two 35-mm Petri dishes and keep them close to the dissecting microscope. 5. Kill the animal under dim red light by pithing the brain and the spinal cord with the Pithing needles. 6. Enucleate the eyes using the long scissors and the student Dumont forceps. 7. The rest of the procedures are carried out under the dissecting microscope using infrared light. Use the infrared

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viewer with illuminator, in case you need to find something outside the field of view of the microscope. 8. Remove any leftover muscle and skin tissue using the long scissors and the student Dumont forceps. 9. Remove the anterior part of the eye, leaving the vitreous behind; use the short scissors to make an incision and cut around just behind the ora serrata. 10. Transfer the eyecup into one of the Petri dishes filled with Ringer’s. Carefully remove the vitreous using the fine forceps. 11. Under the infrared light, the retina is now visible against the dark background of the retinal pigment epithelium. Gently separate the retina from the epithelium; it will remain attached to the eyecup at the optic nerve. With the fine forceps reach underneath the retina and pinch it off at the point of attachment. Separate the retina fully from the eyecup (see Note 3). 12. Using a plastic pipette, draw some solution containing the retina and transfer it to the other, clean Petri dish. It can be kept there in a light-tight box for a few hours (see Note 4). 3.1.3. Isolated Living Photoreceptor Cells

1. Bring pipettes, coated chambers, sylgard-covered dishes close to the dissecting microscope. Grab a piece of razor blade with the blade holder, with the edge of the blade at approximately 45◦ angle to the holder. All subsequent procedures are carried out under the dissecting microscope using infrared light. 2. Using the small scissors, cut the retina into four pieces. With a plastic pipette, draw some solution containing a piece and transfer it into a sylgard-covered dish. The final volume of the solution in that dish should be about 600 μl. 3. With the fine forceps flatten the piece of retina on the sylgard surface, keeping the photoreceptor side up. Using the razor blade, chop the piece in one direction; repeat 3–4 times, then rotate the dish 90◦ , and chop again 3–4 times. Repeat the whole procedure until you see a “cloud” of dissociated cells. It is important to chop finely, while keeping the piece of retina stuck to the sylgard. If the chopping is too coarse, or the retina becomes unstuck, one gets mostly pieces of retina instead of isolated cells (see Note 5). 4. After finishing the chopping, transfer the solution to three experimental chambers, 200 μl in each chamber see Note 6). Keep the chambers with the isolated cells in a light-tight box. 5. Wait for 10 min for the cells to settle, then add 2–3 ml of Ringer’s to each of the dishes that contain the chambers.

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The cells can now be taken to the non-linear optical confocal microscope for experiments. Isolated cells can survive for a few hours (see Note 7). 3.2. FRAP Measurements

1. Tune the Ti:sapphire laser to 720 nm for fluorescence excitation. Set the fluorescence emission measurement from 400 to 650 nm. 2. Take one of the chambers containing cells out of the light– tight box and expose the cells to >530 nm light for 1 min, using the illuminator and the longpass filter. Carry out measurements between 1 and 2 h after bleaching. 3. Find a cell under the bright field. Only rod outer segments with attached ellipsoids can generate retinol. Among those, it is best to use whole intact cells (with outer segment, ellipsoid, and nucleus) instead of ROS-RIS, as the latter might not survive through the full course of a FRAP experiment (see Note 8). 4. Carry out preliminary measurements to optimize the measuring and bleaching intensities. Along with the intensities, you need to optimize the number of time points, time delays between scans, and the time for bleaching. For FRAP experiments, a high intensity of the laser beam is used to bleach retinol, and a lower, non-bleaching intensity is used for scanning and measuring the fluorescence before and after bleaching. Select intensities and number of scans according to the following criteria: (a) avoid bleaching of fluorescence during scanning, (b) avoid visible cell damage or death (due to phototoxicity), and (c) obtain measurements of sufficient resolution to determine the kinetics of retinol fluorescence recovery (see Note 9).

3.2.1. Measurement of Lateral Diffusion

1. Select an intact rod cell. Ensure that the whole cell is included in the scanning area, but keep the area small to minimize the time required for frame acquisition. Set up the FRAP parameters and select the area for bleaching. This area should be a rectangular area, covering one half of the outer segment, on one side of the long axis (Fig. 6.2). It is critical that the inside edge of the area is the long axis of the outer segment. 2. Carry out the experiment. The fluorescence should redistribute and equilibrate between the bleached and unbleached halves over a period of 10–20 s. 3. For each time point after the bleach, measure the fluorescence in a region of interest that covers the unbleached half of the outer segment – opposite to the bleached side.

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4. The fluorescence of retinol declines in this area after bleaching. Fit this decline with a single exponential function (for example, with a graphics software program like Kaleidagraph) and obtain the rate of decline k (Fig. 6.2 g). 5. Dlateral , the coefficient for diffusion of retinol in the plane of the disk, is given as Dlateral = d 2 ×

k π2

[1]

where d is the diameter of the outer segment. For the cell in Fig. 6.2, d = 7.2 μm and k = 0. 25 s−1 , giving Dlateral = 1.3 μm2 s−1 (10).

Fig. 6.2. Measurement of the lateral mobility of all-trans-retinol in the outer segment of an isolated intact frog rod with two-photon FRAP. (a) Diagram of the cell: ROS, rod outer segment; ell, ellipsoid; the bleached area is shaded. (b) Initial image acquired before retinol bleaching, (c–e) images acquired immediately, 2285 ms and 6092 ms after bleaching, respectively. (f) Fluorescence profiles along an outer segment diameter in the bleached region from images (b) (curve 1), (c) (curve 2), (d) (curve 3), and (e) (curve 4). (g) Kinetics of fluorescence recovery in the bleached (•) and unbleached disk areas (). The lines represent single exponential fits, with rate constants of 0.75 s−1 for the bleached area and 0.25 s−1 for the unbleached area. The fluorescence recovery in the bleached disk areas is due to retinol movement from the unbleached areas above and below the plane of bleaching, as well as to retinol movement from the areas in the left side. Images (b–e) are shown at the same intensity scaling. Scale bar is 10 μm. Reprinted from (10) with permission.

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3.2.2. Measurement of Axial Diffusion

1. Select an intact rod cell. Set up the FRAP parameters and select the area for bleaching. This area should be a rectangular area, covering the top half of the outer segment (Fig. 6.3). It is critical that the inside edge of the area is at the half point between the base and tip of the outer segment. 2. Carry out the experiment. For long outer segments, it is unlikely that the fluorescence will equilibrate fully between the bleached and unbleached halves within a reasonable time. 3. Measure the values of fluorescence in the unbleached (F1 ) and bleached (F2 ) halves immediately after bleaching. Calculate F = F1 − F2 . For the cell in Fig. 6.3, F = 50.

Fig. 6.3. Measurement of the axial mobility of all-trans-retinol in the outer segment of an intact frog rod. (a) Diagram of the cell: ROS, rod outer segment; ell, ellipsoid; the bleached area is shaded. (b) Initial image acquired before retinol bleaching, (c, d) images acquired immediately, and 1440 s after bleaching. (e) Fluorescence intensity profiles before (thin line) and immediately after (thick line) bleaching. (f) Fluorescence intensity profile 1440 s after bleaching; the straight dashed line represents the slope of the intensity profile at the boundary between bleached and unbleached regions. The slope of the intensity profile, S = −2.4 μm−1 , gave a rate constant α = 6.9 × 10−5 s−1 . Images (b–d) are shown at the same intensity scaling. The fluorescence intensity profiles are aligned with the images. Reprinted from (10) with permission.

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4. At a time point 20–30 min after the bleach, measure and plot the profile of fluorescence along the length of the outer segment. Measure the slope of the fluorescence S at the middle of the outer segment (Fig. 6.3f). For the cell in Fig. 6.3, S = −2.4 μm−1 . 5. Obtain the rate parameter α by solving the equation (for example, with Mathcad): ∞ 2 × F  −(2m+1)2 αt · e S=− L

[2]

m=0

where L is the length of the outer segment. 6. Daxial , the coefficient for diffusion of retinol along the length of the outer segment, is given as Daxial = L 2 ×

α π2

[3]

where L is the length of the outer segment. For the cell in Fig. 6.3, L = 57 μm, and a = 6.9 × 10−5 s−1 , giving Daxial = 0.023 μm2 s−1 (10).

4. Notes 1. It is critical that the buffer’s composition is accurate within a few percent, as cells are sensitive to the osmolarity of the solution. 2. The health of the cells depends on the health of the animals. Do your best to ensure the health of the animals. 3. Sometimes it is difficult to separate the retina from the pigment epithelium. Be patient and slowly peel off starting from the periphery. If you still cannot separate the retina, a likely possibility is incomplete dark adaptation, which could be caused by bright red lights as well. Ensure that the animal is dark adapted properly and dim the red lights. 4. It is a good idea to cut a small piece of retina and transfer it to a separate Petri dish. Take a look at this piece of retina under room lights: it should be a bright red color. This red color is due to rhodopsin and should fade rapidly under room lights. The presence of rhodopsin indicates the presence of rod outer segments and confirms that you have obtained a healthy retina. The lack of red color is indicative of either an unhealthy retina or a failure to separate the rod outer segments from the retinal pigment epithelium. In such

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case, you should ensure the health of the animals and proper dark adaptation (see Note 3). 5. This is a critical step. If you have a healthy retina (see Note 4), failure to obtain intact cells is most likely due to improper chopping. If the chopping is too coarse, you will see large chunks of retina in the dish when you check under bright field. Release of a “cloud” of cells is usually an indication of successful chopping. 6. The density of the photoreceptor cells in the experimental chamber is important. A very high density will result in cells settling on top of each other, which will not allow an experiment. A very low density might result in failure to find cells for experiment. Optimize the density so that you can find cells suitable for experiments on a regular basis. 7. Before you embark on actual experiments, you need to ensure that the chopping (Note 5) and the cell density (Note 6) have been optimized. Check your isolated cell preparations under the bright field of the microscope and adjust chopping and density until you can regularly obtain isolated intact cells that have settled without cells above or below them. For your experiments it is best to use whole intact cells (with outer segment, ellipsoid, and nucleus). 8. For preliminary experiments for the measurement of lateral diffusion, ROS-RIS can be used. 9. To optimize parameters, you should start with the determination of a range of scanning parameters that do not result in the bleaching of retinol fluorescence. Keep the same frame format (number of pixels) and zoom settings. Begin with a laser intensity that is high enough to obtain a clear retinol fluorescence signal. Scan the cell 10 times – without a time delay – and measure the rod outer segment retinol fluorescence for each of the 10 frames that you have acquired. The change in retinol fluorescence reflects the bleaching due to scanning. If the bleaching is less than ∼0.5% per scan, you can increase the laser intensity to obtain a better signal-tonoise ratio; if not, you need to lower the laser intensity. After establishing a range of acceptable scanning intensities, proceed to the determination of the bleaching intensity for the FRAP experiment. Select an area for bleaching and begin with a high enough laser intensity so that the effect of bleaching can be resolved. You might need to expose the selected area repeatedly to the high laser intensity to achieve sufficient bleaching. If bleaching causes visible cell damage, reduce the laser intensity or the number of repetitions. Optimize intensity and repetitions to obtain resolvable bleaching without visible cell damage. After establishing a range of

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acceptable parameters for bleaching, you can then adjust the time delay between the measurement scans. If you find that you need to shorten the time delay between measurement scans, you might need to reduce the number of bleaching repetitions. If you need to reduce the acquisition time for each frame, you will need to reduce the number of pixels per frame. References 1. Imanishi, Y., Lodowski, K.H., Koutalos, Y. (2007) Two-photon microscopy: Shedding light on the chemistry of vision. Biochemistry 46, 9674–9684. 2. Lamb, T.D., Pugh, E.N., Jr. (2004) Dark adaptation and the retinoid cycle of vision. Prog. Retin. Eye Res. 23, 307–380. 3. Saari, J.C. (2000) Biochemistry of visual pigment regeneration: The Friedenwald lecture. Invest. Ophthalmol. Vis. Sci. 41, 337–348. 4. Szuts, E.Z., Harosi, F.I. (1991) Solubility of retinoids in water. Arch. Biochem. Biophys. 287, 297–304. 5. Moise, A.R., Noy, N., Palczewski, K., Blaner, W.S. (2007) Delivery of retinoid-based therapies to target tissues. Biochemistry 46, 4449–4458. 6. Ala-Laurila, P., Kolesnikov, A.V., Crouch, R.K., Tsina, E., Shukolyukov, S.A., Govardovskii, V.I., Koutalos, Y., Wiggert, B., Estevez, M.E., Cornwall, M.C. (2006) Visual cycle: Dependence of retinol production and removal on photoproduct decay and cell morphology. J. Gen. Physiol. 128, 153–169. 7. Chen, C., Tsina, E., Cornwall, M.C., Crouch, R.K., Vijayaraghavan, S., Koutalos, Y. (2005) Reduction of all-trans retinal to alltrans retinol in the outer segments of frog and mouse rod photoreceptors. Biophys. J. 88, 2278–2287.

8. Tsina, E., Chen, C., Koutalos, Y., AlaLaurila, P., Tsacopoulos, M., Wiggert, B., Crouch, R.K., Cornwall, M.C. (2004) Physiological and microfluorometric studies of reduction and clearance of retinal in bleached rod photoreceptors. J. Gen. Physiol. 124, 429–443. 9. Wu, Q., Blakeley, L.R., Cornwall, M.C., Crouch, R.K., Wiggert, B.N., Koutalos, Y. (2007) Interphotoreceptor retinoid-binding protein is the physiologically relevant carrier that removes retinol from rod photoreceptor outer segments. Biochemistry 46, 8669–8679. 10. Wu, Q., Chen, C., Koutalos, Y. (2006) Alltrans retinol in rod photoreceptor outer segments moves unrestrictedly by passive diffusion. Biophys. J. 91, 4678–4689. 11. Zipfel, W.R., Williams, R.M., Christie, R., Nikitin, A.Y., Hyman, B.T., Webb, W.W. (2003) Live tissue intrinsic emission microscopy using multiphoton-excited native fluorescence and second harmonic generation. Proc. Natl. Acad. Sci. USA 100, 7075–7080. 12. Brown, E.B., Wu, E.S., Zipfel, W., Webb, W.W. (1999) Measurement of molecular diffusion in solution by multiphoton fluorescence photobleaching recovery. Biophys. J. 77, 2837–2849.

Chapter 7 Microfluorometric Measurement of the Formation of All-Trans-Retinol in the Outer Segments of Single Isolated Vertebrate Photoreceptors Yiannis Koutalos and M. Carter Cornwall Abstract The first step in the detection of light by vertebrate photoreceptors is the photoisomerization of the retinyl chromophore of their visual pigment from 11-cis to the all-trans configuration. This initial reaction leads not only to an activated form of the visual pigment, meta II, that initiates reactions of the visual transduction cascade but also to the photochemical destruction of the visual pigment. By a series of reactions termed the visual cycle, native visual pigment is regenerated. These coordinated reactions take place in the photoreceptors themselves as well as the adjacent pigment epithelium and Müller cells. The critical initial steps in the visual cycle are the release of all-trans-retinal from the photoactivated pigment and its reduction to all-trans-retinol. The goal of this monograph is to describe methods of fluorescence imaging that allow the measurement of changes in the concentration of all-trans-retinol as it is reduced from all-trans-retinal in isolated intact salamander and mouse photoreceptors. The kinetics of all-transretinol formation depend on cellular factors that include the visual pigment and photoreceptor cell type, as well as the cytoarchitecture of outer segments. In general, all-trans-retinol forms much faster in cone cells than in rods. Key words: Retina, rod, cone, visual pigment, rhodopsin, visual cycle.

1. Introduction Absorption of incoming light by the visual pigment of vertebrate photoreceptors isomerizes its retinyl chromophore from 11cis to all-trans. This photoisomerization results in the formation of meta II, an enzymatically active visual pigment conformation, and begins the transduction of light to an electrical signal that can be transmitted to the brain (see Ref. (1) for review). The all-trans chromophore is then removed and recycled to reform 11-cis-retinal that can be used to regenerate the pigment (see Ref. H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_7, © Springer Science+Business Media, LLC 2010

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(2–5) for reviews). The first steps of this process take place in the photoreceptor outer segment and culminate in the generation of all-trans-retinol, which is formed through the reduction of the all-trans-retinal released from the photoactivated visual pigment. The conversion of all-trans-retinal to all-trans-retinol is catalyzed by the enzyme retinol dehydrogenase (RDH) and requires metabolic input in the form of NADPH. In the case of rods, all-trans-retinol is transferred to neighboring cells of the retinal pigment epithelium (RPE) where it is esterified by lecithin retinyl acyltransferase. The resultant ester is the substrate for the Rpe65 isomerohydrolase that generates 11-cis-retinol, which is then oxidized to 11-cis-retinal and translocated to the photoreceptor outer segments; there, it condenses with opsin left behind following the release of all-trans-retinal to regenerate the visual pigment. In the case of cones, an additional pathway involving the Müller cells can generate 11-cis-retinal and regenerate the visual pigment independently of the RPE (6, 7). The transfer of alltrans-retinol from the outer segments appears to occur through mass action (8–11) and allows for the recycling of the chromophore to make fresh 11-cis-retinal. In the rest of the chapter, where unspecified, retinal and retinol refer to their all-trans isomers. Several steps comprise the pathway that forms all-transretinol in the outer segment after the photoisomerization of the visual pigment’s 11-cis-retinyl chromophore. The initial step is the hydrolysis of the Schiff base bond between the chromophore and the visual pigment protein and the release of all-trans-retinal. The released all-trans-retinal may be sequestered inside the disks in the form of a Schiff base with phosphatidylethanolamine; it can then become available for reduction after it has been transferred to the cytosol by the ABCA4 transporter (12). The final step is the reduction of all-trans-retinal by RDH, a reaction that uses NADPH as a co-factor. Because of the substantial amount of chromophore present in photoreceptor outer segments (3–4 mmol/l) (13), the kinetics of RDH and the availability of NADPH may have a significant impact on the overall kinetics of retinol formation. In summary, the kinetics of retinol formation depend on the rate of retinal release, local NADPH availability, retinal access to RDH, RDH kinetics, and retinol elimination rate (that depends on outer segment morphology). The kinetics of the formation of retinol after light excitation can be monitored in the outer segments of single photoreceptors from its intrinsic fluorescence, which has been shown to provide a good measure of its concentration (9, 14). Such single-cell fluorescence measurements with their high sensitivity and time resolution provide a unique window for examining the different steps in the formation of retinol. They have been used to study the formation of retinol in the rod and cone photoreceptors from several vertebrate species, including tiger salamander (Ambystoma

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tigrinum), grass frog (Rana pipiens), lizard (Gecko gecko), and mouse (Mus musculus) (8–10, 14–16). From among these species, tiger salamanders and mice offer several distinct advantages as model systems for the study of retinol formation. From the salamander retina, one can obtain different types of rods and cones that include two types of rods – green sensitive and blue sensitive – and three types of cones – red sensitive , blue sensitive, and UV sensitive (10, 17). These salamander cells are large, robust, and can survive for several hours after isolation from the retina, facilitating the experimental manipulations for single-cell imaging. Their large size also allows measurement of the time course and kinetics of retinol formation in local regions of the outer segments. Salamander photoreceptors utilize different pigment types as well as the same pigment in different cell types (10, 18). Thus, they allow the comprehensive examination of the dependence of retinol formation on cell type, visual pigment type, and outer segment architecture. The mouse retina offers complementary advantages. It is dominated by a single-cell type, the rods, allowing comparisons with biochemical measurements from isolated retinas (14) and the eyes of whole animals. Furthermore, the availability of genetically modified animals offers the opportunity to probe specific enzyme involvement and disease relevance. Figure 7.1 shows a diagram of the setup that is used for such measurements, and includes an inverted microscope, fitted with a near UV fluorescence excitation light source and a high-sensitivity

Fig. 7.1. Diagram of the experimental apparatus. Bright-field infrared image of a rod and a cone photoreceptor is shown at the top and a fluorescent image of these cells is shown at the bottom. The fluorescent image was acquired before visual pigment bleaching.

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camera. The orientation and function of these separate components are described later (Section 3) and in the figure legend. An experiment begins with placement of dark-adapted photoreceptor cells in the experimental chamber. After an initial fluorescence measurement in their dark-adapted condition, the cells are illuminated to ensure that virtually all of the visual pigment chromophore has been isomerized to the all-trans conformation. The reason behind this is that the UV light used to excite the fluorescence of retinol is also absorbed by the visual pigment and thus isomerizes the 11-cis chromophore. So, to avoid having each measurement of fluorescence initiating the reactions that lead to additional retinol formation, each experiment begins with photoactivation (bleaching) of virtually all of the visual pigment. Figure 7.2 shows an experiment performed on an isolated salamander green-sensitive rod photoreceptor in this way. Here it can be seen that, following quantitative bleaching of the visual pigment, retinol fluorescence increases within the outer segment, assuming a maximal value at about 30 min, and declining slowly thereafter. The data presented in Fig. 7.3 illustrate a similar experiment performed on an isolated salamander red-sensitive cone cell. In this case, retinol fluorescence increases rapidly after bleaching, reaching a maximal value after ca. 1 min. Subsequently, it declines at a much faster rate than in the rod. Finally, Fig. 7.4 presents an experiment with an isolated mouse rod, performed at 37◦ C.

Fig. 7.2. Formation of all-trans-retinol in an isolated rod photoreceptor from a larval tiger salamander retina. (a) Retinol fluorescence increases in the rod outer segment after rhodopsin bleaching. a, Infrared image of an isolated rod photoreceptor cell; b–g, fluorescence (excitation, 360 nm; emission >420 nm) images of the cell; b, before bleaching; c, immediately after; d, 10 min; e, 30 min; f, 50 min; g, 90 min after bleaching. All fluorescence images are shown at the same intensity scaling. Bar is 10 μm. (b) Rod outer segment fluorescence intensity values before (t = −1 min) and at different times after rhodopsin bleaching for the cell in (a). Bleaching was carried out from t = −1 to 0 min.

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Fig. 7.3. Formation of all-trans-retinol in an isolated cone photoreceptor from a larval tiger salamander retina. (a) Retinol fluorescence increases in the cone outer segment after visual pigment bleaching. a, Infrared image of the isolated cone photoreceptor cell; b–g, fluorescence (excitation, 360 nm; emission, >420 nm) images of the cell; b, before bleaching; c, immediately after; d, 0.5 min; e, 1 min; f, 5 min; g, 30 min after bleaching. All fluorescence images are shown at the same intensity scaling. Bar is 10 μm. (b) Cone outer segment fluorescence intensity values before (t = −10 s) and at different times after visual pigment bleaching for the cell in (a). Bleaching was carried out from t = −10 s to 0 min. The inset shows the same data as the main panel in an expanded time scale.

Fig. 7.4. Formation of all-trans-retinol in an isolated rod photoreceptor from a c57bl/6 mouse. (a) Retinol fluorescence increases in the rod outer segment after rhodopsin bleaching. a, Infrared image of an isolated rod photoreceptor with intact ellipsoid; b–g, fluorescence (excitation, 360 nm; emission, >420 nm) images of the cell; b, before bleaching; c, immediately after; d, 10 min; e, 30 min; f, 50 min; g, 90 min after bleaching. All fluorescence images are shown at the same intensity scaling. Experiment carried out at 37◦ C. Bar is 5 μm. (b) Rod outer segment fluorescence intensity values before (t = −1 min) and at different times after rhodopsin bleaching for the cell in (a). Bleaching was carried out from t = −1 to 0 min.

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2. Materials A dark room is necessary for the fluorescence imaging setup. The same room can be used for dark-adapting animals and for dissection. An area of ∼100–150 ft2 is sufficient. A revolving door for entering is convenient, but a thick black curtain is also adequate. 2.1. Photoreceptor Cell Preparation

1. Red lights for the dark room: These are obtained from photographic equipment stores. A good choice is adjustable Kodak safelights fitted with Kodak Wratten #2 filters. If individual red bulbs are used, an appropriate choice is the Delta 1 Jr. Safelight. It is best to keep the red lights as dim as possible. 2. Larval tiger salamanders (A. tigrinum) are obtained from approved vendors (The Sullivan Company, Nashville, TN; Kons Scientific, Germantown, WI). Salamanders are usually available throughout the year; however, check with the supplier well in advance for availability. 3. Wild-type mice (M. musculus) are obtained from approved vendors (The Jackson Laboratory, Bar Harbor, ME; Harlan Laboratories, Indianapolis, IN). Genetically modified animals can be obtained from appropriate sources. 4. Salamander Ringer’s with composition: 110 mM NaCl, 2.5 mM KCl, 1.6 mM MgCl2 , 1 mM CaCl2 , 5 mM HEPES, pH 7.55. The pH should be adjusted to the final value with NaOH (see Note 1). The Ringer’s solution can be kept well sealed at room temperature for months. At the time of the experiment, glucose (5 mM) (see Note 2) and delipidated bovine serum albumin (concentration 0.01%) are added (see Note 3). 5. Mammalian Ringer’s with composition: 130 mM NaCl, 5 mM KCl, 0.5 mM MgCl2 , 2 mM CaCl2 , 25 mM hemisodium–HEPES, pH 7.40. At the time of the experiment, glucose (5 mM) is added. 6. Stock glucose solution (1 M): This solution has to be kept at −20◦ C to avoid bacterial growth. 7. Dissecting microscope. 8. Infrared (IR) light source: This can be a homemade infrared-LED-based system. Alternatively, an infrared safelight (FJW Optical Systems, Inc.) can be used. 9. Two infrared image viewers are attached to the dissecting microscope eyepieces (FJW Optical Systems, Inc.).

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The same company provides the components necessary for attaching the viewers to the dissecting microscope eyepieces. 10. Infrared viewer with illuminator (FIND-R-SCOPE Infrared Viewer with Illuminator Model 85100A; FJW Optical Systems, Inc.). 11. Petri dishes: 35 mm plastic (Falcon) dishes (Fisher Scientific). 12. Plastic transfer pipettes: 5 ml (Fisher Scientific). 13. Filter paper (Wratten; Fisher Scientific). 14. Dissecting tools (Fine Science Tools or Roboz Surgical Instruments): One pair of delicate iris scissors, straight, 11.5 cm long. One pair of extra fine Bonn scissors, curved, 8.5 cm long. At least two pairs of fine Dumont forceps, numbers 5 or 7. A couple of pairs of inexpensive student Dumont forceps (number 5) are also useful during the dissection. One razor blade holder/breaker. Pithing needles. 15. Sylgard 184 elastomer kit (Essex, Charlotte, NC). 16. Metal cutter (local hardware store or Fisher Scientific). 17. Double-edge razor blades, “Personna Double Edge Platinum Chrome” (local drug stores or pharmacies). 18. Experimental chambers: These can be 35-mm culture dishes with a 12-mm chamber (Warner Instruments, Hamden, CT). 19. Coating solution for chambers: 0.01% Poly-L-ornithine solution or 0.1% poly-L-lysine solution (Sigma-Aldrich Chemical Company, St. Louis, MO). The poly-L-lysine solution is diluted with distilled water to a final 0.01% concentration. 20. Three light-tight boxes that can accommodate 2–3 of 35-mm Petri dishes each. 21. Fiber-optic illuminator and longpass (>530 nm) filter for bleaching the cells (Edmund Optics, Barrington, NJ) (see Note 4). 2.2. Fluorescence Imaging

1. An inverted microscope with a light train that can be used for epifluorescence measurements. An inverted microscope is best, as it allows the use of high numerical aperture oilimmersion objectives that are critical for fluorescence measurements from single cells. The microscope should have a port for the CCD camera, and the microscope optics should allow a setting for 100% of the fluorescence signal to be directed to the camera port.

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2. Table for the microscope: A 3 ft × 3 ft vibration isolation table (Newport, Irvine, CA, or TMC, Peabody, MA), but a solid table of similar size is sufficient. The height of the table should allow the legs of an experimenter sitting in a chair to go under it. 3. The microscope should be placed in a light-tight enclosure (a “cage”) that allows access to microscope controls for the experimenter. A solid, sturdy frame that can support the weight of the cage is essential and can be constructed from wood or metal. The frame should rest on the floor and reach a height a few inches above the top of the microscope; it should have a couple of inches clearance from the sides of the table to allow bringing in optical and electronic cables. The left and right sides, top, and back of the cage can be constructed from aluminum plates screwed on or nailed to the frame and should begin 1–2 in. below the surface of the microscope table (see Note 5); the inside surface of these plates should be painted black (see Note 6). The front of the cage should be left open and covered with a double curtain made from black cloth. The inside curtain can be thin and should have two slits to allow the hands of an experimenter to control the microscope. A wooden horizontal rod attached at the bottom helps to roll the curtain up or down and keep it in place during experiments (see Note 7). The outside curtain should be thick and drop to 1–2 in. below the surface of the microscope table (see Note 8). If a vibration isolation table is used, a hand rest attached to the frame and placed about 1 in. above the height of the microscope table is very helpful (see Note 9). 4. Microscope stage: It should include an adaptor in which the experimental chambers readily fit. Such an adaptor is usually available from the microscope manufacturer, as the size of the experimental chambers is fairly standard. 5. Perfusion components: Slow perfusion of the cells during the course of the experiment improves their viability. A gravity-fed flow rate of 0.1–0.5 ml/min is appropriate; higher rates might disturb or even dislodge the cells (see Note 10). The solution is continuously removed from the chamber and poured into a beaker below the microscope stage through a passive, wick-facilitated system. Alternatively, the solution can be removed through suction. In this case, use a 14-gauge needle connected to the vacuum line through a 3-ml syringe and plastic tubing to remove the solution from the chamber (see Note 11). 6. Infrared light source for microscope: An infrared filter (>850 nm) in front of the microscope’s transmitted light

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source can be used to provide the infrared light for viewing the dark-adapted cells on the microscope stage. The darkadapted cells on the microscope stage have to be protected from any visible light leak from this source. One possible arrangement is for the light source to be placed outside the cage and the light be brought in via a light guide. Another option is to use an infrared-LED (wavelength >850 nm) placed behind the condenser. In either case, it is necessary to be able to easily switch the infrared light on and off. 7. Objective lens: A high-magnification (at least 40×) and numerical aperture (at least 1.3) oil-immersion lens is necessary. Use a lens appropriate for epifluorescence measurements (see Note 12). 8. Lens immersion oil: this should be of high quality and have low intrinsic fluorescence. A suitable one is Cargille Type FF Nonfluorescing immersion oil (Fisher Scientific). 9. Lens cleaning paper and solution (Fisher Scientific). 10. Stage, solution, and objective lens heating: Experiments with mouse photoreceptors have to be carried out at 37◦ C, necessitating heating the experimental chamber, the incoming solution, and the objective lens. A variety of heated stages compatible with the experimental chamber are available (Warner Instruments), along with a separate assembly for heating the incoming solution, heated jacket for the objective lens (see Note 13), and temperature controllers for each component. The temperature of the solution in the experimental chamber is measured independently and is maintained at 37◦ C by adjusting the temperatures of the stage, the incoming solution, and the lens. 11. Excitation light source: A xenon continuous arc light source (Sutter Instrument Company, Novato, CA; or Cairn Research Ltd., Faversham, UK) is used to provide the light to excite the fluorescence of retinol. An electronic shutter (Uniblitz; Vincent Associates, Rochester, NY) is placed in the light path to control the exposure of the cells to the excitation light (see Note 14). The xenon light source is placed on the floor under or next to the microscope table (see Note 15) and the excitation light is brought to the microscope port with a light guide. Adaptors for connecting the light guide to the light source and the microscope port are available (Sutter Instrument Company). The casing for the light source should have slots for neutral density filters to attenuate the excitation light intensity as necessary.

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12. Neutral density filters and holders: The filter holders should fit into the slots in front of the excitation light source. Neutral density filters of 1-log unit (attenuation to 10%) and 2log unit (attenuation to 1%) are necessary (Chroma Technology, Rockingham, VT). 13. Filters for retinol fluorescence measurement: filter set 11000v3 (Chroma Technology) includes parts D350/50× for excitation (bandpass filter centered at 350 nm with bandwidth of 50 nm), 400DCLP for dichroic mirror (reflects light <400 nm), and E420LPv2 for emission (longpass filter >420 nm) (see Note 16). 14. High-sensitivity CCD camera: Adequate sensitivity of the CCD camera is critical for successful imaging of retinol fluorescence in living cells. Suitable cameras include CoolSNAP HQ2 Monochrome (Photometrics, Surrey, BC), Sensicam QE (Cooke Corporation, Auburn Hills, MI), or Orca-285 (Hamamatsu Photonics, Hamamatsu-City, Japan). An image intensifier (VS4-1845; OPELCO Inc., Dulles, VA) coupled to the camera can further improve detection but is expensive. The same camera is used in live mode with infrared illumination to find a dark-adapted cell to begin an experiment. 15. Image acquisition and analysis software: The software coordinates the function of the different hardware components to acquire images with user-specified parameters (for example, exposure time). Suitable packages include Slidebook (Intelligent Imaging Innovations, Denver, CO) and Openlab (Improvision Inc., Waltham, MA). The software package includes routines for image analysis. 16. Computer: The image acquisition and analysis software is installed in a computer that controls all hardware components. Consult with the software provider for appropriate computer specifications. 17. Computer monitor: Larger monitors are easier for the user, so the size of the monitor ultimately depends on the available space next to the microscope table. During an experiment, the screen should be covered with transparent red plastic (gel sheet Roscolux #27, Medium Red; theater lighting companies) to minimize light in the room. 18. Surge protectors for all electrical and electronic components. It is essential that the xenon arc light source has its own separate surge protector. Other components can share a surge protector.

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3. Methods 3.1. Photoreceptor Cell Preparation 3.1.1. Dishes, Chambers, and Razor Blades

1. Coat the bottoms of 35-mm Falcon Petri dishes with Sylgard elastomer. Prepare the elastomer according to the instructions on the box and pour a small amount into each dish to cover its bottom with a thick layer. Replace the covers on the dishes and store them. The elastomer will harden over a period of few days and the dishes will be ready. 2. Coat the bottoms of the experimental chambers with 0.01% poly-L-lysine or poly-L-ornithine; 200 μl of solution per chamber is enough. Cover the chambers with a paper towel to protect them from dust and let them sit until dry. Wash them with distilled water and keep them upside down to dry. Store in a closed box and use within 2 weeks. 3. The chambers can be re-used. After an experiment, wash the chamber with 100% ethanol to remove the oil (from the oilimmersion lens) on the outside and the cell debris on the inside of the chamber. Use cotton-tipped applicators to gently scrub the bottom of the chamber to remove the debris. Wash with distilled water and let dry. 4. Prepare several small razor blades by cutting each doubleedged blade into eight pieces with the metal cutter.

3.1.2. Isolated Retinas

1. Keep the animals healthy and clean, feed them, and provide veterinary care (see Note 17). 2. Dark adapt an animal in a ventilated container (for example, for salamanders, a suitably modified bucket) in the dark room for at least 2–3 h before beginning experiments. 3. Immediately before the experiment add appropriate glucose and/or bovine serum albumin concentrations. Use this Ringer’s for experiments. Discard any leftover solution at the end of the day, as it might grow bacteria. 4. Pour some of the Ringer’s solution to two 35-mm Petri dishes and keep them close to the dissecting microscope. 5. Kill the animal under dim red light. 6. Enucleate the eyes using the long scissors and the student Dumont forceps. 7. The rest of the procedures are carried out under the dissecting microscope using infrared light. Use the infrared viewer with illuminator, if you need to find something outside the field of view of the microscope.

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8. Remove any leftover muscle and skin tissue from the eye using the long scissors and the student Dumont forceps. 9. Tape a small piece of filter paper on the dissecting microscope stage and place the eye on it (see Note 18). Remove the anterior part of the eye, leaving the vitreous in the eyecup. Use the short scissors to make an incision and cut around just behind the ora serrata. 10. Transfer the eyecup into one of the Petri dishes filled with Ringer’s. Carefully remove the vitreous using the fine forceps. 11. Under the infrared light, the retina is now visible against the dark background of the retinal pigment epithelium. Gently separate the retina from the epithelium; it will remain attached to the eyecup at the optic nerve. With the fine forceps, reach underneath the retina and pinch it off at the point of attachment. Separate the retina fully from the eyecup (see Note 19). 12. Using a plastic pipette, draw some solution containing the retina and transfer it to the other, clean Petri dish. It can be kept there in a light-tight box for a few hours (see Note 20). 3.1.3. Isolated Living Photoreceptor Cells

1. Bring pipettes, coated chambers, Sylgard-covered dishes close to the dissecting microscope. Grab a piece of razor blade with the blade holder, with the edge of the blade at approximately 45◦ angle to the holder. All subsequent procedures are carried out under the dissecting microscope using infrared light. 2. Using the small scissors, cut the retina into 2–3 pieces. With a plastic pipette, draw some solution containing a piece and transfer it into a Sylgard-covered dish. The final volume of the solution in that dish should be about 250 μl. 3. With the fine forceps, flatten the piece of retina on the Sylgard surface, keeping the photoreceptor side up. Using the razor blade, chop the piece in one direction; repeat 3–4 times, then rotate the dish 90◦ , and chop again 3–4 times. Repeat the whole procedure until you see a “cloud” of dissociated cells. It is important to chop finely, while keeping the piece of retina stuck to the Sylgard. If the chopping is too coarse, or the retina becomes unstuck, one gets mostly pieces of retina instead of isolated cells. 4. After finishing the chopping, transfer 200 μl of the solution to an experimental chamber. Keep the chamber with the isolated cells in a light-tight box.

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5. Wait for 10 min for the cells to settle and then add 2–3 ml Ringer’s. The cells can now be taken to the microscope stage for the experiment (see Note 21). 3.2. Fluorescence Imaging Measurements

1. Bring the fiber-optic cable of the illuminator to be used for bleaching the cells inside the light-tight cage and above the microscope stage; secure it so that its end is at a distance of about 2 in. from the nose of the objective lens. The nose of the lens should be at the center of the illuminating beam to ensure bleaching of the cells. 2. Put immersion oil on the objective lens (see Note 22). 3. Bring a dish over to the stage with the IR viewer using infrared light from its illuminator (see Note 23). Bring the perfusion components to their positions and begin perfusion. 4. Close the curtains to the light-tight cage surrounding the microscope. The preparation should now be in darkness. 5. Make sure that all electronic equipment is switched off and only then turn on the xenon lamp (see Note 24). Next, turn on the rest of the equipment, including camera, computer, and, for mouse experiments, the heating components. Last, turn on the monitor covered with red plastic. 6. Turn on the microscope IR illumination and, with the camera in live mode, move the stage and find a cell. When working with isolated salamander cells, the overwhelming majority of the cells are green-sensitive rods and redsensitive cones (see Note 25). 7. Only outer segments with attached ellipsoids can generate retinol. Among those, it is best to use whole intact cells (with outer segment, ellipsoid, and nucleus). 8. Carry out preliminary measurements to determine proper focus for fluorescence. Determine what the cell should look like under IR illumination to be in focus for retinol fluorescence. Make sure that the retinol signal is well above background and does not saturate the camera. Perform initial measurements with rod cells to measure the time course of fluorescence production. Carry out tests after a post-bleach period at which time maximum retinol fluorescence is observed. Ensure that the measuring light does not photobleach retinol; make repeated measurements (about 10) of fluorescence in rapid succession. Any significant diminution (more than 0.5% per individual measurement) of the fluorescence signal can be attributed to retinol photobleaching. If significant photobleaching is observed, use the neutral density filters to attenuate the excitation light intensity and reduce the exposure time. With a CCD camera that has

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adequate sensitivity, using 1–10% of the xenon lamp intensity and 100–500 ms exposure times, should provide a clear retinol fluorescence signal without significant photobleaching (see Note 26). 9. For an experiment, find a cell under IR, center it in the field, and adjust the focus for fluorescence measurement. Switch off the IR and capture a fluorescence image. Turn on the fiber-optic illuminator and bleach the cells on the microscope stage. Use 1 min illumination for rods and 10 s for cones. Switch off the illuminator and capture another fluorescence image (for rods) or capture a series of images with a time delay acquisition routine (for cones) (see Note 27). Switch on the IR to check the focus and continue, capturing images at specified times after bleaching. Remember to switch off the IR before capturing a fluorescence image. 10. Keep in mind that the focus drifts and that the cell might move slightly. Refocus for each measurement (except for the one measurement immediately after bleaching, when speed is of the essence). 3.3. Analysis of Fluorescence Imaging Data

1. Use the software to define regions of interest (ROI) in the outer segment and in the background. 2. Use the ROIs to measure average fluorescence intensity for outer segment and background regions. 3. Subtract background fluorescence from that of the outer segment ROI to obtain the outer segment intensity due to the outer segment fluorophore. Obtain outer segment fluorescence intensity for each time point. 4. Continue analysis for your purposes. Example, subtract initial control value to obtain the outer segment fluorescence due to retinol. Alternatively, normalize over the initial control value or analyze kinetics according to different models, etc.

4. Notes 1. It is critical that the buffer composition is accurate within a few percent. The osmolarity of the solution affects the function and viability of the cells, especially the murine ones.

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2. We have obtained the same results using a Ringer’s solution with a slightly different composition: 110 mM NaCl, 2.5 mM KCl, 1.6 mM MgCl2 , 1.0 mM CaCl2 , 10 mM glucose, 10 mM HEPES, pH 7.8. 3. The addition of bovine serum albumin is not strictly necessary, but we have found that it appears to improve the viability of the cells. At this concentration, albumin does not affect the removal of retinol from the outer segment. 4. This longpass filter can be used for bleaching the visual pigments of the salamander green-sensitive rods and redsensitive cones, and mouse rods. These three cell types comprise the overwhelming majority of the cells isolated from the retinas of these two species. For experiments with the salamander blue-sensitive rods and cones and UV-sensitive cones, a much more demanding technical approach is required, including different filters for bleaching the visual pigments of these cells (see Section 3.2, step 5). 5. You can place solutions for perfusion on the top of the cage (instead of inside). In that case, the top plate should have a 2–3-in.-diameter hole to allow the tubes carrying the solutions to enter the cage. 6. You can improve the light tightness of the enclosure by attaching a 5-in. skirt made of black cloth around the bottom edges of the plates. 7. The width of the inside curtain should be the same as the width of the frame; its top side should be permanently attached to the frame. The horizontal rod at the bottom is used to roll the curtain up or down as needed. 8. The outside curtain must fully cover the inside one, so its width is longer than the frame. It is not permanently attached to the frame and is kept in place during experiments with Velcro strips at the top and the right and left sides. Place the Velcro on the frame and the curtain, so that the curtain is securely held but is also loose at the bottom. This is necessary to allow the hands of the experimenter to go under the outside curtain and through the slits of the inside one to reach the microscope. 9. A half-in.-thick piece of wood that is about 5 in. deep and runs the length of the front side of the cage is adequate. 10. The flow rate can be controlled by adjusting the height of the reservoir containing the solution or by compressing the plastic tubing that brings the solution from the reservoir to the chamber.

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11. Placement of the needle is critical for avoiding complete removal of the solution in the chamber and drying the cells. A disadvantage of the suction system is that it is difficult to avoid pulsation, because the height of the solution in the chamber tends to fluctuate. Because the suction generates a sound, however, it allows monitoring of perfusion from the outside of the light-tight cage. Cessation of the sound is an important warning sign, indicating potential flooding on the microscope stage. 12. The high numerical aperture is necessary to collect as much fluorescence signal as possible. A 100× lens might be somewhat more helpful especially for visualizing the small mouse photoreceptor cells, but because of the smaller field of view it makes it more difficult to find a cell. In our experience, an oil-immersion 40× lens with 1.3 numerical aperture (for example, the Zeiss Plan Neofluar or even the Fluar) is perfectly adequate. 13. The objective lens is a major heat sink and heating it helps to stabilize the temperature in the experimental chamber. Temperature fluctuations in the lens change the temperature of the immersion oil, resulting in changes in its refractive index and focus drift. 14. The shutter is essential to avoid photobleaching of retinol. The cells should be exposed to the excitation light only for image capture. 15. Do not crowd or try to cover the xenon light source to reduce light leak. The lamp generates a lot of heat and good ventilation is necessary to avoid overheating. 16. The absorption maximum of retinol is 325 nm, and its fluorescence emission maximum is ∼480 nm (15). The glass optics used (lenses and light guide) transmit poorly <350 nm, hence the use of the particular excitation filter. Use of very expensive quartz optics should allow efficient excitation with 325 nm light, but we have no direct experience with such a system. The dichroic mirror and the longpass emission filter are selected to collect most of the fluorescence emission. 17. Do your best to ensure the health of the animals, because the health of the cells depends on it. 18. It is not necessary to use filter paper, but we have found that it helps for handling the small salamander and mouse eyes by keeping them in place. 19. Sometimes it is difficult to separate the retina from the pigment epithelium. Slowly peel off starting from the periphery. If you still cannot separate the retina, a likely possibil-

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ity is incomplete dark adaptation, which could be caused by too bright a red light. Ensure that the animal is dark adapted properly and dim the red light. 20. It is a good idea to cut a small piece of retina and transfer it to a separate Petri dish. Under room lights, this piece of retina should be a bright red color (due to rhodopsin) that fades rapidly. The red color indicates the presence of rod outer segments and confirms that you have obtained a healthy retina. Lack of red color indicates either an unhealthy retina or a failure to separate the rod outer segments from the retinal pigment epithelium. In such case, you should ensure the health of the animals and proper dark adaptation. 21. Before you embark on actual experiments, you need to ensure that the chopping and the cell density have been optimized. A very high cell density will result in cells settling on top of each other, disallowing an experiment. A very low density might result in failure to find a cell for experiment. Check your isolated cell preparations under the infrared illumination of the microscope, and adjust chopping and density until you can regularly obtain isolated intact cells that have settled without cells above or below them. 22. At the end of the day, remove any residual oil on the lens with lens paper. With use, oil seeps under the lens. Remove and clean the lens regularly with cleaning fluid and lens paper. You can use cotton-tipped applicators to clean the back surface of the lens. 23. After the first experiment of the day, the equipment has been switched on. If you need to switch off the xenon lamp to place a second dish on the microscope stage, then you will need to switch off all pieces of equipment (see Note 24 below). However, with the xenon lamp on the floor, the light leaking from it should not reach the microscope stage inside the cage. If this is indeed the case, switch off only the computer monitor and open the light-tight box containing the chamber with the cells inside the cage. Continue as with the first dish of the day. 24. When the xenon lamp is turned on, it could generate a voltage spike that might damage other electronic equipment. 25. Blue-sensitive salamander rods and blue- and UV-sensitive cones are each morphologically distinct and can be distinguished from the green-sensitive rods and red-sensitive cones (17). For experiments with these types of cells, however, it is best to independently corroborate their identity

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by measuring their spectral sensitivity. Such measurements require additional equipment (10). 26. An important additional parameter for image acquisition is image binning. Binning improves the signal-to-noise ratio at the expense of resolution by grouping the output of camera pixels. With the specified hardware components, and exposure times 100–1000 ms, we find that 2×2 and 4×4 binning provide images with adequate resolution and good signal-to-noise ratio. 27. In cone outer segments, retinol fluorescence increases rapidly after bleaching, so it is necessary to employ a time delay image acquisition routine to capture a series of images in rapid succession. This image acquisition routine is an option provided by the software and is necessary for capturing images with time delays less than 1 min. For longer time delays between images, single image capture is generally better. Using a time delay routine for an experiment lasting for more than 5 min, although not impossible, is usually unsuccessful, because the focus drifts and the cell might move. Refocusing with the IR before each image capture is necessary for these longer experiments.

References 1. Fain, G.L., Matthews, H.R., Cornwall, M.C., Koutalos, Y. (2001) Adaptation in vertebrate photoreceptors. Physiol. Rev. 81, 117–151. 2. Imanishi, Y., Lodowski, K.H., Koutalos, Y. (2007) Two-photon microscopy: Shedding light on the chemistry of vision. Biochemistry 46, 9674–9684. 3. Lamb, T.D., Pugh, E.N., Jr. (2004) Dark adaptation and the retinoid cycle of vision. Prog. Retin. Eye Res. 23, 307–380. 4. McBee, J.K., Palczewski, K., Baehr, W., Pepperberg, D.R. (2001) Confronting complexity: The interlink of phototransduction and retinoid metabolism in the vertebrate retina. Prog. Retin. Eye Res. 20, 469–529. 5. Saari, J.C. (2000) Biochemistry of visual pigment regeneration: The Friedenwald lecture. Invest. Ophthalmol. Vis. Sci. 41, 337–348. 6. Mata, N.L., Radu, R.A., Clemmons, R.C., Travis, G.H. (2002) Isomerization and oxidation of vitamin a in cone-dominant retinas: A novel pathway for visual-pigment regeneration in daylight. Neuron 36, 69–80. 7. Wang, J.S., Estevez, M.E., Cornwall, M.C., Kefalov, V.J. (2009) Intra-retinal visual cycle required for rapid and complete cone dark adaptation. Nat. Neurosci. 12, 295–302.

8. Tsina, E., Chen, C., Koutalos, Y., AlaLaurila, P., Tsacopoulos, M., Wiggert, B., Crouch, R.K., Cornwall, M.C. (2004) Physiological and microfluorometric studies of reduction and clearance of retinal in bleached rod photoreceptors. J. Gen. Physiol. 124, 429–443. 9. Wu, Q., Blakeley, L.R., Cornwall, M.C., Crouch, R.K., Wiggert, B.N., Koutalos, Y. (2007) Interphotoreceptor retinoid-binding protein is the physiologically relevant carrier that removes retinol from rod photoreceptor outer segments. Biochemistry 46, 8669– 8679. 10. Ala-Laurila, P., Kolesnikov, A.V., Crouch, R.K., Tsina, E., Shukolyukov, S.A., Govardovskii, V.I., Koutalos, Y., Wiggert, B., Estevez, M.E., Cornwall, M.C. (2006) Visual cycle: Dependence of retinol production and removal on photoproduct decay and cell morphology. J. Gen. Physiol. 128, 153–169. 11. Wu, Q., Chen, C., Koutalos, Y. (2006) Alltrans retinol in rod photoreceptor outer segments moves unrestrictedly by passive diffusion. Biophys. J. 91, 4678–4689. 12. Weng, J., Mata, N.L., Azarian, S.M., Tzekov, R.T., Birch, D.G., Travis, G.H. (1999)

Microfluorometric Measurement of the Formation of All-Trans-Retinol Insights into the function of Rim protein in photoreceptors and etiology of Stargardt’s disease from the phenotype in abcr knockout mice. Cell 98, 13–23. 13. Harosi, F.I. (1975) Absorption spectra and linear dichroism of some amphibian photoreceptors. J. Gen. Physiol. 66, 357–382. 14. Chen, C., Blakeley, L.R., Koutalos, Y. (2009) Formation of all-trans retinol after visual pigment bleaching in mouse photoreceptors. Invest. Ophthalmol. Vis. Sci. 50, 3589–3595. 15. Chen, C., Tsina, E., Cornwall, M.C., Crouch, R.K., Vijayaraghavan, S., Koutalos, Y. (2005) Reduction of all-trans retinal to alltrans retinol in the outer segments of frog and mouse rod photoreceptors. Biophys. J. 88, 2278–2287.

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16. Kolesnikov, A.V., Ala-Laurila, P., Shukolyukov, S.A., Crouch, R.K., Wiggert, B., Estevez, M.E., Govardovskii, V.I., Cornwall, M.C. (2007) Visual cycle and its metabolic support in gecko photoreceptors. Vision Res. 47, 363–374. 17. Perry, R.J., McNaughton, P.A. (1991) Response properties of cones from the retina of the tiger salamander. J. Physiol. 433, 561–587. 18. Ma, J., Znoiko, S., Othersen, K.L., Ryan, J.C., Das, J., Isayama, T., Kono, M., Oprian, D.D., Corson, D.W., Cornwall, M.C., Cameron, D.A., Harosi, F.I., Makino, C.L., Crouch, R.K. (2001) A visual pigment expressed in both rod and cone photoreceptors. Neuron 32, 451–461.

Chapter 8 HPLC / MSN Analysis of Retinoids James E. Evans and Peter McCaffery Abstract This protocol describes a highly sensitive and selective method to quantify retinoids using normal-phase HPLC with online APCI MSN . The retinoids are key regulators of gene expression, retinol being oxidized via a retinaldehyde intermediate to retinoic acid (RA) which activates specific nuclear receptors, the signalling of which is turned off by oxidative inactivation of the ligand to 4-oxo-RA and other metabolites. Many of these retinoids are present only transiently at low concentrations in tissues and during analysis are labile to heat, light, and oxygen. HPLC with online APCI MSN provides a rapid technique to quantify these retinoids simultaneously. Techniques to extract the retinoids and prevent their degradation are described, with an emphasis on transcriptionally active RA. RA controls patterning of gene expression in the embryo, organizing embryonic morphology in the central nervous system. Similarly, a patterned distribution of RA controls function of the adult CNS, a tissue particularly difficult to analyse for RA because of its high lipid content. To understand how these patterns are organized in the brain and change over time, it is essential to determine the concentration of RA in small areas of tissues, and techniques of exquisite sensitivity are indispensable. Key words: Retinoic acid, retinol, retinaldehyde, 4-oxo retinoic acid, high-performance liquid chromatography, mass spectrometry, brain.

1. Introduction 1.1. Why Assay for Retinoic Acid?

Vitamin A is defined as the group of compounds derived from C20-beta-ionone that demonstrate the biological activity of alltrans retinol (1). It is a member of the ‘retinoids’ family, themselves defined as a group of compounds comprising of four isoprenoid units joined head to tail (2). The major active members of this group are retinol itself and its oxidative metabolites retinaldehyde and retinoic acid (RA) (Fig. 8.1).

H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_8, © Springer Science+Business Media, LLC 2010

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Fig. 8.1. The main active members of the retinoids: retinol, the form carried in the circulation and which can act as a kinase cofactor; retinaldehyde, the chromophore necessary for visual transduction; and retinoic acid, the retinoid which binds to specific nuclear receptors to regulate gene transcription. Retinol and retinaldehyde are interconverted via a retinol dehydrogenase (RDH) and retinaldehyde is irreversibly oxidized to retinoic acid by a retinaldehyde dehydrogenase (RALDH).

The biological activity of vitamin A includes support of visual transduction via the requirement of 11-cis retinaldehyde as the light-responsive component of rhodopsin. Retinol itself can function as a cofactor for a number of kinases and derivatives such as retroretinol and anhydroretinol may act on similar intracellular signalling pathways (3). The best understood retinoid signalling pathway required for a great number of physiological events is RA’s regulation of transcription via its binding to specific nuclear receptors. This is required for events such as organ development in the embryo and, in the adult, control of epithelial cell differentiation and proliferation, lung, bone, skin, and brain function (4). As a consequence of these functions the storage and/or transport forms of the retinoids, such as retinol and retinyl esters, are naturally present at high concentration (μM). Their more polar metabolic products, which are produced locally to function as transient signalling molecules, as well as their inactive metabolites, are inevitably present at much lower concentrations (nM) and are thus more difficult to detect. The brain is a region that has been a topic of recent interest with regard to RA function (5–7) and, for instance, brain-related defects are among the first symptoms of vitamin A deficiency in chick, specifically ataxia, a loss of balance and coordination with unsteady gait, which can be reversed (although not always completely), with RA treatment (8). The brain, however, is a prime example of one of the great problems in retinoid research; it is very difficult to quantify and localize the transcriptionally active metabolite of the retinoids, RA. Unlike proteins, which can be pinpointed using antibodies, or mRNA, localized with antisense, no such probes exist for RA. Particularly in the brain, where RA acts within very discrete regions in a concentration-dependent manner, knowledge of RA’s distribution within the many subcompartments of the brain is essential to understand its function. At present, the most effective way to determine RA distribution is

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through the localization of the protein required for RA synthesis, specifically the enzyme retinaldehyde dehydrogenase (RALDH) which catalyses the last step of RA synthesis and whose pattern of expression corresponds very well with the distribution of RA (9–11) (Fig. 8.1). However, to quantitatively determine how retinoids, or RA specifically, change over time, condition, or vary between tissue or species, their direct measurement is necessary. 1.2. Methods of Retinoid Detection 1.2.1. Problems with the Fragility of RA

The five or more double bonds present in retinoids make them easily oxidized in air, isomerized by light, and altered by heat; this is particularly the case when concentrations are low. Thus, it is essential that these conditions are controlled through use of antioxidants such as butylated hydroxytoluene in solvents and maintaining samples under an inert atmosphere using argon or nitrogen gas, avoiding high temperatures, and preparing samples under long-wavelength gold or yellow light and/or low-light conditions. The retinoids are also labile to strong acids particularly in anhydrous conditions, while during preparation there may be adsorption of retinoids to glass or plastic containers or steel components of the HPLC setup (4, 12).

1.2.2. Solvent Extraction of RA from Tissue

The retinoids in general are insoluble in water and soluble in organic solvents. Certain retinoids have a solvent preference; for instance relatively polar solvents like methanol and ethanol are excellent for retinol and RA but very poor for retinyl esters, which prefer non-polar solvents like hexane while solvents such as diethyl ester, chloroform, dichloromethane, dimethyl sulfoxide, and ethyl acetate are excellent for most retinoids (13). Therefore the choice of solvents is dependent on the retinoid to be extracted. A further useful property of the solvent is its relative volatility if the sample is to be concentrated for later analysis. Obviously, when extracting any retinoid from tissues, a significant amount of water will also be released and solvents that are water-immiscible will not be efficient in extraction. One key step of extraction is the release of RA from the binding proteins, cellular retinoic acid-binding proteins I and II (CRABPI and II), which carry it in the cell; this can be brought about by precipitation in ethanol or methanol.

1.2.3. Chromatography

The earliest methods to detect RA included paper and thin layer (14) chromatography. Lipids extracted with diethyl ether would be separated on paper strips and zones of lipid migration cut out under ultraviolet (UV) light and determined for colour reaction with antimony chloride (8). Alumina and silicic acid were among the first type of material to be used for column chromatography (15, 16) as well as ion exchange chromatography (17) but such

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methods would not provide complete resolution of retinoids (18, 19) and were also prone to generation of artefacts (20, 21). The use of SephadexTM gel filtration chromatography improved resolution and also gave good recovery of retinoids (22). The next step in development was high-performance liquid chromatography (HPLC), initially using normal phase with silica columns (23), but this had some difficulties because of the presence of water from the sample, and reverse-phase chromatography with, for instance, Spherisorb ODS provided good resolution and could be completed over a relatively short period of time (24). Use of gradient elution with reverse-phase HPLC could separate the full spectrum of retinoid isomers (25), important, for instance, because the differing isomers of RA have very different affinities for the RA receptors. Modern high-purity silica-based chromatographic materials are low in metal contaminants (binding sites for retinoids at low levels and promoting their isomerization and oxidation) and are also more robust; thus effective normal-phase chromatography is now achievable (26). Overall, reverse-phase HPLC is often preferred for biological samples because separations are generally less sensitive to slight changes in mobile-phase composition. They also equilibrate more rapidly for gradient separation and thus can be useful for simultaneous runs of different retinoid classes. However, normal-phase (adsorption) chromatography is often better at separating closely related compounds such as cis/trans retinoid isomers. Techniques to obtain the required specificity for low-level retinoid assay through cleanup before analysis by HPLC with UV or fluorescence detection were essential. This was particularly the case for tissues with a high lipid content such as liver, kidney, and brain and the difficulty in separating these lipids from retinoids with similar chromatographic properties. However, the problem with extensive preparation before analysis is the greater possibility for losing sensitive, low-level retinoids during the process. Use of column-switching (27) and online solid-phase extraction (28) has enabled efficient enrichment of samples. Solid-phase extraction based on aminopropyl columns has been found to be helpful with analysis of both polar and apolar retinoids from samples with a high lipid content (29). The importance of such cleanup techniques though has been reduced with the development of detection methods of ever-increasing specificity. 1.2.4. Methods of Detection and Mass Spectrometry

Common HPLC detection methods have included fluorescence detection (for retinol and its esters) and UV absorption. The characteristic UV absorption spectra of the different retinoids have led to the use of diode-array detection allowing identification of the resolved retinoids. Mass spectrometry (MS), however, has led to a large improvement in the capacity to specifically detect and identify low levels of retinoids. One of the first uses of MS was

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in conjunction with gas chromatography; however, this was not ideal because of the necessity for derivativation to form volatile, thermally stable derivatives and the sensitivity of retinoids to disruption by the high temperatures necessary for their GC elution (30). These factors lead to extensive cis/trans isomerization of retinoids and losses due to oxidation. Use of negative ion electrospray ionization (ESI) mass spectrometry (31) allows direct analysis of carboxyl bearing retinoids (RA) with little sample handling but does not allow sensitive detection of neutral retinoids. Atmospheric pressure chemical ionization (APCI)-mass spectrometry provides efficient ionization of both acidic and nonionic retinoids such as retinol and retinyl palmitate directly from a reverse-phase column, avoiding the problem associated with gas chromatography (32). Recent studies have relied upon positive ion APCI MS to quantify simultaneously retinol and RA with better sensitivity than ESI, as well as negative ion APCI, and also to provide simultaneous determination of retinoids (33). These findings were confirmed by Kane et al. (34) who also demonstrated the increased detection specificity that can be obtained with crude extracts by the use of tandem MS (MS/MS). 1.2.5. Example of Detection of Retinoids in Tissue, the Brain

Detection of retinoids in the brain has proven particularly difficult; the myelin that surrounds all nerves in the CNS results in a large amount of lipid in brain extracts, from which the retinoids need to be distinguished. To provide a useful measurement of RA in the brain, the sensitivity has to be sufficient to determine concentration within brain subregions. Determination of RA levels in whole rat brain obtained values of 6.3 pmol/g tissue, the lowest value in all regions tested compared to, for example, testes, liver, fat, and pancreas, the latter particularly high at 29.3 pmol/g tissue (35). It has been proposed that most of the RA in the brain (as well as the liver) was simply derived from the low concentrations (1.8 pmol/g) present in the circulation, in experiments infusing radiolabelled RA at steady state (35). However, quite different results for RA levels in the CNS were found in a study on retinoids in the vitamin A-deficient rat treated with physiological levels of radiolabelled retinol. Of all organs examined the region of the cortex and hippocampus had the highest ratio of RA to retinol, and with regard to total RA only liver and kidney contained higher amounts of RA per gram tissue than brain. Despite the earlier finding that the majority of RA in the brain derived from plasma it was found that this transport was no greater in the brain than any other tissue. One potential explanation for the exceptionally high levels of RA found in the brain in this study is the use of vitamin A-deficient animal models which may result in a compensatory increase in the enzymes that generate RA. If the brain has a particularly high requirement for RA this may result in a greater elevation of these synthetic enzymes in the brain and,

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when radiolabelled retinol is added, this results in greatly elevated levels of RA. 1.2.6. Current Method

This chapter describes our use of normal-phase silica gel-based HPLC with online APCI MSN for the analysis of crude tissue and media extracts allowing the sensitive analysis of a broad range of biologically important retinoids in a single assay (26). The use of a low binding site high-purity silica gel (low trace metal) based HPLC column packing allows trace levels of sensitive retinoids to be assayed without significant losses. Positive ion APCI provides for efficient ionization of all the retinoids as opposed to ESI that only provides sensitive ionization for retinoids that contain free carboxyl groups. Quadrupole ion trap mass spectrometry provides unique capabilities for multistage tandem MS (MSN ) which allows a number of sequential stages of precursor–product ion MS/MS to be performed. This is key to obtaining high specificity while maintaining high sensitivity. The high specificity of MSN (MS3 and MS4 ) used here allows the analysis of low-level retinoids in crude lipid extracts without interference from other sample components. This ability to directly analyse crude extracts without multiple steps of prefractionation prevents sample handling losses that would otherwise be unavoidable. Methods described below are designed to provide high recovery of native retinoids with sensitive, specific, and accurate quantification.

2. Materials All solvents used were HPLC grade or higher. 2.1. Extraction of Retinoids from Tissues

1. Extraction solvent: ethanol–isopropanol (2:1, v/v) containing 1 mg/ml butylated hydroxytoluene antioxidant (>99%, Sigma Chemical Company). 2. Dounce glass homogenizer (7 ml, Wheaton, Millville, NJ).

2.2. Extraction of Retinoids from Media and Biofluids 2.3. HPLC Materials

Hexane–dioxane–isopropanol (50:5:1, v/v/v) containing 1 mg/ml butylated hydroxytoluene antioxidant (>99%, Sigma Chemical Company). 1. Inertsil silica normal-phase HPLC column (150 × 2 mm, 5 μm particle, 5 μm particle; Keystone Scientific, Inc., Bellefonte, PA). This column was chosen for its high-purity silica gel packing and 2 mm ID that has an optimum flow rate of ∼200 μl/min for high APCI sensitivity on the Thermo LCQ mass spectrometry system used. The HPLC

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pumping system used was a Leap Technologies Rheos Flux 2000 ternary gradient system with a Rheodyne 7125 injector equipped with a 10 μl PEEK sample loop. All connection tubing was 0.005 in. ID PEEK. 2. Gradient elution mobile phases for separation of a broad spectrum of retinoids. Solvent A: n-hexane. Solvent B: n-hexane–dioxane–isopropanol (40:8:2, v/v/v). HPLC mobile phases were continuously sparged with helium (99.9%) to remove oxygen that could oxidize retinoids before and during analyses. 3. Isocratic mobile phase for resolving all-trans, 9-cis, and 13-cis RA. n-hexane–dioxane–isopropanol (70:26:4). 4. Retinaldehyde, all-trans, 9-cis, and 13-cis RA and retinol (Sigma Chemical Company) and didehydro-RA and 4-oxoRA (gift from Dr. J Grippo, Hoffmann-La Roche, Inc., Nutley, NJ, USA) stock standards dissolved in DMSO at 0.1 M (see Note 1).

3. Methods 3.1. Tissue Extraction

All extraction procedures were performed under low-intensity yellow light. 1. All tissues were collected as fresh as possible and stored at −80◦ C until extraction (see Notes 2 and 3). 2. The fresh or freshly thawed tissue was deposited into the cold homogenizer containing 1 ml of extraction solvent per 100 mg wet weight tissue and the tissue homogenized. 3. The homogenate was centrifuged at 3000×g for 10 min at 3◦ C and the supernatants collected and stored in the dark at −80◦ C until analysis.

3.2. Media and Biofluid Extraction

The cold fluid (0◦ C, 200 μl) was vortexed with 200 μl of cold extraction solvent in a TeflonTM sealed tube under nitrogen for 1 min and centrifuged at 3000×g for 10 min at 3◦ C. The supernatants were collected, briefly stored in the dark at −20◦ C, and analysed as soon as possible (see Note 4).

3.3. HPLC/MSN Analysis of Retinoids

The methods presented here utilized a Thermo LCQ 3D ion trap mass spectrometer with the ability to perform MS3 or 4 experiments. These MSN capabilities are used to provide highspecificity analyses while maintaining the necessary sensitivity for low-level retinoid quantification. While many of the ion trap instruments that are currently available have these capabilities,

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current-generation linear ion traps can be expected to provide higher sensitivities (compared to 3D ion traps) for MSN analyses due to their better ion trapping efficiencies, higher ion capacities, and more sensitive detection. Two HPLC/MSN methods are presented here. One utilizes gradient solvent elution to provide analysis of the full spectrum of retinoids in less than 20 min. The other is an isocratic method that provides improved resolution of all-trans, 9-cis, and 13-cis RA isomers for their individual quantification. 3.3.1. Gradient Elution Normal-Phase HPLC

Under low-level yellow light, extracts were warmed to 0◦ C immediately before analysis and 3 μl of the extract (see Note 5) was drawn into a cold 10 μl gas-tight Hamilton glass syringe and rapidly loaded and injected onto the column at initial solvent conditions (90% solvent A–10% solvent B flowing at 200 μl/min). Immediately after injection a 18.9 min linear gradient to 58% solvent A–42% solvent B was initiated, also at 200 μl/min.

3.3.2. APCI MSN Analysis of Major Retinoids

The column was directly connected to the APCI source of the mass spectrometer and the source operated with the vaporizer at 375◦ C, the nitrogen sheath gas flow at 35% (relative), the source current at 5 μA, and the heated capillary at 150◦ C. A constant infusion of 1.0 μg/ml RA in n-hexane at 200 μl/min was used to optimize the APCI source and auto-tune the mass spectrometer using the m/z 301 ion (MH+ ) (see Note 6). Each analyte separated by HPLC was detected by a unique series of MSN scan functions that were optimized by trial of multiple precursor/product ion combinations and collision energies to provide maximum selectivity and sensitivity as listed in Table 8.1 and illustrated in Fig. 8.2 (see Note 7). The isolation width used was adjusted to provide the precursor ion free of any adjacent ions while maintaining maximum ion current. Examples of the spectra for predominant retinoids, retinaldehyde, RA, and retinol, are shown in Fig. 8.3.

3.3.3. Isocratic HPLC APCI MSN Analysis of Retinoic Acid Isomers

This mode of operation is preferred when RA isomers are the only analytes of interest. All operating conditions were the same as described above except for the mobile phase and the scan functions that were used. The isocratic mobile phase results in elution of all three isomers in less than 14 min. The scan function used was the one listed in Table 8.1 for RA isomers (m/z 301, 205, and 159). Improved chromatographic resolution resulted from the isocratic separation and the dedicated scan mode used under these conditions lead to improved quantification for the RA isomers and provided shorter overall analysis times because of the faster elution of RA’s and lack of a requirement for column reequilibration following each injection.

12–20

16.2

16.2

Retinol

4-oxo-RA

315.1

269.1

299.1

DidehydroRA

8.8

285.1 301.1

Retinaldehyde 5.7 RA isomers 7.5–8.6

0–6 5–12

Precursor m/z (MH+)

Retinoid

Retention time (min)

Time segment (min)

MS2

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32

30

30 30

297.1

213.1

243.1

193.1 205.1

Collision Precursor energy (m/z)

MS3

Table 8.1 Scan functions and approximate HPLC retention times for analytes

30

32

30

30 30

279.1

Collision Precursor energy (m/z)

MS4

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209.1

156.0

225.1

175.1 159.1

0.90

3.80

35.0

12.0 1.3

Overall Collision Quantification efficiency energy ion (m/z) (%)

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Fig. 8.2. Example of HPLC APCI MSN analysis. Normal-phase HPLC separation (Inertsil silica column) and APCI MSN product ion scans of retinoid standards.

3.3.4. Results

The mass spectrum obtained of RA, retinaldehyde, and retinol analysed at each stage of MSN analysis illustrates the selection of product ions available at each MSN stage for selection as precursors for subsequent utilization as shown in Fig. 8.3.

Fig. 8.3. Representative APCI MSN spectra of retinoids. The precursor and quantitation product ions chosen for analysis of retinaldehyde, retinoic acid, and retinol are shown.

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4. Notes 1. RA standards are relatively stable at high concentrations (e.g. 0.1 M in DMSO). Low-concentration standards (μM and below) should be diluted immediately before use. DMSO is preferable to ethanol as a solvent as it is less volatile and thus less is evaporated when flushed with argon or nitrogen. 2. The high sensitivity of retinoids to oxidation, light, and heat is still the case even when present in tissue, and best results are always obtained when the time between removal of tissue and analysis is kept as short as possible, i.e. the same day. If this is not possible, the tissue should be snap frozen in liquid nitrogen and kept no longer than a week, preferably flushed with argon or nitrogen, and the container in which it is kept sealed. 3. A tissue with high retinoid content to function as a positive control is the eye, which is simple to dissect and retinoids are easy to extract with its lower lipid content compared to other CNS regions. 4. Retinoids have a high affinity for the silanol groups of glass and glassware is best considered disposable. Glassware can be silanized with a 5% (w/v) solution of dichlorodimethylsilane in toluene followed by washing with a 1:l (v/v) mixture of methanol and acetone. 5. Injection volumes of the ethanol–isopropanol extracts must be less than 3 μl to avoid shortening of retention times and degradation of chromatographic resolution by the relatively polar solvent. Because extraction efficiency of RA is decreased when lower polarity solvents are used and that serious losses are encountered when concentration is attempted we have not been able to increase the tissue equivalent amount of extract injected. 6. It is important to tune the APCI ion source and ion optics using a solution of a standard analyte (RA) infused at chromatographic flow rate in a solvent similar to the chromatographic solvent to obtain the highest possible sensitivity. This will assure optimal ionization efficiency and ion transmission with minimal ion losses in the high-temperature and high-pressure regions of the mass spectrometer. 7. In most cases MS3 product ion scans were used to provide high-specificity detection of retinoids; however, for 4-oxo-RA the first two successive high-efficiency collisional

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activation decomposition products were through loss of water, which do not impart high detection specificity. Hence an additional MS4 step from the m/z 279 MS3 product ion was used to give a good yield of a m/z 209 product. References 1. International Union of Nutritional Sciences, C. I., Nomenclature (1978). (1976) Generic descriptors and trivial names for vitamins and related compounds recommendations. Nutr. Abstr. Rev. Ser. A, 831–835. 2. Nomenclature of Retinoids: Recommendations 1981. (1983) IUPAC-IUB Joint Commission on Biochemical Nomenclature (JCBN). Arch. Biochem. Biophys. 224, 728–731. 3. Chiu, H.J., Fischman, D.A., Hammerling, U. (2008) Vitamin A depletion causes oxidative stress, mitochondrial dysfunction, and PARP1-dependent energy deprivation, FASEB. J. 22, 3878–3887. 4. Sporn, M.B., Roberts, A.B., Goodman, D.S. (1994) The Retinoids: Biology, Chemistry, and Medicine, 2nd ed., Raven Press, New York. 5. Mey, J., McCaffery, P. (2004) Retinoic acid signaling in the nervous system of adult vertebrates, Neuroscientist 10, 409–421. 6. Lane, M.A., Bailey, S.J. (2005) Role of retinoid signalling in the adult brain, Prog. Neurobiol. 75, 275–293. 7. Tafti, M., Ghyselinck, N.B. (2007) Functional implication of the vitamin A signaling pathway in the brain. Arch. Neurol. 64, 1706–1711. 8. Krishnamurthy, S., Bieri, J.G., Andrews, E.L. (1963) Metabolism and biological activity of vitamin A acid in the chick. J. Nutr. 79, 503–510. 9. McCaffery, P., Lee, M.-O., Wagner, M.A., Sladek, N.E., Dräger, U.C. (1992) Asymmetrical retinoic acid synthesis in the dorsoventral axis of the retina. Development 115, 371–382. 10. McCaffery, P., Dräger, U.C. (1994) Hotspots of retinoic acid synthesis in the developing spinal cord. Proc. Natl. Acad. Sci. USA 91, 7194–7197. 11. McCaffery, P., Dräger, U.C. (1994) High levels of a retinoic-acid generating dehydrogenase in the meso-telencephalic dopamine system. Proc. Natl. Acad. Sci. USA 91, 7772– 7776. 12. Wyss, R., Bucheli, F. (1988) Quantitative analysis of retinoids in biological fluids by high-performance liquid chromatography using column switching. I. Determination of

13.

14.

15.

16.

17.

18.

19. 20. 21.

22.

23.

isotretinoin and tretinoin and their 4-oxo metabolites in plasma. J. Chromatogr. 424, 303–314. Furr, H.C., Barua, A.B., Olson, J.A. (1994) Analytical methods. In: Sporn, M.B., Roberts, A.B., Goodman, D.S. (eds.), The Retinoids: Biology, Chemistry, and Medicine, Raven Press, New York. Morgan, B., Thompson, J.N., Pitt, G.A. (1969) The uptake and metabolism of retinol, retinoic acid and methyl retinoate by the early chick embryo, Br. J. Nutr. 23, 899–904. Huang, H.S., Goodman, D.S. (1965) Vitamin A and carotenoids. I. Intestinal absorption and metabolism of 14c-labelled vitamin A alcohol and beta-carotene in the rat. J. Biol. Chem. 240, 2839–2844. Roberts, A.B., DeLuca, H.F. (1968) Oxidative decarboxylation of retinoic acid in microsomes of rat liver and kidney. J. Lipid Res. 9, 501–508. Zachman, R.D., Dunagin, P.E., Jr., Olson, J.A. (1966) Formation and enterohepatic circulation of metabolites of retinol and retinoic acid in bile duct-cannulated rats. J. Lipid Res. 7, 3–9. Zile, M.H., Emerick, R.J., DeLuca, H.F. (1967) Identification of 13-cis retinoic acid in tissue extracts and its biological activity in rats. Biochim. Biophys. Acta. 141, 639–641. Zile, M., DeLuca, H.F. (1968) Chromatography of vitamin A compounds on silicic acid columns. Anal. Biochem. 25, 307–316. Lippel, K., Olson, J.A. (1968) Origin of some derivatives of retinoic acid found in rat bile. J. Lipid Res. 9, 580–586. Kleiner-Bossaler, A., Deluca, H.F. (1971) Formation of retinoic acid from retinol in the kidney. Arch. Biochem. Biophys. 142, 371–377. Ito, Y.L., Zile, M., Ahrens, H., DeLuca, H.F. (1974) Liquid-gel partition chromatography of vitamin A compounds; formation of retinoic acid from retinyl acetate in vivo. J. Lipid Res. 15, 517–524. Bridges, C.D. (1975) Storage, distribution and utilization of vitamins A in the eyes of adult amphibians and their tadpoles. Vision Res. 15, 1311–1323.

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24. Frolik, C.A., Tavela, T.E., Sporn, M.B. (1978) Separation of the natural retinoids by high-pressure liquid chromatography. J. Lipid Res. 19, 32–37. 25. Bugge, C.J., Rodriguez, L.C., Vane, F.M. (1985) Determination of isotretinoin or etretinate and their major metabolites in human blood by reversed-phase high-performance liquid chromatography. J. Pharm. Biomed. Anal. 3, 269–277. 26. McCaffery, P., Evans, J., Koul, O., Volpert, A., Reid, K., Ullman, M. (2002) Retinoid quantification by HPLC/MS(n). J. Lipid Res. 43, 1143–1149. 27. Campins-Falco, P., Herraez-Hernandez, R., Sevillano-Cabeza, A. (1993) Columnswitching techniques for high-performance liquid chromatography of drugs in biological samples. J. Chromatogr. 619, 177–190. 28. Gundersen, T.E., Lundanes, E., Blomhoff, R. (1997) Quantitative high-performance liquid chromatographic determination of retinoids in human serum using on-line solid-phase extraction and column switching. Determination of 9-cis-retinoic acid, 13-cis-retinoic acid, all-trans-retinoic acid, 4-oxo-all-trans-retinoicacid and 4-oxo13-cis-retinoic acid. J. Chromatogr. B Biomed. Sci. Appl. 691, 43–58. 29. Schmidt, C.K., Brouwer, A., Nau, H. (2003) Chromatographic analysis of endogenous retinoids in tissues and serum. Anal. Biochem. 315, 36–48.

30. Ranalder, U.B., Lausecker, B.B., Huselton, C. (1993) Micro liquid chromatographymass spectrometry with direct liquid introduction used for separation and quantitation of all-trans- and 13- cis-retinoic acids and their 4-oxo metabolites in human plasma. J. Chromatogr. 617, 129–135. 31. Van Breemen, R.B., Huang, C.R. (1996) High-performance liquid chromatographyelectrospray mass spectrometry of retinoids. FASEB. J. 10, 1098–1101. 32. van Breemen, R.B., Nikolic, D., Xu, X., Xiong, Y., van Lieshout, M., West, C.E., Schilling, A.B. (1998) Development of a method for quantitation of retinol and retinyl palmitate in human serum using high-performance liquid chromatographyatmospheric pressure chemical ionizationmass spectrometry. J. Chromatogr. A. 794, 245–251. 33. Wang, Y., Chang, W.Y., Prins, G.S., van Breemen, R.B. (2001) Simultaneous determination of all-trans, 9-cis, 13-cis retinoic acid and retinol in rat prostate using liquid chromatography-mass spectrometry. J. Mass Spectrom. 36, 882–888. 34. Kane, M.A., Chen, N., Sparks, S., Napoli, J.L. (2005) Quantification of endogenous retinoic acid in limited biological samples by LC/MS/MS. Biochem. J. 388, 363–369. 35. Kurlandsky, S.B., Gamble, M.V., Ramakrishnan, R., Blaner, W.S. (1995) Plasma delivery of retinoic acid to tissues in the rat. J. Biol. Chem. 270, 17850–17857.

Chapter 9 Binding of Retinoids to ABCA4, the Photoreceptor ABC Transporter Associated with Stargardt Macular Degeneration Ming Zhong and Robert S. Molday Abstract ABCA4 is a member of the superfamily of ATP-binding cassette (ABC) transporters, which has been implicated in the clearance of all-trans retinal derivatives from rod and cone photoreceptor cells following photoexcitation as part of the visual cycle. Mutations in ABCA4 are known to cause Stargardt macular degeneration and related disorders, associated with a severe loss in vision. Recently, a solid-phase binding assay has been developed to identify retinoids that likely serve as substrates for this transporter. In this procedure, monoclonal antibodies directed either against an epitope within ABCA4 (Rim 3F4 antibody) or against the 9 amino acid 1D4 epitope tag engineered onto the C-terminus of expressed ABCA4 (Rho 1D4 antibody) are covalently bound to a Sepharose matrix. This immunoaffinity matrix is then used to isolate ABCA4 from photoreceptor outer segments or transfected cells. All-trans retinal is added to immobilized ABCA4 in the presence of a phospholipid mixture containing phosphatidylethanolamine. The bound retinoid is then analyzed directly by spectrophotometry or identified by HPLC and/or mass spectrometry following extraction with organic solvents. Using this procedure, it has been shown that unprotonated N-retinylidene-phosphatidylethanolamine binds with high affinity to ABCA4 and is released by the addition of ATP. These procedures and related radiometric assays using titrated retinal have been used to study the binding of N-retinylidene-PE to wild-type and mutant ABCA4 in the absence and presence of nucleotides for structure–function studies. Key words: ABCA4, ABC transporters, retinoids, visual cycle, photoreceptor cells, Stargardt macular degeneration, retinal degenerative diseases, N-retinylidene-phosphatidylethanolamine, immunoaffinity chromatography, monoclonal antibody.

1. Introduction ABCA4, also known as the rim protein or ABCR, is a member of the superfamily of ATP-binding cassette (ABC) transporters H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_9, © Springer Science+Business Media, LLC 2010

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expressed in photoreceptor cells (1, 2). It is localized along the rim region of rod and cone photoreceptor outer segment disc membranes where it has been implicated in the removal of retinal derivatives as part of the visual cycle (3–7). ABCA4 is a relatively abundant 250 kDa glycoprotein which, like most other full-length eukaryotic ABC transporters, is organized as two tandem halves, each consisting of a transmembrane domain followed by a cytoplasmic nucleotide-binding domain (8). Over 500 mutations in the gene encoding ABCA4 are known to cause Stargardt macular degeneration and a subset of autosomal recessive cone– rod dystrophy and retinitis pigmentosa (2, 9–12). Individuals heterozygous for selected Stargardt disease mutations in the ABCA4 gene have also been suggested to be at increased risk for developing age-related macular degeneration (13). Several studies have implicated ABCA4 in the clearance of retinoids from photoreceptor outer segments following photoexcitation. Abca4 knockout mice exposed to cyclic or continuous lighting show elevated levels of all-trans retinal, protonated N-retinylidene-phosphatidylethanolamine (N-retinylidene-PE), and phosphatidylethanolamine (PE) in retinal extracts (4, 14). Individuals with Stargardt disease and abca4 knockout mice display a progressive accumulation of lipofuscin deposits in their retinal pigment epithelial (RPE) cells (15–17). The diretinal pyridinium compound A2E is one of the major components of lipofuscin (17–19). The A2E precursor known as A2PE is formed in photoreceptor outer segments through the condensation of all-trans retinal and N-retinylidene-PE when these retinoids are not efficiently cleared from outer segments following photoexcitation. Upon phagocytosis of outer segments, A2PE is hydrolyzed to A2E in the phagolysosomes of RPE cells. A2E progressively accumulates as fluorescent lipofuscin deposits since A2E is not readily metabolized in these cells. In a separate approach, the enzymatic properties of purified and reconstituted ABCA4 have been studied to explore the role of ABCA4 in photoreceptors (5, 20). All-trans retinal and 11-cis retinal stimulate the ATPase activity of purified ABCA4 up to fourfold in the presence of PE, suggesting that retinoid compounds may serve as substrates transported by ABCA4. This was further supported by the finding that most disease-associated mutations in ABCA4 result in a marked decrease or complete loss in retinal-stimulated ATPase activity (21, 22). Since the aldehyde moiety of retinal is known to reversibly react with the primary amine of PE to form the Schiff base adduct, N-retinylidene-PE (Fig. 9.1), it was unclear from these studies whether retinal or N-retinylidene-PE serves as the substrate for ABCA4. This was investigated using a solid-phase retinoidbinding assay (23). In this procedure, the Rim 3F4 monoclonal antibody directed against an epitope (YDLPLHPRT) near the

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Fig. 9.1. Structures and reactions of all-trans retinal with phosphatidylethanolamine. All-trans retinal released upon photobleaching of rhodopsin or cone opsin can reversibly react with phosphatidylethanolamine (PE) to form N-retinylidenephosphatidylethanolamine (N-retinylidene-PE) in disc membranes. The unprotonated form is in equilibrium with the protonated form. N-retinylidene-PE can be stabilized by reduction with NaBH4 to form N-retinyl-PE. ABCA4 binds N-retinylidene-PE as well as N-retinyl-PE.

C-terminus of ABCA4 was purified on a Protein G-Sepharose column and directly coupled to CNBr-activated Sepharose 4B. The Rim 3F4-Sepharose immunoaffinity matrix was then used to isolate ABCA4 from detergent-solubilized bovine rod outer segment (ROS) membranes (Fig. 9.2). Various potential retinoid compounds were added to immobilized ABCA4 in the presence of a defined phospholipid mixture. After thoroughly washing the affinity matrix to remove unbound material, the bound retinoid was extracted from ABCA4 with an organic solvent and identified by high-performance liquid chromatography (HPLC). When all-trans retinal was added to ABCA4 in the presence of phospholipids containing PE, the bound retinoid was identified as N-retinylidene-PE on the basis of its retention time relative to known standards and its spectral properties as illustrated in Fig. 9.3 (23). Since a trifluoroacetic acid-containing solvent was used in the HPLC analysis, the protonated form of N-retinylidene-PE with an absorption maximum at 450 nm was detected by HPLC. N-retinylidene-PE can be stabilized by reduction of the Schiff base with sodium borohydride to form

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Fig. 9.2. Purification of ABCA4 and ABCA4-1D4. Endogenous ABCA4 was purified from detergent-solubilized ROS membranes on a Rim 3F4-Sepharose matrix and ABCA4-1D4 was purified from membrane preparations of transfected HEK 293 cells on a Rho 1D4Sepharose matrix. Samples of ROS membranes and purified ABCA4 were analyzed on SDS-PAGE gels stained with Coomassie blue (CB) and Western blots labeled with the Rim 3F4 (ABCA4) or Rho 1D4 (ABCA4-1D4) monoclonal antibodies.

N-retinyl-PE (Fig. 9.1). This compound, like N-retinylidene-PE, was found to bind to ABCA4 in stoichiometric amounts using the solid-phase binding assay (Fig. 9.3). In contrast, all-trans retinol in the presence of PE did not bind to ABCA4 nor did all-trans retinal in the absence of PE. This solid-phase binding assay was also used to study the effect of nucleotides on retinoid binding to ABCA4 (23). The addition of ATP or GTP resulted in the quantitative release of Nretinylidene-PE from ABCA4 suggesting that the binding of these nucleotides to the nucleotide-binding domains results in a conformational change in ABCA4 which converts the high-affinity N-retinylidene-PE binding site to a low-affinity site. The release of N-retinylidene-PE from ABCA4 in the presence of ATP has been suggested to be related to the transport of substrate across the membrane. Since analysis of retinoid binding by HPLC requires considerable amounts of ROS membranes and immunoaffinity reagents, a more sensitive radiometric binding assay was developed employing [3 H]-all-trans retinal (23). This assay requires less than onetenth the amount of immobilized ABCA4 and immunoaffinity matrix, thereby facilitating the analysis of multiple samples. From these studies, the apparent dissociation constant for the binding of N-retinylidene-PE to detergent-solubilized immobilized ABCA4 was found to be ∼5 μM in agreement with analysis by HPLC as shown in Fig. 9.4 (23).

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Fig. 9.3. HPLC chromatographs and spectra of retinoid compounds that bind to ABCA4. Upper panel: all-trans retinal was added to ABCA4 in the presence of a DOPE/DOPC phospholipid mixture. The HPLC chromatogram of bound retinoid was measured at 450 nm (λmax of protonated N-retinylidene-PE). Lower panel: N-retinyl-PE, the reduced adduct of N-retinylidene-PE, was added to ABCA4 in the presence of DOPE/DOPC phospholipid mixtures and the bound N-retinyl-PE was measured at 330 nm (λmax of N-retinyl-PE). Retention times and spectra (insets) of major peak in each chromatogram were used to identify the bound retinoid. Modified from (23).

To determine the protonation state of N-retinylidene-PE bound to ABCA4, a variation of this solid-phase binding assay was used. In this procedure, the binding of N-retinylidene-PE was carried out by adding all-trans retinal to immobilized ABCA4 in the presence of a PE as discussed above. After the column was washed to remove unbound retinoid, ABCA4 containing bound N-retinylidene-PE was released from the affinity matrix by the addition of 0.2 mg/ml of synthetic 3F4 competing peptide for analysis by spectrophotometry. N-retinylidene-PE bound to ABCA4 had an absorption maximum of 370 nm, characteristic of the unprotonated form of N-retinylidene-PE (Fig. 9.5). Recently, a variation of this solid-phase retinoid-binding assay has been developed to examine the binding of N-retinylidene-PE

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Fig. 9.4. Binding of retinoid to ABCA4 using the radiolabeling method. [3 H] all-trans retinal was added to ABCA4 in the presence of DOPE/DOPC phospholipid mixture and the bound retinoid (N-retinylidene-PE) was determined after extraction with organic solvent. The binding curve was fitted with a single apparent Kd of 5.4 μM. Inset: Scatchard plot of the binding data. Modified from (23).

Fig. 9.5. Absorption spectra of ABCA4 containing bound retinoid. ABCA4 was immobilized on Rim 3F4-Sepharose matrix, incubated with all-trans retinal in the presence of PE, and subsequently treated with buffer in the presence or absence of 0.5 mM ATP. The matrix was then washed and ABCA4 was eluted with the 3F4 competing peptide. The absorption spectra were measured in a UV–Vis spectrophotometer. The absorption maximum of bound retinoid was characteristic of unprotonated retinylidene-PE.

to wild-type and mutant ABCA4 expressed in HEK 293 cells (22). For these studies, the 9 amino acid 1D4 epitope tag (TETSQVAPA) was engineered onto the C-terminus of wild-type and mutant ABCA4. ABCA4-1D4 was isolated from CHAPSsolubilized extracts of transfected HEK 293 cells on a Rho-1D4Sepharose immunoaffinity matrix (Fig. 9.2). This procedure has

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been used to investigate the binding of N-retinylidene-PE to C-terminal deletion mutants of ABCA4 which are devoid of the 3F4 epitope (22) (Fig. 9.6). Deletion of the 30 amino acid C-terminus of ABCA4 abolished N-retinylidene-PE binding due to severe protein misfolding, whereas removal of up to 24 amino acids retained substrate binding. The rho 1D4 immunoaffinity tag has been widely used in the detection and localization of heterologously expressed proteins, as well as in the rapid and efficient purification of membrane and soluble proteins from cell extracts for functional characterization and proteomic analysis (24).

Fig. 9.6. Binding of N-retinylidene-PE to ABCA4 C-terminal deletion mutants in the absence and presence of ATP. Wild-type and ABCA4 deletion mutants tagged with the 1D4 epitope were immobilized on a Rho 1D4-Sepharose matrix and incubated with [3 H]-all-trans retinal in the presence of PE. The matrix was washed to remove unbound substrate and incubated in the absence or presence of 0.5 mM ATP. Bound N-retinylidene-PE was eluted with ethanol and quantified by scintillation counting. Data represent the average of three or more experiments ± SD. Modified from (22).

2. Materials 2.1. Retinoid Binding to ABCA4 as Measured by HPLC 2.1.1. Purification of ABCA4 from Rod Outer Segment

1. Column buffer: 50 mM HEPES, pH 7.5, 0.1 M NaCl, 10 mM CHAPS, 1 mM DTT, 3 mM MgCl2 , 10% glycerol, 0.32 mg/ml DOPE, and 0.32 mg/ml DOPC (Avanti Polar Lipid, Alabaster, AL), store at 4◦ C (see Note 1).

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2. Solubilization buffer: 50 mM HEPES, pH 7.5, 0.1 M NaCl, 18 mM CHAPS, 1 mM DTT, 3 mM MgCl2 , 10% glycerol, 0.32 mg/ml DOPE, and 0.32 mg/ml DOPC, store at 4◦ C. 3. Hypotonic buffer: 10 mM HEPES, pH 7.5. 4. Rim 3F4-Sepharose 2B beads, store in equal volume of 20 mM Tris–HCl, pH 8.0, 150 mM NaCl, 0.01% NaN3 at 4◦ C (see Note 2). 5. Rod outer segment preparation (about 8 mg/ml) in 20% (w/v) sucrose, 20 mM Tris–HAc, pH 7.4, 10 mM taurine, 10 mM β-D-glucose, and 0.25 mM MgCl2 , store at −80◦ C (see Note 3). 2.1.2. Binding of Retinoids to ABCA4 and HPLC Analysis

1. The concentration of all-trans retinal (Sigma, St. Louis, MO) in ethanol was determined spectrophotometrically using an extinction coefficient of 42,900 M−1 cm−1 . Stock solution of all-trans retinal (5 mM) in ethanol was added to column buffer to achieve a final concentration of 50 μM or desired concentration (see Note 4). 2. A 1:1 mixture of chloroform and methanol, keep on ice. 3. Mobile phase for reversed-phase HPLC: A: 85% methanol in water, 0.1% trifluoroacetic acid. B: 100% methanol, 0.1% trifluoroacetic acid.

2.1.3. Protein Determination

1. Stock BSA solution (1 mg/ml). 2. SDS-PAGE loading buffer: 8% SDS, 20% glycerol, 0.6 M Tris–HCl, pH 8.8, bromophenol blue.

2.2. Binding of Radiolabeled Retinoid to ABCA4 2.2.1. Titration of All-trans Retinal

1. 50 mM NaOH in water. 2. 1 mg/ml all-trans retinal in ethanol. 3. Mobile phase for normal-phase HPLC: 10% ethyl acetate in hexane.

2.2.2. Purification of 1D4-Tagged ABCA4 Mutants from HEK 293 Cells

1. Rho 1D4-Sepharose 2B beads, store in equal volume of 20 mM Tris–HCl, pH 8.0, 150 mM NaCl, 0.01% NaN3 at 4◦ C (see Note 5).

2.2.3. Binding of Radiolabeled Retinoid and Scintillation Counting

1. 0.5 mM ATP in column buffer.

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3. Methods 3.1. Retinoid Binding to ABCA4 as Measured by HPLC 3.1.1. Purification of ABCA4 from Rod Outer Segment

1. Equilibrate 0.7 ml of gravity packed Rim 3F4-Sepharose 2B beads in a Bio-Rad Econo column (7.0 mm diameter × 4.0 cm length) with column buffer. 2. Solubilize 12–15 mg of ROS membrane in 10 ml solubilization buffer (see Note 6), stir at 4◦ C for 30 min, and incubate with the Rim 3F4-Sepharose beads at 4◦ C for 60 min (see Note 7). 3. Wash the beads six times with 2 ml of column buffer by low-speed centrifugation in a clinical centrifuge to remove unbound proteins.

3.1.2. Binding of Retinoids to ABCA4 and HPLC Analysis

1. Using the capped column as an incubation tube, mix Rim 3F4-Sepharose 2B beads containing immobilized ABCA4 with 50 μM (or desired concentration) all-trans retinal or other retinoid in 2 ml of column buffer for 30 min at 4◦ C on a rotating wheel. 2. Wash column five times with 2 ml of column buffer by centrifugation (see Note 8). 3. Resuspend the column matrix with 2 ml of column buffer in the presence or absence of 0.5 mM ATP (or another nucleotide) for 15 min to determine the effect of ATP on retinoid binding. 4. Wash the column to remove unbound retinoid and transfer contents to a glass test tube. 5. Add 2 ml of ice-cold chloroform–methanol mixture to resuspended column matrix and mix by pipetting gently up and down without generating bubbles. 6. Add 2 ml of ice-cold hexane to the mixture and mix. Centrifuge the solution for 3–5 min to generate a phase separation and carefully remove the upper hexane phase. 7. Repeat the extraction procedure two times and pool the hexane phases from the three extractions. 8. Re-extract pooled hexane phases with 1.5 ml of ice-cold distilled and deionized water. 9. Dry and seal the hexane phase under nitrogen and store overnight in the dark at −80◦ C.

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10. Resuspend dried sample in 300–400 μl of ice-cold methanol and load 50–100 μl on Phenomenex Primesphere 5 C18 HC column (150 mm × 3.2 mm). 11. Elute the HPLC column using a continuous gradient from mobile phase A to B over a period of 30 min, followed by isocratic elution with B at a flow rate of 0.5 ml/min. 12. Identify bound retinoids by comparison of retention time and spectra with standards run on the same column (see Note 9). 3.1.3. Protein Determination

1. Collect immunoaffinity matrix after retinoid extraction and wash extensively with column buffer. 2. Incubate with 500 μl of SDS-PAGE loading buffer without reducing agent for 20 min at room temperature (see Note 10). 3. Add additional 500 μl of SDS-PAGE buffer and repeat elution procedure. 4. Combine eluates, add β-mercaptoethanol (final concentration 2%), and run on an 8% SDS-polyacrylamide gel, together with BSA standards of known concentrations (100 ng–1 μg). 5. Stain gel with Coomassie blue and quantify the density of protein bands on a LICOR infrared imager. 6. Generate standard curve from BSA standards and determine the concentration of ABCA4.

3.2. Radiolabeled Binding Assay 3.2.1. Titration of All-trans Retinal

1. Mix [3 H]NaBH4 (5 mCi, 0.33 μmol, American Radiolabeled Chemicals, St. Louis, MO) in 100 μl of 50 mM NaOH with 0.13 ml of 1 mg/ml all-trans retinal in ethanol and incubate at room temperature for 15 min in a capped tube (see Note 11). 2. Add 400 μl of 50 mM NaOH in water and 600 μl ethanol to bring up volume. 3. Extract the mixture with 1 ml of hexane and collect the upper hexane phase. Repeat the extraction procedure. 4. Combine the two extractions and mix with 30 mg MnO2 and stir the contents for 15 min at 37◦ C. 5. Centrifuge down the MnO2 particles and dry down hexane solution to 200 μl. 6. Inject 100 μl of sample at a time onto a Supelcosil LCSi column (15 cm × 4.6 mm, 3 μl particle size, Supelco

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Park, Bellefonte, PA) and elute isocratically at a flow rate of 1 ml/min with 10% ethyl acetate in hexane for 15 min. Collect fractions containing [3 H]all-trans retinal (see Note 12). 7. Dry down fractions containing [3 H]all-trans retinal under N2 and re-dissolve in ethanol. 8. Mix purified [3 H]all-trans retinal with cold all-trans retinal to achieve a final concentration of 1 mM and specific activity of 500–1000 pm/pmol. Store ethanol solution at −30◦ C. 3.2.2. Purification of 1D4-Tagged ABCA4 Mutants from HEK 293 Cells

1. Equilibrate 12.5 μl Rho 1D4-Sepharose 2B beads in a 500 μl microcentrifuge tube with column buffer (see Note 13). 2. Solubilize transfected HEK 293 T cells from one 10-cm Petri dish in 0.5 ml of solubilization buffer, stir at 4◦ C for 30 min, and incubate with beads at 4◦ C for 30 min. 3. Wash the beads in microcentrifuge tube two times with 0.4 ml of column buffer by spinning down the beads and aspirating off supernatant to remove unbound protein.

3.2.3. Binding of Radiolabeled Retinoid and Scintillation Counting

1. Dilute 1 mM [3 H]all-trans retinal ethanol stock 1:100 in 0.25 ml column buffer (10 μM final concentration, 2.5 × 106 dpm total activity) and mix with Rho 1D4-Sepharose 2B beads containing immobilized 1D4-tagged ABCA4 mutants at 4◦ C for 30 min in microcentrifuge tube (see Note 14). 2. Wash the matrix four times with 0.4 ml column buffer to remove unbound [3 H] all-trans retinal. 3. Incubate the matrix in the presence or absence of 0.5 mM ATP in 0.4 ml of column buffer for 15 min. 4. Resuspend matrix in 0.4 ml column buffer and transfer into an Amicon Ultrafree MC 0.45 μm centrifugal filter device (Millipore, Billerica, MA). 5. Wash three more times in the filter device by low-speed centrifugation. 6. Extract bound [3 H] N-retinylidene-PE by incubation with 0.5 ml ice-cold ethanol for 15 min at room temperature in the filter device. 7. Mix the 0.5 ml eluate with 2 ml of scintillation fluid and count in a liquid scintillation counter (see Note 15).

4. Notes 1. Resuspend CHAP, DOPE, and DOPC in small volume of water and sonicate in a water bath sonicator for about 2 h to dissolve the lipid mixture before addition of other

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ingredients. All buffers should be made fresh or stored for only short periods of time at 4◦ C. DTT should be made fresh and added to the buffers on the same day as the experiment. 2. Rim 3F4 monoclonal antibody can be obtained from Santa Cruz Biotechnology, Inc., Santa Cruz, CA, or PhosphoSolutions, Aurora, CO. 3. Rod outer segments are typically purified from freshly dissected or frozen bovine retina by sucrose gradient centrifugation (25). 4. Final concentration of ethanol should be less than 2%. 5. Rho 1D4 monoclonal antibody can be obtained from Millipore/Chemicon, Billerica, MA, or http://www. flintbox.com/. 6. Resuspend ROS membranes in small volume hypotonic buffer and add dropwise to solubilization buffer with constant stirring. 7. All procedures are carried out under dim red light. Wrap tubes in aluminum foil during incubations. 8. When washing immunoaffinity columns by centrifugation, it is important to centrifuge for only a short time at low speed so that the immunoaffinity matrix does not dry out. 9. To ensure the specificity of retinoid binding to ABCA4, several controls are performed: (1) buffer without both solubilized ROS and retinoid substrate; (2) buffer with the retinoids substrate, but without solubilized ROS; and (3) buffer with solubilized ROS, but without retinoid substrate were added to separate immunoaffinity columns. No detectable retinoid compounds were extracted from immunoaffinity matrix in these control samples. 10. When eluting ABCA4 from the immunoaffinity matrix with SDS loading buffer, β-mercaptoethanol or DTT should not be present as these reducing agents will release the immunoglobulin from the immunomatrix resulting in additional immunoglobulin protein bands. 11. This reaction must be run with an appropriate trap because tritium gas is produced during the reaction. 12. All-trans retinal (λmax = 368 nm) elutes earlier than alltrans retinol (λmax = 325 nm). 13. The higher sensitivity of the radiolabeled assay allows the usage of much less immunoaffinity matrix (1/50 of that for HPLC method) and makes it possible to test limited amounts of protein heterologously expressed in one or two 10-cm Petri dishes of transfected HEK 293 cells.

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14. Incubation and washing is carried out in a microcentrifuge tube before the transfer of the matrix to an Amicon Ultrafree MC 0.45 μm centrifugal filter device for elution. This minimizes background caused by non-specific binding of [3 H]all-trans retinal to the membrane in the filter device. 15. 1D4-tagged Na/K ATPase or another membrane protein is treated in the same way as 1D4-tagged ABCA4 and used as a control for background retinoid binding. The counts in the control sample, typically about 20% of the test sample, are subtracted from the test samples to determine specific retinoid binding. References 1. Illing, M., Molday, L.L., Molday, R.S. (1997) The 220-kDa rim protein of retinal rod outer segments is a member of the ABC transporter superfamily. J. Biol. Chem. 272, 10303–10310. 2. Allikmets, R., Singh, N., Sun, H., et al. (1997) A photoreceptor cell-specific ATPbinding transporter gene (ABCR) is mutated in recessive Stargardt macular dystrophy. Nat. Genet. 15, 236–246. 3. Papermaster, D.S., Schneider, B.G., Zorn, M.A., Kraehenbuhl, J.P. (1978) Immunocytochemical localization of a large intrinsic membrane protein to the incisures and margins of frog rod outer segment disks. J. Cell Biol. 78, 415–425. 4. Weng, J., Mata, N.L., Azarian, S.M., Tzekov, R.T., Birch, D.G., Travis, G.H. (1999) Insights into the function of rim protein in photoreceptors and etiology of Stargardt’s Disease from the phenotype in abcr knockout mice. Cell 98, 13–23. 5. Sun, H., Molday, R.S., Nathans, J. (1999) Retinal stimulates ATP hydrolysis by purified and reconstituted ABCR, the photoreceptorspecific ATP-binding cassette transporter responsible for Stargardt disease. J. Biol. Chem. 274, 8269–8281. 6. Molday, R.S. (2007) ATP-binding cassette transporter ABCA4: Molecular properties and role in vision and macular degeneration. J. Bioenerg. Biomembr. 39, 507–517. 7. Molday, L.L., Rabin, A.R., Molday, R.S. (2000) ABCR expression in foveal cone photoreceptors and its role in Stargardt macular dystrophy. Nat. Genet. 25, 257–258. 8. Molday, R.S., Zhong, M., Quazi, F. (2009) The role of the photoreceptor ABC transporter ABCA4 in lipid transport and Stargardt macular degeneration. Biochim. Biophys. Acta. 179, 573–583.

9. Allikmets, R. (2000) Simple and complex ABCR: Genetic predisposition to retinal disease. Am. J. Hum. Genet. 67, 793–799. 10. Martinez-Mir, A., Paloma, E., Allikmets, R., et al. (1998) Retinitis pigmentosa caused by a homozygous mutation in the Stargardt disease gene ABCR. Nat. Genet. 18, 11–12. 11. Maugeri, A., Klevering, B.J., Rohrschneider, K., et al. (2000) Mutations in the ABCA4 (ABCR) gene are the major cause of autosomal recessive cone-Rod dystrophy. Am. J. Hum. Genet. 67, 960–966. 12. Cideciyan, A.V., Swider, M., Aleman, T.S., et al. (2009) ABCA4 disease progression and a proposed strategy for gene therapy. Hum. Mol. Genet. 18, 931–941. 13. Allikmets, R., Shroyer, N.F., Singh, N., et al. (1997) Mutation of the Stargardt disease gene (ABCR) in age-related macular degeneration. Science 277, 1805–1807. 14. Mata, N.L., Tzekov, R.T., Liu, X., Weng, J., Birch, D.G., Travis, G.H. (2001) Delayed dark-adaptation and lipofuscin accumulation in abcr+/– mice: Implications for involvement of ABCR in age-related macular degeneration. Invest. Ophthalmol. Vis. Sci. 42, 1685–1690. 15. Delori, F.C., Staurenghi, G., Arend, O., Dorey, C.K., Goger, D.G., Weiter, J.J. (1995) In vivo measurement of lipofuscin in Stargardt’s disease – Fundus flavimaculatus. Invest. Ophthalmol. Vis. Sci. 36, 2327–2331. 16. Mata, N.L., Weng, J., Travis, G.H. (2000) Biosynthesis of a major lipofuscin fluorophore in mice and humans with ABCRmediated retinal and macular degeneration. Proc. Natl. Acad. Sci. USA 97, 7154–7159. 17. Parish, C.A., Hashimoto, M., Nakanishi, K., Dillon, J., Sparrow, J. (1998) Isolation and one-step preparation of A2E and iso-A2E,

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Zhong and Molday fluorophores from human retinal pigment epithelium. Proc. Natl. Acad. Sci. USA 95, 14609–14613. Ben-Shabat, S., Parish, C.A., Vollmer, H.R., et al. (2002) Biosynthetic studies of A2E, a major fluorophore of retinal pigment epithelial lipofuscin. J. Biol. Chem. 277, 7183–7190. Eldred, G.E., Lasky, M.R. (1993) Retinal age pigments generated by self-assembling lysosomotropic detergents. Nature 361, 724–726. Ahn, J., Wong, J.T., Molday, R.S. (2000) The effect of lipid environment and retinoids on the ATPase activity of ABCR, the photoreceptor ABC transporter responsible for Stargardt macular dystrophy. J. Biol. Chem. 275, 20399–20405. Sun, H., Smallwood, P.M., Nathans, J. (2000) Biochemical defects in ABCR protein variants associated with human retinopathies. Nat. Genet. 26, 242–246.

22. Zhong, M., Molday, L.L., Molday, R.S. (2009) Role of the C terminus of the photoreceptor ABCA4 transporter in protein folding, function, and retinal degenerative diseases. J. Biol. Chem. 284, 3640–3649. 23. Beharry, S., Zhong, M., Molday, R.S. (2004) N-retinylidene-phosphatidylethanolamine is the preferred retinoid substrate for the photoreceptor-specific ABC transporter ABCA4 (ABCR). J. Biol. Chem. 279, 53972–53979. 24. Wong, J.P., Reboul, E., Molday, R.S., Kast, J. (2009) A carboxy-terminal affinity tag for the purification and mass spectrometric characterization of integral membrane proteins. J. Proteome Res. 8, 2388–2396. 25. Molday, R.S., Molday, L.L. (1987) Differences in the protein composition of bovine retinal rod outer segment disk and plasma membranes isolated by a ricin-gold-dextran density perturbation method. J. Cell Biol. 105, 2589–2601.

Chapter 10 Fluorescence-Based Technique for Analyzing Retinoic Acid Leslie J. Donato and Noa Noy Abstract Retinoic acid (RA) is a potent transcriptional activator whose actions are mediated by members of the nuclear hormone receptor family. In addition to playing key roles in embryonic development and in tissue maintenance in the adult, RA is a potent anticarcinogenic agent currently in clinical use for treatment of various cancers. Here, we describe an optical method for measuring the concentrations of RA in biological samples. This method uses cellular retinoic acid-binding protein I (CRABP-I), a protein that binds RA with high affinity and specificity, as a “read-out” for its ligand. Replacing 28 Leu of CRABP-I with a Cys residue allows for covalently attaching an environmentally sensitive fluorescent probe to the protein at a region that undergoes a significant conformational change upon ligand binding. Association of RA with the modified protein thus results in changes in the fluorescence of the probe, enabling reliable measurements of RA concentrations as low as 50 nM. We show that the method can be effectively used to measure RA concentrations in serum and to monitor the biosynthesis and the degradation of RA in cultured mammalian cells. Key words: Retinoic acid, intracellular lipid-binding proteins, retinoic acid-binding protein, equilibrium dissociation constant, 5-bromomethyl fluorescein, fluorescence titration, retinoic acid biosynthesis, retinoic acid degradation.

1. Introduction The vitamin A metabolite all-trans-retinoic acid (RA) controls biological functions by virtue of its ability to regulate the rate of transcription of multiple target genes. The transcriptional activities of this hormone are mediated by two ligandactivated transcription factors that are members of the nuclear hormone receptor family, the RA receptor (RAR) and the peroxisome proliferator-activated receptor β/δ (PPARβ/δ) (1, 2). In addition to associating with these nuclear receptors, RA also H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_10, © Springer Science+Business Media, LLC 2010

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binds with sub-nanomolar affinities to two small (∼14 kDa) cytosolic proteins called cellular RA-binding proteins (CRABP-I and CRABP-II) (3). CRABP-II shuttles RA from the cytosol to the nucleus where it directly “channels” it to RAR, thereby facilitating the delivery of the ligand and enhancing the transcriptional activity of the receptor (4, 5). The details of the mechanism of action of CRABP-I are incompletely understood, but it is believed that this protein directs RA to degradation pathways (6, 7). RA plays important roles in embryonic development and in regulating proliferation and differentiation in adult mammals, and it exhibits chemotherapeutic and chemopreventive activities in a number of human cancers, including bladder, liver, lung, pancreas, head and neck, prostate, and breast cancers (8–11). However, usage of RA in chemotherapy is complicated by the pronounced toxicity of this compound at pharmacological doses (12, 13). The pharmacokinetics of RA has been reported to vary widely between different patients, and hence, minimizing toxic side effects while optimizing the efficacy of the drug may be significantly improved by tailoring therapeutic regimes to individual patients. Tailored therapies will require constant monitoring of plasma drug levels. However, usual methodologies for measurement of RA concentrations in biological samples utilize multi-phase organic extraction followed by HPLC and/or mass spectrometry analyses (14, 15) and are too expensive and complicated to be applied in many laboratories and in clinical settings. Consequently, as currently practiced, RA treatment is not individualized but is administered by “standard” dosing. Here we describe a fluorescence-based method that allows for measurements of RA concentrations in biological samples using widely available instrumentation. In this, CRABP-I, a protein that binds RA with a high affinity (4), is used as a “read-out” for its ligand. Association of RA with proteins results in a marked decrease in the intrinsic fluorescence of a protein (4, 16). Protein fluorescence emanates primarily from the fluorescent amino acid residues tryptophan and tyrosine and is characterized by absorption and emission maxima that center around 280 and 340 nm, respectively. As the absorption spectra of RA, peaking at 350 nm, extensively overlap with the fluorescence emission spectra of proteins, RA binding is accompanied by a significant decrease in protein fluorescence. This decrease has been widely used to study retinoid–protein interactions using purified proteins. However, using this method for analyzing retinoids in biological samples is complicated by optical artifacts originating from contaminating biological fluorophores, many of which possess optical properties at short wavelengths, and from the presence of multiple proteins in the samples. In addition, the quantum yield of protein fluorescence is quite low, limiting the sensitivity of the method. These difficulties were bypassed by covalently attaching to CRABP-

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I a synthetic fluorescent probe which displays long-wavelength fluorescence characteristics. The probe was placed at a location on the protein whose conformation is altered upon ligand binding. Hence, the environment of the probe, and thus its fluorescence, changes upon ligand binding and the fluorescence change can be used to monitor the concentration of RA in a sample. Attempts to conjugate a fluorescent moiety to either CRABP-I or CRABP-II using various reagents failed, suggesting that the proteins lack reactive residues that are readily accessible to covalent modifications. To bypass this problem, a CRABP-I mutant that can be labeled efficiently was generated. Inspection of the reported three-dimensional crystal structures of CRABPs in the presence and absence of RA revealed that, while these proteins do not undergo dramatic conformational changes upon ligation, ligand binding results in subtle changes within their helix-loophelix region (17, 18). Notably, the Leu28 side chain appears to be solvent-exposed and to acquire an altered configuration in the holo vs. apo forms of CRABPs (Fig. 10.1). Hence, if a fluorescent moiety can be attached at this location, its environment may change upon ligand binding and such a change may result in altered fluorescence. Leu28 of CRABP-I was replaced by a cysteine residue to generate a CRABP-I-L28C mutant. The fluorescent probe fluorescein was then covalently attached to the mutant using the thiol-reactive agent 5-bromomethyl fluorescein.

Fig. 10.1. X-ray crystal structures of apo- and holo-CRABP-II. Superposition of the three-dimensional structures of apo-CRABP-II ((18), PDB entry 1XCA) and the RAbound protein ((17), PDB entry 1CBS). Apo- and holo-CRABP-II are depicted in gray and black, respectively. The position of Leu28 shifts in response to ligand binding. The helix-loop-helix region of the protein is boxed. Structures were visualized using Pymol (www.pymol.org).

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The equilibrium dissociation constant (Kd ) that characterizes RA binding by the labeled mutant was measured using standard binding assays in which the progress of titrations was monitored by following the decrease in the intrinsic fluorescence of the protein (ex. 280 nm, em. 340 nm, Fig. 10.2a). The RA-binding affinity of the fluorescein-labeled protein (F-CRABP-L28C) was found to be essentially identical to that of the wild-type protein. RA binding to F-CRABP-I-L28C was then monitored by following ligand-induced changes in the fluorescence of the proteinbound fluorescein which was measured at excitation and emission wavelengths of 492 and 519 nm, respectively (Fig. 10.2b). The Kd derived from these measurements was indistinguishable from that obtained using the standard assay. In addition, titrations with a closely related retinoid, retinaldehyde, resembled control vehicle titrations (Fig. 10.2b, inset), indicating that the fluorescence of the probe specifically reports on binding of RA.

Fig. 10.2. Fluorescein-labeled CRABP-I-L28C as a sensor for RA. (a) RA binding was monitored by following the intrinsic fluorescence of the protein (ex. 280 nm, em. 340 nm). (b) Titration followed by monitoring changes the fluorescence of protein-bound probe (ex. 492 nm, em. 520 nm). Inset: F-CRABP-I-L28C was titrated with RA, retinaldehyde (RAL), or ethanol (veh).

F-CRABP-I-L28C was used to measure RA concentrations in serum and in cultured MCF-7 mammary carcinoma cells. Appropriate calibration curves were constructed. To this end, known concentrations of RA were added to serum, or cultured cells were extracted in ethanol containing known concentrations of RA. F-CRABP-I-L28C was then titrated with RA-containing serum or with cell ethanol extracts. Titrations with samples devoid of exogenously added RA did not result in significant fluorescence changes, demonstrating that concentrations of endogenous RA in these samples were below the sensitivity of the method. Initial slopes of titrations with increasing RA concentrations in serum (Fig. 10.3a) or cell extracts (Fig. 10.3c) were plotted as a function of RA concentrations to generate calibration curves (Fig. 10.3b and d).

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Fig. 10.3. Calibration curves for RA in serum and in cells. Fluorescein-labeled CRABP-I-L28C (45 nM) was titrated by consecutive additions of 5 μl of either fetal bovine serum (FBS) (a) or ethanol extracts of MCF-7 cells (c) containing the denoted concentrations of RA. Fluorescence (ex 492 nm, em 520 nm) was recorded after each addition. Initial phases of titrations with standard solutions containing 0.25, 0.5, and 1.0 μM RA are shown. Absolute values of initial slopes (F/μl) were plotted as a function of RA concentration to obtain calibration curves for RA in FBS (b) or in cell extracts (d).

The utility of the method was demonstrated by measurements of the rates of degradation and of biosynthesis of RA in cultured cells. To examine RA degradation, MCF-7 mammary carcinoma cells were treated with 1 μM RA for 1 h to allow for accumulation of ligand. The medium was removed, cells washed, and RA extracted into ethanol at various time points following the removal of the ligand. F-CRABP-I-L28C was titrated with the extracts, and RA concentrations were determined by using a calibration curve and normalized to the concentrations of cell protein. The data (Fig. 10.4a) showed that the process of RA degradation in MCF-7 cells displays a half-time of 10–20 min. Biosynthesis of RA was examined in MCF-7 and in COS-7 cells. Cells were treated with the metabolic precursor of RA, retinaldehyde. At various time points following addition of the substrate, ethanol extracts were obtained and RA concentrations determined. In agreement with the report that MCF-7 cells lack the enzymatic machinery for RA synthesis (19), no RA was found in MCF-7 cell extracts. In contrast, the hormone was readily generated in COS-7 cells, reaching a steadystate concentration of 0.3 nmol RA/mg protein within 200 min (Fig. 10.4b).

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Fig. 10.4. Measurements of RA in cell extracts. (a) RA degradation in MCF-7 cells. Cells were treated with RA for 1 h, RA removed from the medium, and cells extracted in ethanol at the indicated times. (b) RA synthesis in MCF-7 and COS-7 cells. Cells were treated with retinaldehyde and extracted into ethanol at the denoted times. F-CRABP-I-L28C was titrated with each extract and the concentration of RA was obtained from initial slopes of titrations and appropriate calibration curves. Data were normalized to the amount of cellular protein.

2. Materials 2.1. Purification of Recombinant CRABP-I-L28C

1. Bacterial expression vector (pET28a) harboring cDNA of CRABP-I-L28C 2. LB medium: 10 g tryptone, 5 g yeast extract, 10 g NaCl, dissolve in 1 l distilled water, pH 7.0 3. Kanamycin 4. Isopropyl B-D-1-thiogalactopyranoside (IPTG) 5. Lysozyme 6. Binding buffer (BB): 5 mM imidazole, 500 mM NaCl, 20 mM Tris–HCl (pH 8.0), 0.2 mM phenylmethylsulfonyl fluoride (PMSF) 7. HEK buffer: 10 mM HEPES (pH 8.0), 0.1 mM ethylenediaminetetraacetic acid (EDTA), 100 mM KCl 8. Protein concentrator (Centrifugal filter device, Centricon YM 10 membrane, Millipore) 9. Dialysis tubing

2.2. Fluorescent Labeling of Protein

1. 5-(Bromomethyl)fluorescein (BMF) (Molecular Probes/ Invitrogen) dissolved in DMF 2. HEK buffer, pH 7.3 3. Dialysis tubing

Fluorescence-Based Technique for Analyzing Retinoic Acid

2.3. Retinoic Acid Preparation

1. Retinoic acid (CalBiochem)

2.4. Protein Determination

1. 1 M NaOH in dH2 O

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2. BSA standard 0.1 mg/ml in dH2 O 3. Bradford assay reagent (Bio-Rad, Hercules, CA, USA)

3. Methods 3.1. Protein Purification

1. Transform CRABP-I-L28C plasmid into BL21 (DE3) Escherichia coli containing 30 μg/mL kanamycin. Place at 37◦ C overnight 2. Pick a single colony and inoculate to 50 mL start culture containing 30 μg/mL kanamycin in LB medium. Grow at 37◦ C overnight 3. Inoculate 2L LB (containing 30 μg/mL kanamycin). Grow culture at 37◦ C to OD600 nm = 0.6–0.8 4. Induce protein production by addition of 0.5 mM IPTG directly to culture and continue growing for 3 h 5. Harvest bacteria by centrifugation 6. Resuspend bacterial pellet in binding buffer containing 1 mg/ml lysozyme 7. Incubate for 20 min at 4◦ C while stirring 8. Sonicate suspension twice 9. Centrifuge and collect supernatant 10. Rotate supernatant with ∼1 ml of Ni2+ beads (e.g., HiTrap, GE Healthcare, Waukesha, WI, USA) for 2 h at 4◦ C 11. Wash beads sequentially with 5 ml BB, 5 ml binding buffer (BB) containing 50 mM imidazole, 5 ml BB 12. Elute protein with 20 ml BB containing 500 mM imidazole 13. Concentrate eluted protein to 0.2–1 ml 14. Dialyze concentrated protein against HEK buffer 15. Add 50% glycerol (v/v), mix gently, and store at −20◦ C. 16. Measure protein concentration using the Bradford assay (Bio-Rad, Hercules, CA) 17. Resolve protein by SDS-PAGE and stain using Coomassie blue to verify purity

3.2. Fluorescent Labeling of Protein

1. Prepare 1–10 mM stock solution of BMF in DMF (see Note 2). 2. Dilute protein in HEK pH 7.3 to a concentration in the range of 10–50 μM and add BMF (dissolved in DMF)

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at a mole ratio of protein/BMF of 1:3. Keep DMF concentration to a minimum (do not exceed 2% v/v). Protect from light and incubate for 2 h at ambient temperature (see Note 3). 3. Extensively dialyze labeled protein (in the dark) against HEK buffer, pH 7.3, containing 1 mM dithiothreitol (DTT) and 5% glycerol (see Note 4). 4. Measure the concentration of the protein using Bradford assay (Bio-Rad, Hercules, CA). 3.3. Preparation of Retinoic Acid Solutions

1. Dissolve a small amount of lyophilized RA in ethanol. Use spatula to transfer two to three grains from vial into a dark tube, add 0.5–2 ml ethanol, and mix until dissolved (see Note 1). 2. Determine the RA concentration by measuring the absorbance of the solution at 350 nm and using the extinction coefficient 45,300 M−1 . The stock solution is likely to be too concentrated for direct measurement. To measure, dilute in ethanol (typically 1:10). To ensure that the absorption maximum is indeed at 350 nm and to subtract background, an absorption spectrum in the 250–450 nm range should be obtained rather than a single wavelength measurement. Note: If the OD of the solution at 350 nm is higher than the linear range of the spectrophotometer (usually 1–1.5), measure using a more diluted solution. The concentration of RA is OD350 nm × dilution factor/45,300 M.

3.4. Fluorescence Titrations

To verify that the protein is viable and properly reports on RA binding, titrate with RA. 1. Place a stoppered cuvette containing F-CRABP-I-L28C (0.1–1 μM in HEK buffer, pH 7.3, final volume 1 ml) in the sample chamber of a spectrofluorometer. Measure fluorescence at excitation and emission wavelengths of 492 and 519 nm, respectively. 2. Aim to complete a titration within 10–15 steps. RA is added from an ethanolic stock solution in 1–2 μl increments. Hence, use a RA solution which is approximately 100-fold higher than the concentration of the protein in the cuvette. A 1 μl addition of RA from such a solution will yield an RA concentration corresponding to 10% of the protein. Saturation will thus be expected following 10 such additions. Mix RA and protein by inverting the cuvette two to three times and measure the fluorescence after each addition. Fluorescence will decrease upon titration until a plateau is reached at saturation (see Fig. 10.2b). Titration curves can be analyzed to yield the number of binding sites and Kd of the labeled protein (16). Note: Protein preparations that display

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a Kd in the 1–30 nM range may be used for subsequent measurement. Importantly, a linear initial phase should be observed, reflecting high-affinity binding. 3.5. Quantitation of Retinoic Acid in Cultured Cells 3.5.1. Construction of Standard Curve

1. Grow cells in 100 mm plates until approximately 80% confluence. 2. Remove medium, wash cells with PBS. 3. Scrape cells into 1 ml PBS. 4. Pellet cells gently and remove supernatant. 5. To cell pellet, add 100 μl ethanol containing RA in the range of 0–10 μM, pipet up and down 20 times. 6. Pellet cell debris and collect ethanol supernatant. Supernatant will be used for titrations. 7. Fluorescence titrations. Place 0.05–0.2 μM F-CRABP-IL28C in a cuvette and titrate consecutively with 2–5 μl ethanol extract. Following each step, measure fluorescence at excitation and emission wavelengths of 492 and 519 nm, respectively. 8. For each RA extract, plot fluorescence as a function of volume added, normalizing to the initial fluorescence (see Fig. 10.3a and c). Compute initial slope for each extract. 9. Calibration curve. Plot the initial slope of each ethanol extract as a function of the concentration of RA (see Fig. 10.3b and d).

3.5.2. Retinoic Acid Synthesis

1. Seed 350,000 cells in a 60 mm cell culture plate. 2. Once cells have attached, remove medium and wash cells with PBS. 3. Replace medium with serum-free medium containing desired concentration of retinol or retinal. 4. At desired time points, remove medium, wash cells with PBS, and scrape into 1 ml PBS. 5. Pellet cells gently and remove supernatant. 6. Resuspend cell pellet in 100 μl ethanol. Pipet up and down 20 times. 7. Pellet cell debris. Collect supernatant for RA measurements. 8. Add 0.5 ml 1 M NaOH to cell pellet. Incubate for at least 5 h. 9. Measure protein concentration in cell pellet using Bradford assay.

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10. Fluorescence titrations. Place 0.15–0.3 μM F-CRABP-IL28C in a cuvette and titrate consecutively with 2–5 μl of each ethanol extract. Following each step, measure fluorescence at excitation and emission wavelengths of 492 and 519 nm, respectively. 11. For each extract, plot fluorescence as a function of volume added, normalizing to the initial fluorescence (see Fig. 10.3a and c). Compute an initial slope for each extract. 12. Use the calibration curve (Section 2.5.1) to obtain the concentration of RA in each extract. 13. Normalize RA concentrations to cell protein. 3.5.3. Retinoic Acid Degradation

1. Seed 350,000 cells in a 60 mm cell culture plate 2. Treat cells with desired concentration of RA 3. Remove medium 4. Wash cells with PBS 5. Replace medium with serum-free medium 6. At desired time points, remove medium, wash cells with PBS, and scrape into 1 ml PBS 7. Pellet cells gently and discard supernatant 8. Resuspend cell pellet in 100 μl ethanol. Pipet up and down 20 times 9. Pellet cell debris. Collect supernatant for RA measurements 10. Add 0.5 ml 1 M NaOH to cell pellet. Incubate for at least 5h 11. Measure protein concentration in cell pellet using Bradford assay 12. Use ethanol extracts for RA measurements as in Section 2.5.2

4. Notes 1. To minimize photodegradation and oxidation, protect solutions containing retinoids and fluorescent probes from light or use light with a cutoff of 400–420 nm. Bubble argon or nitrogen through buffers prior to use. 2. Stock RA and BMF solutions should be made fresh prior to each experiment. 3. Avoid over-labeling the protein with the fluorescent probe. The mole ratio of label/protein should be 0.5–0.8.

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4. F-CRABP-I-L28C should not be stored for longer than a few days. References 1. Chambon, P. (1996) A decade of molecular biology of retinoic acid receptors. FASEB. J. 10, 940–954. 2. Schug, T.T., Berry, D.C., Shaw, N.S., Travis, S.N., Noy, N. (2007) Opposing effects of retinoic acid on cell growth result from alternate activation of two different nuclear receptors. Cell 129, 723–733. 3. Noy, N. (2000) Retinoid-binding proteins: Mediators of retinoid action, Biochem. J. 348(Pt 3), 481–495. 4. Dong, D., Ruuska, S.E., Levinthal, D.J., Noy, N. (1999) Distinct roles for cellular retinoic acid-binding proteins I and II in regulating signaling by retinoic acid. J. Biol. Chem. 274, 23695–23698. 5. Budhu, A.S., Noy, N. (2002) Direct channeling of retinoic acid between cellular retinoic acid-binding protein II and retinoic acid receptor sensitizes mammary carcinoma cells to retinoic acid-induced growth arrest. Mol. Cell. Biol. 22, 2632–2641. 6. Boylan, J.F., Gudas, L.J. (1991) Overexpression of the cellular retinoic acid binding protein-I (CRABP-I) results in a reduction in differentiation-specific gene expression in F9 teratocarcinoma cells. J. Cell Biol. 112, 965–979. 7. Boylan, J.F., Gudas, L.J. (1992) The level of CRABP-I expression influences the amounts and types of all- trans-retinoic acid metabolites in F9 teratocarcinoma stem cells. J. Biol. Chem. 267, 21486–21491. 8. Soprano, D.R., Qin, P., Soprano, K.J. (2004) Retinoic acid receptors and cancers. Annu. Rev. Nutr. 24, 201–221. 9. Hong, W.K., Itri, L. (1994) Retinoids and human cancer. In: Sporn, M.B., Roberts, A.B., Goodman, D.S. (eds.), The Retinoids: Biology, Chemistry, and Medicine, Raven Press, New York, pp. 597–630. 10. Lotan, R. (1996) Retinoids in cancer chemoprevention. FASEB. J. 10, 1031–1039. 11. Yang, L.M., Tin, U.C., Wu, K., Brown, P. (1999) Role of retinoid receptors in the

12.

13.

14.

15.

16.

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prevention and treatment of breast cancer. J. Mammary Gland Biol. Neoplasia. 4, 377–388. Miller, W. H., Jr. (1998) The emerging role of retinoids and retinoic acid metabolism blocking agents in the treatment of cancer. Cancer 83, 1471–1482. Frankel, S.R., Eardley, A., Lauwers, G., Weiss, M., Warrell, R.P., Jr. (1992) The “retinoic acid syndrome” in acute promyelocytic leukemia. Ann. Intern. Med. 117, 292–296. De Leenheer, A.P., Lambert, W.E., Claeys, I. (1982) All-trans-retinoic acid: Measurement of reference values in human serum by high performance liquid chromatography. J. Lipid Res. 23, 1362–1367. Kane, M.A., Folias, A.E., Wang, C., Napoli, J.L. (2008) Quantitative profiling of endogenous retinoic acid in vivo and in vitro by tandem mass spectrometry. Anal. Chem. 80, 1702–1708. Norris, A.W., Cheng, L., Giguere, V., Rosenberger, M., Li, E. (1994) Measurement of subnanomolar retinoic acid binding affinities for cellular retinoic acid binding proteins by fluorometric titration. Biochim. Biophys. Acta 1209, 10–18. Kleywegt, G.J., Bergfors, T., Senn, H., Le Motte, P., Gsell, B., Shudo, K., Jones, T.A. (1994) Crystal structures of cellular retinoic acid binding proteins I and II in complex with all-trans-retinoic acid and a synthetic retinoid. Structure 2, 1241–1258. Chen, X., Tordova, M., Gilliland, G.L., Wang, L.C., Li, Y., Yan, H.G., Ji, X.H. (1998) Crystal structure of apo-cellular retinoic acid-binding protein type II (R111M) suggests a mechanism of ligand entry. J. Mol. Biol. 278, 641–653. Mira, Y.L.R., Zheng, W.L., Kuppumbatti, Y.S., Rexer, B., Jing, Y., Ong, D.E. (2000) Retinol conversion to retinoic acid is impaired in breast cancer cell lines relative to normal cells. J. Cell Physiol. 185, 302–309.

Chapter 11 The Interaction Between Retinol-Binding Protein and Transthyretin Analyzed by Fluorescence Anisotropy Claudia Folli, Roberto Favilla, and Rodolfo Berni Abstract The retinol carrier retinol-binding protein (RBP) forms in blood a complex with the thyroid hormone carrier transthyretin (TTR). The interactions of retinoid–RBP complexes, as well as of unliganded RBP, with TTR can be investigated by means of fluorescence anisotropy. RBP represents the prototypic lipocalin, in the internal cavity of which the retinol molecule is accommodated. Due to the tight binding of retinol within a substantially apolar binding site, an intense fluorescence emission characterizes the RBP-bound vitamin. The addition of TTR to the retinol–RBP complex (holoRBP) causes a marked increase in the fluorescence anisotropy of the RBP-bound retinol within the system, due to the formation of the holoRBP–TTR complex, which allows the interaction between the two proteins to be monitored. The fluorescence anisotropy technique is also suitable to study the interaction of TTR with apoRBP and RBP in complex with non-fluorescent retinoids. In the latter cases, the fluorescence signal is provided by a fluorescent probe covalently linked to TTR rather than by RBP-bound retinol. We report here on the preparation of recombinant human RBP and TTR, the covalent labeling of TTR with the fluorescent dansyl probe, and fluorescence anisotropy titrations for RBP and TTR. Key words: Retinol-binding protein, transthyretin, fluorescence, fluorescence anisotropy, protein–protein interactions, macromolecular complex, vitamin A.

1. Introduction Natural retinoids need to be bound to specific retinoid-binding proteins to be protected, solubilized, and transported in body fluids and within the cell. The physiologically occurring prototypic retinoid is retinol (vitamin alcohol) (Fig. 11.1). The transport of retinol in blood and its delivery to recently identified specific cell surface receptors (1) are uniquely accomplished H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_11, © Springer Science+Business Media, LLC 2010

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Fig. 11.1. Structural formulae of all-trans retinol (a) and fenretinide (b).

by retinol-binding protein (RBP, also designated RBP4), a monomeric protein of 21 kDa belonging to the lipocalin family (2). RBP is synthesized primarily in the liver, the major storage site of vitamin A, where its secretion into the general circulation is triggered by the binding of retinol (3). RBP is a singledomain protein which represents the prototypic lipocalin, being the first lipocalin for which an X-ray structure was described (4, 5). Most characterizing in the RBP structure is an eightstranded up-and-down β-barrel, in the internal cavity of which the retinol molecule is accommodated. Mainly due to the tight binding of retinol within a substantially apolar binding site (5), an intense fluorescence emission characterizes the RBP-bound vitamin (Fig. 11.2). RBP circulates in the blood of terrestrial vertebrates bound to another transport protein, transthyretin (TTR), a transporter of thyroid hormones (thyroxine and triiodothyronine). TTR has been associated with human diseases; in fact, it is one of a number of proteins that can produce the extracellular accumulation in tissues of protein aggregates responsible for degenerative diseases known as amyloidoses, which are especially caused by a large number of amyloidogenic mutations in the case of TTR (6). TTR is a tetrameric protein of 55 kDa, formed by the assembly of four chemically identical subunits, whose structure is known at high resolution (7, 8). The crystal structures of heterologous (human TTR – chicken RBP (9)) and homologous (human TTR – human RBP (10–11)) TTR–holoRBP complexes,

Fluorescence Intensity (arbitrary units)

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Fig. 11.2. Absorption (a) and fluorescence emission (b) spectra of holoRBP prepared as described in Section 3.3. The excitation wavelength for the fluorescence emission spectrum was 330 nm.

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both characterized by a 1:2 TTR:RBP stoichiometry, have been determined. Accordingly, binding data in solution have indicated that a maximum of two RBP molecules can be bound by one TTR tetramer for both human (12, 13) and chicken (14) RBP–TTR complexes. Despite the high symmetry of TTR, consistent with four virtually identical binding sites for RBP, the binding of two RBP molecules to the TTR tetramer partially hinders the potential binding of two nearby RBP molecules, thereby limiting the possible interactions with tetrameric TTR to two RBP molecules (9). It should be noted, however, that a 1:1 TTR:RBP complex is normally present in human plasma due to a concentration of TTR significantly higher than that of RBP (3). The association of RBP with TTR increases the stability of the retinol–RBP complex (15, 16), consistent with the opening of the RBP β-barrel cavity and bound retinol being totally buried within the holoRBP–TTR complex (9–11). Moreover, the association of RBP with TTR is believed to reduce the glomerular filtration of the relatively small RBP molecule (21 kDa), by forming a complex of 76 kDa (17). In turn, the stability of the RBP–TTR complex is strongly affected by the presence of retinol bound to RBP within the complex, a feature that is also believed to be of physiological significance. In fact, the affinity of holoRBP for TTR (Kd ≈ 0.3 μM (11, 14) (Fig. 11.3a)) is significantly higher than that of apoRBP (Kd ≈ 1.2 μM (14) (Fig. 11.3b)), in keeping with the retention of holoRBP in the

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Fig. 11.3. (a) Representative fluorescence anisotropy titration of human holoRBP (3 μM) with human TTR (filled triangles). Fluorescence anisotropy (excitation and emission at 330 and 460 nm, respectively) values are plotted as a function of human TTR concentration. The line represents a theoretical binding curve corresponding to a dissociation constant of 0.34 μM and a stoichiometry of 2.0 RBP:1 TTR (see Section 3.7). The fluorescence anisotropy values for the titration of holoRBP with the amyloidogenic Ile84Ser TTR variant, prepared according to (25), are also shown (open triangles), revealing a negligible binding affinity of the TTR variant for holoRBP. (b) Representative fluorescence anisotropy titrations of human DNS-TTR (4 μM) with human apoRBP (filled triangles) and with the fenretinide–RBP complex (open triangles). Fluorescence anisotropy (excitation and emission at 380 and 480 nm, respectively) values are plotted as a function of RBP concentration. The line represents a theoretical binding curve corresponding to a dissociation constant of 1.2 μM and a stoichiometry of 1.8 RBP:1 TTR (see Section 3.7).

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circulation as the protein–protein complex and with the clearance from the circulation by glomerular filtration of the uncomplexed apoRBP molecule resulting from the delivery of retinol. Indeed, the direct participation, through H-bond interaction, of the RBPbound retinol hydroxyl end group in the binding of TTR has been established (9–11). Accordingly, an important role played by the retinol hydroxyl end group in protein–protein interactions is revealed by the observation that the replacement of RBP-bound retinol by synthetic retinoids affects RBP–TTR recognition to an extent that appears to be well correlated with the nature and steric hindrance of the groups substituting the retinol hydroxyl group (14, 18–20). In particular, a drastic interference with RBP–TTR interactions has been demonstrated for RBP-bound fenretinide, a pharmacologically active retinoid bearing a bulky end group in place of the retinol hydroxyl group (14, 20) (Figs. 11.1 and 11.3b). Moreover, the conformational change affecting one of the loops (in particular, residues Leu35 and Phe36 (21)) surrounding the opening of the β-barrel cavity in apoRBP relative to holoRBP is likely to contribute to the weakening of the interaction of apoRBP with TTR, due to the involvement of such a loop in RBP–TTR recognition (9–11). The relatively weak interaction between holoRBP and TTR (Kd ≈ 0.3 μM) is possibly correlated with the need for the presence in plasma of a small but significant amount of uncomplexed holoRBP, which can thus leave more easily the circulation to deliver the retinol to the target tissues (3, 22). The main RBP–TTR interactions, both polar and apolar, involve the retinol hydroxyl group and a limited number of solvent-exposed residues (9–11). The relevance of TTR residues in complex formation with RBP has been examined by mutational analysis, and the structural consequences of some TTR point mutations affecting protein–protein recognition have been investigated (11). Despite a few exceptions, in general the substitution of a hydrophilic for a hydrophobic side chain in contact regions results in decrease or even loss of binding affinity, thus revealing the importance of interfacial hydrophobic interactions and a high degree of complementarity between RBP and TTR (11). Remarkably, the amyloidogenic Ile84Ser TTR mutation, which affects a residue that is crucial for protein–protein interactions, results in the lack of recognition between RBP and TTR and in an altered plasma transport of RBP by TTR (13) (Fig. 11.3a). Finally, the lack of binding affinity between piscine RBP and TTR has been established (23), in accordance with the presence of relevant amino acid differences in piscine TTR relative to the TTR of terrestrial vertebrates that affect regions involved in RBP–TTR recognition (11). The above-described RBP–TTR interactions have mostly been analyzed by the use of the fluorescence anisotropy

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technique. The addition of TTR to the retinol–RBP complex causes a substantial increase in the fluorescence anisotropy of the RBP-bound retinol within the system, due to the formation of the holoRBP–TTR complex, which allows the interaction between the two proteins to be monitored (24) (Fig. 11.3a). The technique has also proved suitable to study the interaction of TTR with apoRBP and RBP in complex with non-fluorescent retinoids (14). In the latter cases, the fluorescence signal was provided by the fluorescent dansyl (DNS) probe covalently linked to TTR rather than by RBP-bound retinol (Fig. 11.3b). We report here on the preparation of recombinant human RBP and TTR and the covalent labeling of TTR with the DNS probe. Fluorescence anisotropy titrations for RBP and TTR are described.

2. Materials 2.1. Heterologous Expression, Unfolding/Refolding, and Purification of Human RBP 2.1.1. Cloning the cDNA Sequence Coding for Human RBP

1. cDNA sequence: EST sequence BQ645928 encoding for human complete sequence of RBP from American Type Culture Collection (LGC Promochem, Milan, Italy). 2. PCR amplification: MasterTaq DNA polymerase (Eppendorf, Hamburg, Germany), NdeI-tailed upstream primer, and BamHI-tailed downstream primer (MWG, Ebersberg, Germany). 3. T/A cloning: pGEM-T-Easy Vector system (Promega, Madison, WI, USA). 4. Expression vector: pET11b (Novagen, Madison, WI, USA). 5. Isolation of the coding DNA fragment and linearization of pET11b vector: NdeI and BamHI restriction enzymes (New England Biolabs, Beverly, MA, USA). 6. Elution and purification of the restriction fragments: Gel Extraction Kit (QIAGEN, Hilden, Germany). 7. Cloning in the expression vector: T4 DNA Ligase (USB, Staufen, Germany). 8. Transformation of Escherichia coli: MiniPulser Electroporation System (Bio-Rad, Hercules, CA, USA).

2.1.2. Expression and Purification of Human RBP

1. Expression system: E. coli Origami (DE3) cells (Stratagene, La Jolla, CA, USA) transformed with the pET11b vector containing the mature human RBP cDNA.

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2. Luria broth: 1% (w/v) tryptone, 0.5% (w/v) yeast extract, and 1% (w/v) NaCl. Adjust the pH to 7.0 with 2 M NaOH and autoclave. 3. Antibiotic solution: 5% (w/v) ampicillin (Sigma, St. Louis, MO, USA) in sterile water, 0.5% (w/v) tetracycline (Sigma) in ethanol, and 10% (w/v) kanamycin (Sigma) in sterile water. Store at −20◦ C. 4. Isopropyl β-D-1-thiogalactopyranoside (IPTG): 0.1 M IPTG (Inalco, Milan, Italy) solution prepared just before use in sterile water. 5. Lysis buffer: 0.05 M sodium phosphate (pH 7.2), 0.3 M NaCl, 10% (v/v) glycerol, 1 mM 2-mercaptoethanol, 1 μM pepstatin, 1 μM leupeptin, 1 μM PMSF. 6. Denaturing buffer: 0.05 M Tris–HCl (pH 7.5), 2 mM EDTA, 2 mM PMSF, 0.1% (v/v) Triton X-100, 8 M urea. 7. Refolding buffer: 0.05 M Tris–HCl (pH 9.3). 8. Concentrating cell: Ultrafiltration Amicon Cell equipped with a YM 10 membrane (Amicon, Beverly, MA, USA). 9. Gel filtration chromatography: Bio-gel P-60 (Bio-Rad) equilibrated and developed with 0.1 M ammonium sulfate, 0.05 M Tris–HCl (pH 7.4). 10. Affinity chromatography: Sepharose 4B (GE Healthcare Biosciences, Milan, Italy) coupled with human TTR as described (26), equilibrated with 0.05 M sodium phosphate buffer (pH 7.4), 0.15 M NaCl, and eluted with 1 mM sodium phosphate (pH 7.0). 11. Protein concentration: Centricon centrifugal filter device equipped with YM-10 membrane (Millipore, Milan, Italy). 2.2. Heterologous Expression and Purification of Human TTR 2.2.1. Cloning the cDNA Sequence Coding for Human TTR

2.2.2. Expression and Purification of Human TTR

Materials are those reported in Section 2.1.1, except for the following: 1. cDNA sequence: the vector pcDNA3 containing the sequence encoding for mature human TTR was a kind gift from D. Bellovino (INRAN, Rome, Italy). 1. Expression system: E. coli BL21 (DE3) cells (Stratagene) transformed with the pET11b vector containing the mature human RBP cDNA.

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2. Luria broth: 1% (w/v) tryptone, 0.5% (w/v) yeast extract, and 1% (w/v) NaCl. Adjust the pH to 7.0 with 2 M NaOH and autoclave. 3. Ampicillin solution: 5% (w/v) ampicillin (Sigma) in sterile water. Store at −20◦ C. 4. Isopropyl β-D-1-thiogalactopyranoside (IPTG): 0.1 M IPTG (Inalco) solution prepared just before use in sterile water. 5. Lysis buffer: 0.05 M sodium phosphate (pH 7.2), 0.3 M NaCl, 10% (v/v) glycerol, 1 μM pepstatin, 1 μM leupeptin, 1 μM PMSF. 6. Concentrating cell: Ultrafiltration Amicon Cell equipped with a YM 10 membrane (Amicon). 7. Gel filtration chromatography: Bio-gel P-60 (Bio-Rad) equilibrated and developed with 0.05 M Tris–HCl (pH 7.5), 0.3 M ammonium sulfate. 8. Anion exchange chromatography: Q Sepharose (GE Healthcare Biosciences) equilibrated with 0.03 M Tris–HCl (pH 7.5) and eluted with a linear gradient of NaCl (from 0 to 0.6 M) in 0.03 M Tris–HCl (pH 7.5). 9. Hydrophobic interaction chromatography: Phenyl Sepharose (GE Healthcare Biosciences) equilibrated with 0.05 M Tris–HCl buffer (pH 7.5) and 1 M ammonium sulfate and eluted with a linear gradient of ammonium sulfate (from 1 to 0 M) in 0.05 M Tris–HCl buffer (pH 7.5). 2.3. Preparation of Retinol–RBP and Fenretinide–RBP Complexes

1. Prepare just before use a concentrated ethanolic solution of all-trans retinol (Fluka, Buchs, Switzerland) and quantify it by using an extinction coefficient of 46,000 M−1 cm−1 at 325 nm (27). 2. Fenretinide (N-(4-hydroxyphenyl)retinamide) was a gift from R.W. Johnson Pharmaceutical Research Institute (Spring House, PA, USA). Prepare just before use of ethanolic solutions of 0.5–1 mM fenretinide, using an extinction coefficient of 55,630 M−1 cm−1 at 361.5 nm (28).

2.4. Fluorescence Labeling of Human TTR with the Dansyl (DNS) Probe

1. A solution of 5-dimethylaminonaphthalene-1-sulfonyl chloride (DNS-Cl) (Fluka) is prepared just before use by dissolving 0.1 mg of the compound in 1 ml of acetone. 2. Dextran-coated charcoal solution is prepared by suspending 25 mg of charcoal (Sigma) and 0.25 mg of dextran (Sigma) in 1 ml of 0.1 M sodium bicarbonate buffer (pH 8.8).

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3. Methods 3.1. Heterologous Expression, Unfolding/Refolding, and Purification of Human RBP 3.1.1. Cloning the cDNA Sequence Coding for Human RBP

1. Amplify the mature coding sequence adding a NdeI restriction site at the 5 end and a BamHI restriction site at the 3 end by using a specific NdeI-tailed upstream primer (5 -ATACATATGGAGCGCGACTGCCGAGTG-3 ) and a specific BamHI-tailed downstream primer (5 -TAAGGAT CCGATTCTTGATATTGCTACAAAAGG-3 ). 2. Clone the sequence into pGem-T-easy vector following the manufacturer’s procedure. 3. Digest for 3 h at 37◦ C the recombinant plasmid by using three units of NdeI and BamHI enzymes per microgram of DNA. Separate the restriction fragments by electrophoresis on a 0.8% (w/v) agarose gel, elute, and purify the fragment corresponding to the coding sequence from the gel by using the Gel Extraction Kit following the manufacturer’s procedure and quantify the DNA by absorbance at 260 nm. 4. Perform the ligation reaction by mixing 75 ng of the coding fragment with 250 ng of the pET11b vector previously digested with NdeI and BamHI and dephosphorylated. Precipitate the DNA and redissolve it in 3 μl of water. 5. Use 1.5 μl of DNA to transform E. coli Origami (DE3) cells (Stratagene) by electroporation.

3.1.2. Expression and Purification of Human RBP

1. Grow up overnight, at 37◦ C, a 20 ml culture of transformed E. coli Origami (DE3) cells in freshly prepared Luria broth containing 40 μl of ampicillin solution, 40 μl of tetracycline solution, and 20 μl of kanamycin solution. 2. Inoculate 100 ml of Luria broth containing 200 μl of ampicillin solution, 200 μl of tetracycline solution, and 100 μl of kanamycin solution with 2.5 ml of the aforementioned culture. Incubate for about 150 min at 37◦ C until an absorbance of 0.6–0.8 at 600 nm is attained. Add 1 ml of IPTG solution to induce the expression of RBP and continue the incubation overnight at 37◦ C. 3. Separate the cells by centrifugation at 7,500 rpm for 15 min at 4◦ C, resuspend the pellet with 20 ml of 0.05 M Tris–HCl, 1 mM EDTA (pH 8), and repeat the centrifugation.

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4. Resuspend the pellet with 6 ml of lysis buffer, sonicate the suspension (20 bursts of 30 s with pauses of 30 s) keeping the temperature below 10◦ C, and separate the soluble cell extract by centrifugation at 7,500 rpm for 15 min at 4◦ C. 5. Verify the expression of RBP and its presence in the insoluble fraction by SDS-PAGE. 6. Eliminate the supernatant and resuspend the insoluble fraction containing RBP in 20 ml of denaturing buffer. 7. Refold RBP by dialysis against 10 l of refolding buffer for 48 h at 4◦ C. 8. Recover the protein solution, centrifuge at 7,500 rpm for 15 min to remove the insoluble fraction, and adjust the pH to 7.0 with HCl. 9. Concentrate the protein solution with the ultrafiltration Amicon cell. 10. Perform the gel filtration chromatography at 4◦ C, monitoring the elution profile at 280 nm. The refolded RBP elutes with a relative retention volume (Ve /V0 ) of approximately 1.7 and can be identified by SDS-PAGE (see Note 1). 11. Collect the fractions containing RBP and concentrate the protein by means of a Centricon centrifugal filter device equipped with YM-10 membrane. 12. Perform the TTR-affinity chromatography at 4◦ C, monitoring the elution profile at 280 nm. The refolded RBP possesses affinity for TTR, but is eluted from the column when a low ionic strength solution is applied. RBP can be unambiguously identified by monitoring, upon addition of retinol, the typical absorption and fluorescence spectra of the retinol–RBP complex (see Section 3.4 and Fig. 11.2). 13. Quantify apo- and holoRPB using absorption coefficients (A 1 mg/ml, 1 cm) at 279 nm of 1.74 and 2.02, respectively (29). 3.2. Heterologous Expression and Purification of Human TTR 3.2.1. Cloning the cDNA Sequence Coding for Human TTR

For the cloning of the sequence encoding for the mature human TTR, the procedure is essentially the same described for RBP (Section 3.1.1), except for the following: 1. Amplify the coding sequence adding a NdeI restriction site at the 5 end and a BamHI restriction site at the 3 end by using a specific upstream primer (5 -ATA

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CATATGGAGCGCGACTGCCGAGTG-3 ) and a specific downstream primer (5 -TAAGGATCCGATTCTTGATATT GCTACAAAAGG-3 ). 2. Perform the ligation reaction in pET11b by mixing 54 ng of the coding fragment with 250 ng of the expression vector. 3. Transform E. coli BL21 (DE3) cells (Stratagene) by electroporation. 3.2.2. Expression and Purification of Human TTR

1. Grow up overnight, at 37◦ C, a 20 ml culture of transformed E. coli BL21 (DE3) cells, in freshly prepared Luria broth containing 40 μl of ampicillin solution. 2. Inoculate 1 l of Luria broth containing 2 ml of ampicillin solution with 15 ml of the aforementioned culture. Incubate for about 120 min at 37◦ C until an absorbance of 0.6–0.8 at 600 nm is attained. Add 10 ml of IPTG solution to induce the expression of RBP and continue the incubation for 4 h at 28◦ C. 3. Separate the cells by centrifugation at 7,500 rpm for 15 min at 4◦ C, resuspend the pellet with 20 ml of 0.05 M Tris–HCl buffer (pH 8.0), 1 mM EDTA, and repeat the centrifugation. 4. Resuspend the pellet with 60 ml of lysis buffer, sonicate the suspension (20 bursts of 30 s with pauses of 30 s) keeping the temperature below 10◦ C, and separate the soluble cell extract by centrifugation at 7,500 rpm for 15 min at 4◦ C. 5. Verify the expression of TTR and its solubility by SDSPAGE. 6. Concentrate the supernatant with the ultrafiltration Amicon cell. 7. Perform the gel filtration chromatography at 4◦ C, monitoring the elution profile at 280 nm. TTR is eluted from the column with a relative retention volume (Ve /V0 ) of approximately 1.25. 8. Collect the fractions containing the TTR identified by SDS-PAGE and concentrate the protein with the ultrafiltration Amicon cell. 9. Perform the anion exchange chromatography at 4◦ C, monitoring the elution profile at 280 nm. TTR is eluted at an NaCl concentration of about 0.5 M. 10. Collect the fractions containing the TTR identified by SDS-PAGE and concentrate the protein with the ultrafiltration Amicon cell. 11. Perform the hydrophobic interaction chromatography anion exchange chromatography at 4◦ C, monitoring the

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elution profile at 280 nm. TTR is eluted at an ammonium sulfate concentration of about 0.5 M. 12. Quantify TTR using an absorption coefficient (A 1 mg/ml, 1 cm) at 279 nm of 1.43 (12). 3.3. Absorption and Fluorescence Spectra of the Retinol–RBP Complex (holoRBP)

To verify the specificity of the interaction between retinol and recombinant human RBP, the absorption and fluorescence emission spectra of the RBP-bound retinol are analyzed. 1. Add 0.5 μl of an ethanolic solution of 1 mM retinol to the spectrophotometer cuvette containing 6 μM apoRBP in 100 μl of 0.05 M sodium phosphate buffer (pH 7.3), 0.15 M NaCl (see Note 3). A slight molar excess of RBP relative to retinol (the RBP/retinol ratio in the system is approximately 1.1) ensures that nearly all the ligand is bound to the protein (a correction for the contribution of free ligand to the spectra of the retinol–RBP complex is, therefore, not needed). Stir gently and let retinol bind to RBP in the dark at 20◦ C for 15–20 min. Use this solution to record subsequently both absorption and fluorescence spectra. 2. Record the absorption spectrum of holoRBP in the 250–380 nm range (see Fig. 11. 2a and Note 4). 3. Record the fluorescence emission spectrum in the 400–550 range (see Fig. 11.2b and Note 5) of holoRBP using an excitation wavelength of 330 nm and a reduced excitation slit width (<3 nm) (see Note 6).

3.4. The Interaction of Recombinant Human holoRBP with TTR Analyzed by Fluorescence Anisotropy

To investigate the binding of TTR to holoRBP, titrations are carried out by monitoring the change in fluorescence anisotropy of the RBP-bound retinol upon addition of TTR (see Fig. 11.3a and Note 7). Steady-state fluorescence anisotropy values are measured at 20 ± 0.5◦ C by means of a Perkin-Elmer LS-50B spectrofluorometer, which uses the L-format geometry and is equipped with the accessory for fluorescence anisotropy measurements. 1. Add to the spectrofluorometer cuvette containing 3 μM recombinant holoRBP in 100 μl of 0.05 M sodium phosphate buffer (pH 7.3), 0.15 M NaCl, increasing aliquots of a concentrated solution of TTR. Stir the solution gently after each addition and measure the fluorescence anisotropy values using excitation and emission wavelengths of 330 and 460 nm, respectively, and a reduced excitation slit width (<3 nm) (see Note 6). Fluorescence anisotropy is determined according to Eq. [13] reported in Appendix. 2. Plot the fluorescence anisotropy values as a function of TTR concentration in the cuvette.

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3.5. Fluorescence Labeling of Human TTR with the Dansyl (DNS) Probe

The affinity labeling of human TTR with DNS-Cl (see Note 2) is accomplished as follows: 1. Incubate 2 mg of human TTR, purified from human plasma (14), with 2.3-fold molar excess of DNS-Cl solution in 0.1 M sodium bicarbonate buffer (pH 8.8) (total volume: 2.5 ml) for 90 min at 4◦ C, stirring continuously the solution. 2. Add 0.25 ml of dextran-coated charcoal solution, stir for 30 min at 4◦ C, and centrifuge for 30 min at 4◦ C to remove the excess of DNS. 3. Dialyse the supernatant for 48 h at 4◦ C against 0.05 M sodium phosphate buffer (pH 7.3), 0.15 M NaCl. 4. Concentrate the protein by means of a Centricon centrifugal filter device.

3.6. The Interactions of apoRBP and of the Fenretinide–RBP Complex with DNS-Labeled TTR (DNS-TTR) Analyzed by Fluorescence Anisotropy

To investigate the binding of apoRBP and of the fenretinide–RBP complex to DNS-TTR, titrations are carried out by following the change in the fluorescence anisotropy of the TTR-bound DNS (see Fig. 11.3b and Note 8). 1. Obtain apoRBP from holoRBP, purified from human plasma (14), by extraction of the RBP-bound retinol with diethyl ether as described (21). 2. Prepare the fenretinide–RBP complex by incubating apoRBP (5 × 10−5 M) with a slight molar excess of the retinoid in 0.05 M sodium phosphate buffer (pH 7.3), 0.15 M NaCl, for 2 h at 4◦ C. 3. Add to the spectrofluorometer cuvette containing 4 μM DNS-TTR in 100 μl 0.05 M sodium phosphate buffer (pH 7.3), 0.15 M NaCl, increasing aliquots of concentrated solutions of apoRBP or of the fenretinide–RBP complex. Stir the solution gently after each addition and measure the fluorescence anisotropy values using excitation and emission wavelengths of 380 and 480 nm, respectively. 4. Plot the fluorescence anisotropy values as a function of RBP concentration in the cuvette.

3.7. Analysis of Fluorescence Anisotropy Titration Data

Assuming that monomeric holoRBP binds to n equivalent and independent sites on tetrameric TTR, the corresponding equilibrium dissociation constant Kd can be derived from the mass law equation: Kd = [holoRBP]n[TTR]/[holoRBP − TTR]

[1]

where [holoRBP] is the concentration of free holoRBP and n[TTR] and [holoRBP–TTR] are the concentrations of binding sites on TTR, respectively free and complexed with holoRBP.

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Equation [1] is used to derive a working equation for the evaluation of the binding parameters n and Kd : α 2 [holoRBP]0 − α(n[TTR]0 + [holoRBP]0 + Kd ) + n[TTR]0 = 0 [2]

where α(α = [holoRBP − TTR]/[holoRBP]0 ) is the fraction of holoRBP bound by TTR and [holoRBP]0 and [TTR]0 are total molar concentrations of holoRBP and TTR, respectively. By taking the negative square root for the solution of the quadratic equation, the fraction of protein bound can be expressed as α = {n[TTR]0 + [holoRBP]0 + Kd − [(n[TTR]0 + [holoRBP]0 +Kd )2 − 4[holoRBP]0 n[TTR]0 ]0.5 }/2[holoRBP]0 [3] The value of α is calculated for every point of the titration curve using the equation α = (r − r0 )/(rmax − r0 )

[4]

where r represents the fluorescence anisotropy value of RBPbound retinol for a certain molar concentration of TTR and rmax and r0 are the two limiting anisotropy values, i.e., in the presence of an excess saturating TTR and in the absence of TTR, respectively. For the analysis of binding data in the case of the titration of DNS-TTR with apoRBP or RBP in complex with non-fluorescent retinoids, Eqs. [2] and [3] can be rewritten to assume the form shown in Eqs. [5] and [6], respectively: α  2 n[DNS − TTR]0 − α  ([RBP]0 + n[DNS − TTR]0 + Kd ) + [RBP]0 = 0

[5] α  = {[RBP]0 + n[DNS − TTR]0 + Kd − [([RBP]0 + n[DNS − TTR]0 +Kd )2 − 4n[DNS − TTR]0 [RBP]0 ]0.5 }/2n[DNS − TTR]0

[6]

where α  (α  = [RBP − TTR]/n[DNS − TTR]0 ) is the fraction of the sites of DNS-TTR bound by RBP, n is the number of RBPbinding sites present on the DNS-TTR molecule, [RBP]0 is the total molar concentration of RBP, and n[DNS-TTR]0 is the total molar concentration of binding sites on DNS-TTR. The value of α  is calculated for every point of the titration curves using the equation α  = (r − r0 )/(rmax − r0 )

[7]

where r represents the fluorescence anisotropy value of DNSTTR for a certain molar concentration of RBP and rmax and r0

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are the two limiting anisotropy values of DNS-TTR, i.e., in the presence of an excess saturating RBP and in the absence of RBP, respectively. The experimentally determined values of α and α  are fitted to Eqs. [3] and [6], respectively, by means of non-linear least squares regression (see Fig. 11.3a and b), allowing us to determine both the dissociation constant and the binding stoichiometry for the interactions of RBP with TTR. The non-linear regression analyses of binding data have been performed by means of Sigma Plot (Jandel, Corte Madera, CA, USA).

4. Notes 1. Two distinct peaks containing RBP, on the basis of SDSPAGE, were eluted from the gel filtration column. The first peak corresponds to the V0 of the column and contains proteins of large dimensions, while the second, which corresponds to the expected molecular mass of RBP, is collected and further processed. 2. DNS is covalently and specifically bound to TTR upon reaction of DNS-Cl with TTR Lys-15 (30). 3. It is very important to use freshly prepared alcoholic solutions of retinol, protected from light and kept at 0◦ C to avoid its degradation. In fact, retinol is particularly unstable in the aqueous medium but a certain instability is present also in media where retinoids are generally dissolved, such as ethanol and methanol. 4. The absorption spectrum of holoRBP is characterized by a single, well-shaped peak centered at approximately 328 nm. 5. Retinol exhibits a drastically higher fluorescence emission when bound to RBP relative to uncomplexed retinol. 6. When fluorescence spectra or fluorescence intensities are recorded, the excitation slit width should be rather small to minimize the possible decomposition of retinol induced by the excitation light. Likewise, the time of exposure of the spectrofluorometer cuvette to the exciting light should be as short as possible (5–10 s) for each measurement. 7. The fluorescent probe is retinol itself bound to RBP. Since the RBP monomer has a molecular mass of 21 kDa and up to 2 RBP molecules can be bound by the TTR tetramer whose molecular mass is 55 kDa, the dimensions of the RBP–TTR 2:1 complex increase up to 97 kDa as compared to the RBP monomer, i.e., by 361%, consistent with the large increase in

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fluorescence anisotropy observed upon RBP–TTR complex formation. When retinol binds into the hydrophobic internal cavity of RBP, its fluorescence lifetime increases considerably up to 12–13 ns, but does not change appreciably when RBP–retinol is complexed by TTR (24). Since retinol is rigidly bound to RBP, being lodged inside a hydrophobic internal cavity of the carrier protein, the observed change in the fluorescence anisotropy of the RBP-bound retinol upon formation of the RBP–TTR complex can be well attributed to the change in molecular dimensions associated with the formation of the protein–protein complex. Assuming τ =12–13 ns for both holoRBP and the holoRBP–TTR complex, the observed change of r (r ≈ 0.11, Fig. 11.3a) agrees fairly well with the molecular dimensions of uncomplexed holoRBP (21 kDa) and of the holoRBP–TTR complex (2:1 stoichiometry, 97 kDa), according to Appendix Eqs. [14] and [15]. 8. The fluorescent probe is DNS bound to TTR. Since TTR is a tetramer of 55 kDa, able to bind up to two molecules of RBP, the dimensions of the RBP–TTR 2:1 complex increase up to 97 kDa, i.e., by 76%, consistent with a smaller, but significant, increase in fluorescence anisotropy relative to that observed when the fluorescence of the RBP-bound retinol is exploited for fluorescence anisotropy measurements. Assuming a value of τ for the DNS fluorofore covalently bound to TTR similar to that of the RBP-bound retinol (31), the observed fluorescence anisotropy change (r ≈ 0.03, Fig. 11.3b) due to the increment in molecular size from TTR up to the RBP–TTR complex (2:1 stoichiometry) is in very good agreement with that expected theoretically according to Appendix Eqs. [14] and [15].

5. Appendix: Fluorescence Anisotropy Theory

Fluorescence anisotropy is a measure of polarized fluorescence emission from samples excited with polarized light and derives from the fact that both absorption and emission transition moments of a molecule are oriented in a given direction. If these dipoles are not parallel, the emitted light will be depolarized to some extent, depending on the angle they form. Because molecules in solution are randomly oriented, only a fraction of them, with the absorption dipole oriented in the same direction of the polarized incident light, will be excited. During their excited state lifetime, these molecules will also undergo

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rotational diffusion, i.e., will reorient to some extent causing further depolarization. Rotational diffusion depends on a number of parameters, among which viscosity, size, and shape of the molecules are the most important. Usually, the fluorescence anisotropy of small molecules is very small, because they rotate very fast, but for macromolecules, such as proteins, the rotational correlation time (θ ), the inverse of the rotational diffusion rate, becomes comparable to the excited state lifetime (τ ), and the corresponding fluorescence anisotropy (r) may be large enough to be measured. For a globular protein of 20 kDa in water at room temperature θ is close to 10 ns, a value similar to the lifetime of many fluorescent probes conjugated to proteins. The dependence of r on the rotational diffusion rate is the reason why anisotropy has often been exploited in biochemical studies, where changes of size and/or shape of macromolecules can take place, such as upon formation of complexes between small fluorescent ligands and proteins and protein–protein or protein–DNA associations. The fluorescence anisotropy r measured under continuous illumination is defined as steady-state anisotropy, i.e., the time average value of the anisotropy decay curve. The time-resolved anisotropy can be measured, instead, only under ns- or pspulsed excitation. Since in this context we deal with steady-state anisotropy only, it will be merely mentioned here as anisotropy. Fluorescence anisotropy measured under constant illumination is given by r=

F − F⊥ F + 2F⊥

[8]

where F and F⊥ are the fluorescence intensities measured using two polarizers, positioned along the excitation and emission light pathway, oriented parallel and perpendicular with respect to the direction of the plane polarized excitation beam, respectively. The difference of these two terms is normalized over the total fluorescence intensity, given by the denominator (32). In earlier literature, fluorescence polarization P was used in place of anisotropy (33): P=

F − F⊥ F + F⊥

[9]

Though the two parameters are clearly related, r is preferred, because its formulation, for a sample containing a mixture of fluorophores, is simpler: r¯ =

 i

fi ri

[10]

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where r¯ is the average anisotropy and fi ri are the fractional anisotropies of each component. According to Eqs. [8] and [9], if the emitted light is completely polarized, r = P = 1, and if it is completely depolarized, r = P = 0. However, for homogeneous samples in solution, the limiting anisotropy value is much lower and equal to 0.4, because molecules are randomly oriented and only a fraction of them can be excited by the incident polarized light (34). Furthermore, this maximum value can only be reached if the excitation and emission dipole moments are collinear and rigid, which is rarely the case. In practice, Eqs. [8] and [9] need to be corrected for the different transmission efficiency of the emission monochromator for vertically and horizontally polarized light. The correction can be achieved using horizontally polarized excitation light. It can be shown (34) that FHH FVV F = F⊥ FHV FVH

[11]

where H and V refer to the horizontal and vertical orientation of excitation and emission polarizers, respectively. The necessary correction factor is thus given by G=

FHV FHH

[12]

and the true anisotropy can then be rewritten as r=

FVV − GFVH FVV + 2GFVH

[13]

Apart from trivial causes, such as light scattering, reabsorption, and misalignment of polarizers, the main cause of depolarization is given by the rotational diffusion of the particles in solution, as already mentioned. For spherical particles, fluorescence anisotropy is given by the Perrin equation (35): r=

r0 r0 = 1 + 6Dτ 1 + τ/6

[14]

where r0 is the fundamental anisotropy (i.e., the anisotropy in the absence of Brownian rotation), τ is the fluorescence lifetime of the fluorescent reporter group, and D is diffusion coefficient of the protein, related to the rotational correlation time by D = 1/(6θ). θ is, in turn, related to V, the hydrated volume, and Mh , the hydrated molecular mass, of the particle by θ=

ηV = ηMh /RT RT

[15]

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where η is the viscosity of the solution, R is the gas constant, and T is the absolute temperature. For non-spherical particles, the values of θ are larger than those predicted by Eq. [15], not only for a change of shape but also for larger hydration shells. Because of these two factors, the experimental values of θ are usually found to be almost twice as large as those expected for anhydrous globular proteins. For globular proteins in solution, θ has been empirically found to increase linearly with the molecular weight by ≈7 ns every 10 kDa, at least in the 20–70 kDa range (36). References 1. Kawaguchi, R., Yu, J., Honda, J., Hu, J., Whitelegge, J., Ping, P., Wiita, P., Bok, D., Sun, H. (2007) A membrane receptor for retinol binding protein mediates cellular uptake of vitamin A. Science 315, 820–825. 2. Zanotti, G., Berni, R. (2004) Plasma retinolbinding protein: Structure and interactions with retinol, retinoids, and transthyretin. Vitam. Horm. 69, 271–295. 3. Goodman, D.S. (1984) Plasma retinolbinding protein. In: Sporn, M.B., Roberts, A.B., Goodman, D.S. (eds.), The Retinoids, Academic Press, New York, pp. 41–88s. 4. Newcomer, M.E., Jones, T.A., Aqvist, J., Sundelin, J., Eriksson, U., Rask, L., Peterson, P.A. (1984) The three-dimensional structure of retinol-binding protein. EMBO J. 3, 1451–1454. 5. Cowan, S.W., Newcomer, M.E., Jones, T.A. (1990) Crystallographic refinement of human serum retinol binding protein at 2A resolution. Proteins 8, 44–61. 6. Benson, M.D., Kincaid, J.C. (2007) The molecular biology and clinical features of amyloid neuropathy. Muscle Nerve 36, 411–423. 7. Blake, C.C., Geisow, M.J., Oatley, S.J., Rerat, B., Rerat, C. (1978) Structure of prealbumin: Secondary, tertiary and quaternary interactions determined by Fourier refinement at 1.8 A. J. Mol. Biol. 121, 339–356. 8. Hornberg, A., Eneqvist, T., Olofsson, A., Lundgren, E., Sauer-Eriksson, A.E. (2000) A comparative analysis of 23 structures of the amyloidogenic protein transthyretin. J. Mol. Biol. 302, 649–669. 9. Monaco, H.L., Rizzi, M., Coda, A. (1995) Structure of a complex of two plasma proteins: Transthyretin and retinol-binding protein. Science 268, 1039–1041. 10. Naylor, H.M., Newcomer, M.E. (1999) The structure of human retinol-binding protein (RBP) with its carrier protein transthyretin reveals an interaction with the

11.

12.

13.

14.

15.

16.

17.

18.

carboxy terminus of RBP. Biochemistry 38, 2647–2653. Zanotti, G., Folli, C., Cendron, L., Alfieri, B., Nishida, S.K., Gliubich, F., Pasquato, N., Negro, A., Berni, R. (2008) Structural and mutational analyses of proteinprotein interactions between transthyretin and retinol-binding protein. FEBS J. 275, 5841–5854. Tragardh, L., Anundi, H., Rask, L., Sege, K., Peterson, P.A. (1980) On the stoichiometry of the interaction between prealbumin and retinol-binding protein. J. Biol. Chem. 255, 9243–9248. Berni, R., Malpeli, G., Folli, C., Murrell, J.R., Liepnieks, J.J., Benson, M.D. (1994). The Ile-84–>Ser amino acid substitution in transthyretin interferes with the interaction with plasma retinol-binding protein. J. Biol. Chem. 269, 23395–23398. Malpeli, G., Folli, C., Berni, R. (1996). Retinoid binding to retinol-binding protein and the interference with the interaction with transthyretin. Biochim. Biophys. Acta 1294, 48–54. Goodman, D.S., Raz, A. (1972) Extraction and recombination studies of the interaction of retinol with human plasma retinol-binding protein. J. Lipid Res. 13, 338–347. Folli, C., Viglione, S., Busconi, M., Berni, R. (2005) Biochemical basis for retinol deficiency induced by the I41N and G75D mutations in human plasma retinol-binding protein. Biochem. Biophys. Res. Commun. 336, 1017–1022. van Bennekum, A.M., Wei, S., Gamble, M.V., Vogel, S., Piantedosi, R., Gottesman, M., Episkopou, V., Blaner, W.S. (2001) Biochemical basis for depressed serum retinol levels in transthyretin-deficient mice. J. Biol. Chem. 276, 1107–1113. Zanotti, G., Malpeli, G., Berni, R. (1993) The interaction of N-ethyl retinamide with plasma retinol-binding protein (RBP) and

Interaction Between Retinol-Binding Protein and Transthyretin

19.

20.

21.

22.

23.

24.

25.

26.

the crystal structure of the retinoid-RBP complex at 1.9-A resolution. J. Biol. Chem. 268, 24873–24879. Zanotti, G., Marcello, M., Malpeli, G., Folli, C., Sartori, G., Berni, R. (1994). Crystallographic studies on complexes between retinoids and plasma retinol-binding protein. J. Biol. Chem. 269, 29613–29620. Coward, P., Conn, M., Tang, J., Xiong, F., Menjares, A., Reagan, J.D. (2009) Application of an allosteric model to describe the interactions among retinol binding protein 4, transthyretin, and small molecule retinol binding protein 4 ligands. Anal. Biochem. 384, 312–320. Zanotti, G., Berni, R., Monaco, H.L. (1993) Crystal-structure of liganded and unliganded forms of bovine plasma retinol-binding protein. J. Biol. Chem. 268, 10728–10738. Blomhoff, R., Green, M.H., Norum, K.R. (1992) Vitamin A: Physiological and biochemical processing. Ann. Rev. Nutr. 12, 7–57. Folli, C., Pasquato, N., Ramazzina, I., Battistutta, R., Zanotti, G., Berni, R. (2003). Distinctive binding and structural properties of piscine transthyretin. FEBS Lett. 555, 279–284. Kopelman, M., Cogan, U., Mokady, S., Shinitzky, M. (1976) The interaction between retinol-binding proteins and prealbumins studied by fluorescence polarization. Biochim. Biophys. Acta 439, 449–460. Pasquato, N., Berni, R., Folli, C., Alfieri, B., Cendron, L., Zanotti, G. (2007) Acidic pH-induced conformational changes in amyloidogenic mutant transthyretin. J. Mol. Biol. 366, 711–719. Berni R., Stoppini M., Zapponi M.C. (1992) The piscine plasma retinol-binding protein. Purification, partial amino acid sequence and interaction with mammalian transthyretin

27.

28.

29.

30.

31. 32. 33.

34. 35.

36.

207

of rainbow trout (Oncorhynchus mykiss) retinol-binding protein. Eur. J. Biochem. 204, 99–106. Horwitz, J., Heller, J. (1973) Interactions of all-trans, 9-, 11-, and 13-cis-retinal, alltrans-retinyl acetate, and retinoic acid with human retinol-binding protein and prealbumin. J. Biol. Chem. 248, 6317–6324. Berni, R., Clerici, M., Malpeli, G., Cleris, L., Formelli, F. (1993) Retinoids: In vitro interaction with retinol-binding protein and influence on plasma retinol. FASEB J. 7, 1179–1184. Bychkova, V.E., Berni, R., Rossi, G.L., Kutyshenko, V.P., Ptisyn, O.B. (1992) Retinol-binding protein is in the molten globule state at low pH. Biochemistry 31, 7566–7571. Cheng, S.Y., Cahnmann, H.J., Wilchek, M., Ferguson, R.N. (1975) Affinity labelling of the thyroxine binding domain of human serum prealbumin with dansyl chloride. Biochemistry 14, 4132–4136 Hemmila, I.A. (1991) Applications of Fluorescence Immunoassays, John Wiley & Sons, New York. ´ Jabłonski, A. (1960) On the notion of emission anisotropy. Bull. Acad. Pol. Sci. Ser. A 8, 259–264. Weber, G. (1952) Polarization of the fluorescence of macromolecules. I. Theory and experimental method. Biochem. J. 51, 145–155. Lakowitz, J.R. (1999) Principles of Fluorescence Spectroscopy, Kluver Academy, Plenum Publishers, New York. Perrin, A. (1929) La fluorescence des solutions. Induction moléculaire. Polarization et durée d’émission. Photochimie. Ann. Phys. Ser. 10 12, 169–275. Yguerabide, J., Epstein, H. F., Stryer, L. (1970) Segmental flexibility in an antibody molecule. J. Mol. Biol. 51, 573–590.

Chapter 12 Assay of Retinol-Binding Protein–Transthyretin Interaction and Techniques to Identify Competing Ligands Nathan L. Mata, Kim Phan, and Yun Han Abstract The principles of fluorescence resonance energy transfer have been utilized to develop a high-throughput assay which detects compounds that interfere with interaction between retinol-binding protein (RBP) and transthyretin (TTR). In this assay, the intrinsic fluorescence from the RBP–retinol complex excites a probe molecule which is covalently coupled to TTR. Generation of an emission signal from the TTR probe indicates interaction between RBP–retinol and TTR. Importantly, the inclusion of retinol in the assay allows discrimination of test compounds which bind RBP versus those which bind to TTR. Thus, compounds which bind to RBP must compete with retinol in order to affect RBP–TTR interaction. This feature of the assay will be useful to identify test compounds which are more likely to have an effect in vivo. Key words: Fluorescence resonance energy transfer (FRET), inner filter effect, Cogan plot, retinol, retinol-binding protein (RBP), transthyretin (TTR), N-(4-hydroxyphenyl)retinamide (HPR).

1. Introduction Numerous scientific reports have implicated serum retinolbinding protein (RBP) as an important modulator in the pathogenesis of degenerative diseases such as diabetes (1–7) and macular dystrophies which are characterized by accelerated accumulation of lipofuscin fluorophores (e.g., Stargardt disease and dry age-related macular degeneration) (8–12). Collectively, these findings support the notion that pharmacological modulation (reduction) of RBP could be a potential therapeutic strategy for the treatment of these diseases. RBP is secreted from the liver as a holo protein and relies upon interaction with transthyretin (TTR) H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_12, © Springer Science+Business Media, LLC 2010

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to attain a high steady-state level in the circulation (13, 14). In the absence of TTR binding, RBP–retinol is eliminated through glomerular filtration due to its small size (∼21 kDa). Therefore, a compound which competes with retinol for binding to RBP and also prevents interaction with TTR would be a good candidate for RBP modulation. In order to identify candidate compounds for RBP modulation, a high-throughput, fluorescence-based assay which monitors RBP–TTR interaction has been developed. This assay relies upon intrinsic fluorescence from the RBP–retinol complex and fluorescence emanating from a probe which is covalently bound to TTR to monitor interaction between RBP and TTR. Although this assay was developed primarily as a screening tool to identify candidate therapeutic compounds, the assay principles and techniques can be easily applied to other retinoid-binding protein– ligand systems. Important considerations for the development of this assay are discussed below. A well-established method to study interaction between retinoids and retinoid-binding proteins is fluorescence titration (15). In this technique, the intrinsic fluorescence emission from the protein source is monitored in the presence of increasing concentrations of a presumptive ligand. Aromatic amino acids in a protein will produce a fluorescence emission at ∼340 nm following excitation at ∼280 nm. If a presumptive ligand binds to the protein, and the ligand absorbs light energy within the wavelength range of the protein fluorescence emission, the emitted energy will be transferred to the ligand. The result will be a reduction or “quenching” of the fluorescence from the protein. This process is referred to as Förster resonance energy transfer (FRET), named after the German photochemist Theodor Förster (16). The term fluorescence resonance energy transfer is also used to describe this same phenomenon. Importantly, FRET only occurs when the donor emission and acceptor excitation spectra overlap and the compounds are in close proximity to one another (50–80 Å) (17). A representative example of fluorescence quenching is shown in Fig. 12.1. In this experiment, fluorescence emission intensity from RBP was monitored in the presence of increasing concentrations of retinol (excitation = 280 nm, emission = 335 nm). Reduction in RBP fluorescence as a function of increasing retinol concentration indicates that retinol binds to RBP (Fig. 12.1a). The change in fluorescence intensity can then be used to calculate the number of ligand-binding sites on the protein (n) and the apparent affinity (KD ), which governs the protein–ligand interaction. It is also possible to observe fluorescence quenching in the absence of true protein–ligand binding. This phenomenon occurs when the absorbance of the solution is very high (e.g., at excessive ligand concentrations). Under this condition, the fluorescence

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Fig. 12.1. Fluorescence titration of RBP with retinol: RBP emission. The kinetics of RBP–retinol binding were measured using 0.5 μM RBP and increasing concentrations of retinol (up to 4 μM). Excitation wavelength was 280 nm and emission was monitored at 335 nm. The reduction in 335 nm emission indicates binding of retinol to RBP (a). Application of the mass law equation to the fluorescence data and generation of a Cogan plot yield a slope and y-intercept from which the binding constants were determined to be n = 0.911 and KD = 83 nM (b).

emission of the protein is blocked, or “filtered,” by the ligand in a concentration-dependent manner. This effect, referred to as “inner filter effect” (15), must be identified and compensated for using an appropriate correction factor (see Note 1). Upon establishing that a particular protein and ligand do interact, the fluorescence emission intensity data can be used to determine n and KD . The following mathematical formula, derived from the mass law equation by Cogan et al. (18), is a widely accepted method to calculate n and KD from fluorometric titration experiments: P0 α =

R0 α KD 1 × − , n 1−α n

wherein P0 = total protein concentration R0 = ligand concentration α = (Fmax − F )/(Fmax − Fo ); Fmax is the fluorescence at saturation; F is the fluorescence at each ligand concentration; F0 is the initial fluorescence n = number of independent binding sites The first three parameters (P0 , R0 , and α) can be readily obtained from the titration experiment. However, in some cases the n value may not be known. In these cases, one can first generate a plot of P0 α against R0 α/(1–α). A regression line can then be fitted through the data points to obtain the slope and y-intercept. Values for n and KD can then be determined from the plot. A practical application of the mass law equation is shown in Fig. 12.1b. The plot shown, referred to as a Cogan plot, was

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obtained using the fluorescence emission intensity data shown in Fig. 12.1a. A linear regression analysis of this data yields a slope of 1.097 and a y-intercept of 0.0915. The binding site number (n) is given by the reciprocal of the slope and KD is given by the product of the y-intercept and n. In this example, n = 0.911 and KD = 83 nM. RBP is known to have a single binding site (n = 1) with a KD of 70–190 nM (19, 20). Thus, the techniques described here produce binding constants which are consistent with the published literature. In the example described above, the decreasing fluorescence emission from RBP (at ∼335 nm) was used to determine the binding constants of RBP–retinol interaction. Importantly, the FRET event between RBP and retinol produces a second fluorescence emission (at ∼470 nm) which emanates simultaneously from bound retinol and increases as a function of increasing retinol concentration. A representative emission spectrum taken during the RBP–retinol binding experiment shows the two fluorescence peaks (Fig. 12.2). By monitoring the fluorescence emission from retinol, we observe increased fluorescence intensity as energy from RBP is transferred to bound retinol (Fig. 12.3a). As expected, a Cogan plot of the 470 nm emission data reveals binding constants which are comparable to those obtained from the 335 nm emission data shown in Fig. 12.1 (Fig. 12.3b, n = 1.01 and KD = 49 nM).

Fig. 12.2. Emission spectrum of RBP–retinol reaction mixture. A representative emission spectrum of the RBP–retinol reaction mixture was acquired to identify the two fluorescence peaks representing emission from RBP (at ∼335 nm) and bound retinol (at ∼470 nm). Excitation was at 280 nm.

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Fig. 12.3. Fluorescence titration of RBP with retinol: retinol emission. The kinetics of RBP–retinol binding was measured using 0.5 μM RBP and increasing concentrations of retinol (up to 4 μM). Excitation wavelength was 280 nm and emission was monitored at 470 nm. The increase in 470 nm emission indicates binding of retinol to RBP (a). Application of the mass law equation to the fluorescence data and generation of a Cogan plot yield a slope and y-intercept from which the binding constants were determined to be n = 1.01 and KD = 49 nM (b).

To detect binding of TTR to the RBP–retinol complex, an appropriate fluorescent probe is required to label TTR. Ideally, this probe should absorb light energy in the region of retinol emission and emit fluorescence outside of this region. The Alexa Fluor series of fluorescent dyes, developed by InvitrogenTM , is perhaps the most comprehensive collection to choose from as it includes dyes that cover the entire visible spectrum. From this collection, Alexa Fluor 430 (AF-430) was selected as the most appropriate probe to label TTR. As shown in Fig. 12.4a, the absorbance spectrum of AF-430 overlaps with the emission spectrum of RBP–retinol. Additionally, the emission spectrum of AF430 is sufficiently broad and red-shifted relative to the RBP– retinol emission. Although the maximum emission wavelength of AF-430 is ∼550 nm, 580 nm was chosen as the optimal monitoring wavelength in order to minimize any contribution from the RBP–retinol emission. A schematic depiction of RBP– retinol interaction with AF-430-labeled TTR (designated TTR∗ ) is shown in Fig. 12.4b. Briefly, in the presence of retinol, 330 nm excitation produces an emission at ∼ 470 nm which is transferred to TTR∗ ; bound TTR∗ then produces a fluorescence emission which is monitored at 580 nm. Compounds which interfere with RBP–TTR∗ interaction should prevent FRET to TTR∗ resulting in no 580 nm emission. In order to confirm that the developed assay can detect modulators of RBP–TTR interaction, a known RBP antagonist was utilized. N-(4-hydroxyphenyl)retinamide (HPR) has been shown to compete with retinol for binding to RBP and cause dosedependent reductions in serum RBP–retinol by preventing interaction with TTR (21, 22). The ability of HPR to act as an RBP antagonist is due largely to its chemical similarity with retinol in

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Fig. 12.4. Spectral profiles of RBP–retinol and AF-430 and utilization in the RBP–TTR∗ FRET assay. In order to select an appropriate probe to label TTR for the RBP–TTR∗ FRET assay, an analysis of the RBP–retinol absorbance and emission spectra was performed. Optimal excitation of the RBP–retinol complex (at 330 nm) leads to a donor emission signal at 470 nm. AF-430 possesses an absorbance spectrum which overlaps with the emission of RBP–retinol and an emission spectrum which is sufficiently broad to detect without interference from the RBP–retinol emission (i.e., at the optimal monitoring wavelength, 580 nm) (a). Utilization of TTR∗ in the RBP–TTR∗ FRET assay is shown schematically to illustrate the binding event which leads to excitation of AF-430 and the emission signal at 580 nm indicating RBP–TTR interaction (b).

the region which interacts with the RBP binding pocket (i.e., β-ionone ring and saturated double-bond system, see Fig. 12.5). Despite the chemical similarity with retinol, HPR does not fluoresce. Importantly, the bulky, phenyl hydroxyl moiety on the terminal end of HPR provides a steric hindrance which prevents interaction with TTR. Thus, 330 nm excitation of the RBP–HPR complex will not produce a fluorescence emission. It is important to note that excitation of retinol at 330 nm will also not produce an emission as free retinol in aqueous solution does not fluoresce. A schematic depiction of this binding scenario is provided in Fig. 12.6a. An added benefit of this assay design is the ability to discriminate between compounds which interact with RBP versus those which interact with TTR∗ . Thus, increased emission at 470 nm with no increase at 580 nm indicates that the test

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Fig. 12.5. Structural comparison of retinol and HPR. The chemical structures of retinol and HPR are shown to illustrate the structural similarity of HPR to retinol. The RBP binding pocket accommodates the β-ionone ring and saturated double-bond system of retinol. HPR possesses these features allowing it to compete with retinol for binding to RBP. The phenyl hydroxyl moiety on the terminal end of HPR prevents TTR interaction.

Fig. 12.6. Schematic drawing of the RBP–TTR∗ FRET assay. The utility of the RBP–TTR∗ FRET assay to monitor RBP–TTR interaction is shown with HPR as an example antagonist. Binding of HPR to RBP prevents TTR interaction and fails to generate a fluorescence emission following 330 nm excitation. Free retinol in the reaction mixture will also fail to produce an emission signal (a). A benefit of the RBP–TTR∗ FRET assay is the ability to differentiate compounds which bind to TTR. A condition in which emission is detected at 470 nm, but not at 580 nm, indicates a presumptive TTR inhibitor (b).

compound is not interfering with RBP–retinol interaction, but is likely occupying a binding site on TTR∗ (Fig. 12.6b). In order to test these theoretical predictions, TTR∗ was prepared and the assay was conducted as described in Section 3.

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Briefly, apo-RBP and TTR∗ (0.5 μM each) were pre-incubated (30 min at 37◦ C) either in the absence or in the presence of HPR (2 μM). Following pre-incubation, retinol (1 μM) was added and incubation is resumed for another 30 min. Samples were then excited at 330 nm and emission spectra were acquired in the range of 400–600 nm. Sample spectra taken at the conclusion of the assay are shown in Fig. 12.7. The sample containing HPR shows a significant reduction of fluorescence emission from both RBP–retinol and TTR∗ indicating reduced interaction between RBP and TTR∗ . The residual fluorescence shown in the dashed trace indicates the population of RBP–retinol– TTR∗ which escaped inhibition by HPR. Importantly, the inherent dynamic range of this assay system permits a rapid determination of dose response for comparison of candidate antagonist compounds. Analysis of the dose–response curve for inhibition of RBP–TTR∗ interaction by HPR revealed half-maximal inhibition at ∼0.5 μM (Fig. 12.8). As described above, the ability of HPR to compete with retinol for binding to RBP is due to its structural similarity with retinol. Indeed, other metabolites and geometric isomers of retinol have also been shown to bind RBP (23). Investigators using circular dichroism to monitor the bound state of RBP have determined that all-trans isomers have a greater binding affinity to RBP compared to 13-cis isomers (23). This difference in binding affinity is likely due to the altered geometry of

Fig. 12.7. Effect of HPR on the RBP–TTR∗ emission spectrum. The RBP–TTR∗ FRET assay was performed in the absence and presence of HPR (2 μM). A representative emission spectrum from each reaction mixture was acquired at the conclusion of the assay. The reaction mixture containing HPR shows a significant reduction in emission intensity from both RBP–retinol and TTR∗ (compare solid and dashed traces). This result confirms a reduced interaction between RBP and TTR∗ .

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Fig. 12.8. Dose–response curve for inhibition of RBP–TTR∗ interaction by HPR. To demonstrate the dynamic range of the RBP–TTR∗ FRET assay system, inhibition of RBP–TTR interaction was measured at various HPR concentrations (0.01–25 μM). Analysis of the dose–response curve shows half-maximal inhibition at ∼0.5 μM.

13-cis isomers which are not as easily accommodated within the RBP binding pocket. Therefore, to further validate the developed RBP–TTR∗ assay, we examined binding properties of a variety of retinoids which share a structural similarity with retinol. The chemical structures of these retinoids are provided in Fig. 12.9. Assays were conducted as described above using two concentrations of each retinoid (retinol concentration was 1 μM). Data obtained from this study were consistent with the published literature. The order of inhibition potency was HPR > all-trans retinal = all-trans retinoic acid > 13-cis retinoic acid (Fig. 12.10). An alternate method to detect protein–ligand or protein– protein interaction is fluorescence anisotropy. This method, which measures binding interaction between two molecules based upon rotational diffusion, or “tumbling time”, has been used successfully in analytical studies to measure interaction between RBP– retinol and TTR (21, 24, 25). In this approach, the fluorescence emission of RBP–retinol is monitored at 0◦ and 90◦ angles in the presence and absence of TTR. Although this technique is quite sensitive, it cannot discriminate between the degree of bound versus unbound retinol because unbound retinol does not fluoresce. Thus, the output value would be the same for all levels of RBP–retinol–TTR binding. Complete binding of RBP–retinol to TTR could not be distinguished from a binding of only 10%. This shortcoming of the anisotropy technique is illustrated in Fig. 12.11. Here, the effect of HPR on RBP–TTR interaction is measured using the developed

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Fig. 12.9 Structural relationship of test retinoids to be used for validation of RBP–TTR∗ FRET assay. The chemical structures of four related retinoids are shown to illustrate similarities in β-ionone ring and saturated double-bond system and differences in geometry and side chain constituents which may affect binding to RBP and TTR.

Fig. 12.10. Validation of RBP-TTR∗ FRET assay. The ability of four structurally related retinoids to act as RBP antagonists was examined using the RBP–TTR∗ FRET assay. Test retinoids were examined at two concentrations (2 and 8 μM). Consistent with previous studies which utilized techniques such as circular dichroism, data from the RBP–TTR∗ FRET assay revealed greater antagonism by retinoids with all-trans configuration and large side-chain constituents. Thus, HPR was most potent at inhibiting RBP–TTR∗ binding while 13-cis retinoic acid was the least effective.

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Fig. 12.11. Comparison of RBP–TTR∗ FRET assay with fluorescence anisotropy. To illustrate the advantage of the RBP–TTR∗ FRET assay over fluorescence anisotropy, the effect of HPR on RBP–TTR∗ interaction was measured using these two assay techniques. Data from the RBP–TTR∗ FRET assay show that HPR inhibits RBP–TTR∗ interaction by ∼50%. Meanwhile, data from fluorescence anisotropy show no inhibition. The data clearly show that measurement of the inhibition of RBP–TTR interaction cannot be performed using fluorescence anisotropy.

RBP–TTR∗ FRET assay and by traditional fluorescence anisotropy. Although RBP–TTR binding is reduced by ∼50% in the presence of HPR, measurement by fluorescence anisotropy reports no inhibition of binding as there is still a population of intact RBP–retinol–TTR. In addition to the inherent technical impasses of fluorescence anisotropy to screen for compounds which affect RBP– TTR interaction, a technological constraint also exists. There are currently very few commercially available instruments with highthroughput capability which offer fluorescence anisotropy with detection in the near-UV range. This limitation renders fluorescence anisotropy much less desirable compared to the RBP-TTR∗ FRET assay which can be employed on any conventional fluorescent microplate reader.

2. Materials 2.1. Preparation of Recombinant RBP

1. BL21(DE3) expression cell line transformed with Human RBP/pET3a plasmid (obtained by a license with NIH).

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2. LB media containing 50 μg/ml carbenicillin. 3. IPTG stock solution 200 mM. 4. TBS (200 ml): 50 mM Tris–HCl, pH 7.5, 150 mM NaCl. 5. Lysis Buffer (100 ml): 50 mM Tris–HCl, pH 7.5, 2 mM EDTA, 2 mM PMSF, 0.1% Triton X-100. 6. Solubilization Buffer (50 ml): 25 mM Tris–HCl, pH 8.5, 8 M urea, 10 mM DTT. 7. All-trans retinol (at-ROL) stock solution: dissolved in ethanol and purged with argon in darkness to avoid oxidation and photo-bleaching; concentration determined by absorbance (ε325 nm = 52, 770 M−1 cm−1 ); stored at −20◦ C in darkness. 8. Redox Refolding Buffer (1 l): 25 mM Tris–HCl, pH 8.5, 0.5 mM cystine, 5 mM cysteine, 1 mM EDTA, protease inhibitor cocktail; degassed by nitrogen purging for 20 min; prepared fresh and kept on ice until use. 9. Pellicon XL ultrafiltration device (8 kDa MWCO) (Millipore). 10. Buffer A: 25 mM Tris–HCl, pH 8.5. 11. Buffer B: 25 mM Tris–HCl, pH 8.5, 1 M NaCl. 12. Anion Exchange Column (IEX): two tandem HiTrap Q Sepharose columns (5 ml × 2) (GE Healthcare). 13. Size Exclusion Column (SEC): Superdex 75 HiLoad 16/60 column (GE Healthcare). 14. PBS: 50 mM Na2 HPO4 , pH 7.4, 150 mM NaCl. 15. Ethyl ether. 16. Fluorimeter and 3 ml quartz cuvette. 2.2. Preparation of Recombinant TTR

1. BL21(DE3) expression cell line transformed with Human TTR/pMMHA plasmid (obtained by a license with University of California at San Diego). 2. LB media containing 50 μg/ml carbenicillin. 3. IPTG stock solution 200 mM. 4. TBS: 50 mM Tris–HCl, pH 7.5, 150 mM NaCl. 5. (NH4 )2 SO4 solid. 6. Buffer C: 25 mM Tris–HCl, pH 8.0, 0.1 M NaCl. 7. Buffer D: 25 mM Tris–HCl, pH 8.0, 1 M NaCl. 8. IEX Column: two tandem HiTrap Q Sepharose columns (5 ml × 2) (GE Healthcare). 9. Hydrophobic Interaction Column (HIC): Octyl FF HiPrep 16/10 column (GE Healthcare). 10. Buffer E: 25 mM Tris–HCl, pH 7.5, 150 mM NaCl, 1.5 M (NH4 )2 SO4 .

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11. Buffer F: 25 mM Tris–HCl, pH 7.5. 12. SEC Column: Superdex 200 HiLoad 16/60 column (GE Healthcare). 13. PBS: 50 mM Na2 HPO4 , pH 7.4, 150 mM NaCl. 2.3. Conjugation of TTR with Alexa Fluor 430 (AF-430)

1. 1 M NaHCO3 , pH 9.0. 2. Purified TTR in PBS solution: concentration determined using 0.1% M = 73, 770 M−1 cm−1 . ε280 = 1.34 mg/ml−1 cm−1 or ε280 3. Alexa Fluor 430 carboxylic acid, succinimidyl ester (AF-430) (Invitrogen) stock solution: dissolve AF-430 in DMSO, concentration determined using ε430 = 16, 000 M−1 cm−1 . 4. 1.5 M Tris–HCl, pH 8.8. 5. Desalting Column: two tandem HiTrap desalting columns (5 ml × 2) (GE Healthcare).

2.4. Fluorescence Titration of apo-RBP with Retinol

1. At-ROL stock solution: 250 μM in DMSO.

2.5. RBP–TTR∗ Screening Assay

1. Assay mixture: 0.5 μM Apo-RBP and 0.5 μM TTR∗ in PBS.

2. Apo-RBP solution: 0.5 μM in PBS. 3. Fluorimeter and 3 ml quartz cuvette.

2. 384-well black assay plate (flat bottom, non-treated). 3. At-ROL stock solution: 50 μM in DMSO. 4. Fluorescence microplate reader. 5. Test compounds (retinoid analogs) stock solution in DMSO.

3. Methods 3.1. Expression and Purification of Recombinant RBP

1. (Day 1) Inoculate single colony from LB/carbenicillin plate into 5 ml LB media. Grow cells for 2–3 h at 37◦ C with constant shaking at 250 rpm. Add the 5 ml cell culture to 100 ml LB media and continue cell growth for another 2–3 h. Split the 100 ml culture into four 500 ml LB media. Grow cells for an additional 2–4 h until OD600 reaches 0.5–0.8. Induce culture with 1 mM IPTG and harvest cells 3–4 h later (see Note 2). 2. Centrifuge cells at 5,000×g for 10 min at 4◦ C. Wash cells once by resuspending cell pellet with 200 ml ice-cold TBS and centrifuge at 7,000×g for 10 min at 4◦ C. Resuspend cell pellet with 100 ml ice-cold Lysis Buffer and store cell lysate at −80◦ C until further processing (see Note 3).

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3. (Day 2) Thaw cell lysate in room temperature water bath and sonicate cells on ice for 6 min (15 s on and 30 s off pulse intervals). 4. Centrifuge at 15,000×g for 15 min at 4◦ C and save the pellet. RBP should be in the inclusion body (see Note 4). 5. Homogenize the pellet with 50 ml Solubilization Buffer. Stir homogenate at room temperature for 1 h or longer to allow complete denaturation and release of RBP from inclusion body. Centrifuge at 30,000×g for 15 min at 20◦ C and filter the supernatant using a 0.45 μm bottle top filter. Determine total protein recovery in filtrate using Bradford Assay. Estimate relative amount of RBP (21 kDa) from SDS-PAGE (usually ∼90–95% total protein, see Note 4). 6. Under dim red lighting, add at-ROL stock solution to 1 l Redox Refolding Buffer with rapid stirring shortly before adding filtrate containing denatured RBP (use 10X molar excess of at-ROL to RBP and keep final EtOH concentration below 1%) (see Note 5). 7. Add filtrate dropwise to the stirring Redox Refolding Buffer containing at-ROL. Stir solution in darkness at 4◦ C for 5 h or longer to allow proper refolding of RBP (see Note 6). 8. (Day 3) Centrifuge the refolded RBP at 30,000×g for 20 min at 4◦ C to remove aggregated retinol and proteins. Filter supernatant using a 0.45 μm bottle top filter and concentrate filtrate using a Pellicon XL filtration device. Exchange the sample buffer to Buffer A three to five times using Pellicon XL. This process also removes excess at-ROL. Keep sample in darkness at 4◦ C for protein purification. 9. (Day 4) Load the refolded protein sample onto IEX column pre-equilibrated with Buffer A. Elute proteins in the same buffer with a salt gradient from 0 to 700 mM NaCl (Buffer B). Analyze eluted fractions by SDS-PAGE and UV/vis spectroscopy and pool fractions containing RBP (see Note 7). 10. To purify holo-RBP, concentrate the pooled fractions from step 9, filter with a 0.45 μm filter, and apply the sample to SEC column pre-equilibrated with PBS. Analyze fractions by SDS-PAGE and UV/vis spectroscopy. Pool fractions containing purified RBP. Concentrate and determine holo-RBP pro0.1% M = tein concentration (ε279 = 2.02 mg/ml−1 cm−1 or ε279 −1 42, 420 M cm−1 ). Retinol absorbs at 330 nm. The ratio A280 /A330 should be about 1 for pure holo-RBP. 11. To obtain apo-RBP, dilute the pooled fractions from step 9 to 0.5 mg/ml with PBS. Mix one volume of sample with two volumes of ethyl ether in a separatory funnel. Extract the aqueous phase and discard the organic phase. Repeat this step two more times. This extraction removes about 70–90% retinol.

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12. Purge the extracted sample with nitrogen to remove the residual ethyl ether. Keep sample on ice with constant stirring. While purging, bleach sample with a handheld UV lamp (∼360 nm) for 5–6 h. This step removes 10–20% of the remaining retinol. Further bleach sample at 4◦ C using a fluorimeter to remove the residual retinol (2–3 ml at a time for 45 min to 1 h with stirring and maximum excitation bandpass. ex: 330 nm, em: 470 nm). 13. Concentrate sample, filter with a 0.45 μm filter, and purify the apo-RBP by gel filtration as described in step 10. Pool fractions containing RBP, concentrate, and determine apo0.1% = 1.74 mg/ml−1 cm−1 or RBP protein concentration (ε279 −1 M −1 ε279 = 36, 540 M cm ). 3.2. Expression and Purification of Recombinant TTR

1. (Day 1) Inoculate single colony from LB/carbenicillin plate into 20 ml LB media. Grow cells overnight at 37◦ C with constant shaking at 250 rpm. 2. (Day 2) Inoculate the 20 ml overnight culture into four 500 ml LB media. Grow cells for 3–5 h until OD600 reaches 0.5–0.8. Induce culture with 1 mM IPTG and harvest cells 4–8 h later (see Note 8). 3. Centrifuge cells at 8,000×g for 10 min at 4◦ C. Resuspend cell pellet with 100 ml ice-cold TBS and store cell suspension at −80◦ C until further processing (see Note 9). 4. (Day 3) Thaw cell suspension in room temperature water bath and sonicate cells on ice for 6 min (15 s on and 30 s off pulse intervals). 5. Centrifuge at 10,000×g for 15 min at 25◦ C and discard the pellet. Slowly add (NH4 )2 SO4 with stirring to supernatant to 50% saturation (31.4 g for 100 ml). Stir sample for 15–20 min at room temperature after the salt has dissolved. 6. Centrifuge at 10,000×g for 10 min at 25◦ C and discard the pellet. Add (NH4 )2 SO4 to supernatant to 90% saturation (additional 30.2 g). Mix sample as in step 5 and centrifuge at 10,000×g for 10 min. Discard supernatant and resuspend pellet in 15 ml Buffer C (see Note 10). 7. Dialyze the resuspended pellet in 1 l Buffer C (exchange buffer once) overnight at 4◦ C using 10 kDa MWCO dialysis membrane. 8. (Day 4) Determine protein concentration using Bradford Assay, filter with a 0.45 μm syringe filter, and purify protein using IEX column pre-equilibrated with Buffer C. Elute proteins in the same buffer with a gradient from 0 to 40% Buffer D. Analyze eluted fractions by SDS-PAGE and pool fractions containing TTR.

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9. If TTR expression level and purity are relatively high, proceed directly to step 10. Otherwise, purify the pooled fractions by hydrophobic interaction. Add (NH4 )2 SO4 (MW: 132.14, d = 1.77 g/ml) to the pooled eluant to a final concentration of 1.5 M. Filter with a 0.45 μm filter and load content onto HIC column pre-equilibrated with Buffer E. Elute proteins with decreasing salt gradient 0–100% Buffer F. Analyze eluted fractions by SDS-PAGE and pool fractions containing TTR. 10. Concentrate and filter sample with 0.45 μm filter and further purify TTR using SEC column pre-equilibrated with PBS. Analyze fractions by SDS-PAGE and pool fractions containing purified TTR. 11. Concentrate sample and determine TTR concentration 0.1% = 1.34 mg/ml−1 cm−1 or ε M = (MW: 55 kDa, ε280 280 −1 −1 73, 700 M cm ). 3.3. Conjugation of TTR with Alexa Fluor 430 (AF-430)

1. Adjust TTR solution pH to 8.2–8.5 with NaHCO3 . 2. Add 2.5X molar excess AF-430 to TTR. Incubate reaction with constant mixing in darkness at room temperature for 1 h (see Note 11). 3. Stop reaction by adding Tris–HCl, pH 8.8, to a final concentration of 50 mM. Incubate for another hour at room temperature in darkness for excess probe to react with free amine. 4. Remove unconjugated probe with desalting column. 5. Determine protein concentration and labeling efficiency TTR = 73,700 M−1 cm−1 or using UV/vis spectroscopy (ε280 AF430 ε430 = 16, 000 M−1 cm−1 ). ODTTR = OD280 − OD430 × correction factor (0.3 for AF-430) Labeling effic iency = [AF-430]/[TTR] (see Note 12).

3.4. Fluorescence Titration of apo-RBP with Retinol

1. Under dim lighting, add at-ROL at incremental concentrations to 0.5 μM apo-RBP. Concentrations for retinol in μM: 0.1, 0.2, 0.3, 0.4, 0.5, 0.75, 1, 1.25, 1.5, 2. Assay volume: 2.5 ml in PBS in a quartz cuvette. 2. After each retinol addition, incubate at room temperature with constant stirring for 5 min and obtain RBP and RBP–retinol emissions using a fluorimeter (ex: 280 nm, em: 335 nm and 470 nm, see Note 13). 3. Apply Cogan plot to determine the binding affinity (KD ) of retinol to RBP.

3.5. Measurement of holo-RBP–TTR∗ Interaction

1. Dispense 50 μl aliquot of assay mixture in each well of a 384well plate. Use duplicate for each sample.

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2. Add 1 μl at-ROL stock solution to each well or 1 μl DMSO as negative control. Mix well and incubate at 37◦ C in darkness for 30 min. 3. Obtain fluorescence readings with a plate reader (ex: 330 nm, em: 460 nm and 580 nm). 4. For positive control, directly mix 0.5 μM holo-RBP (in place of apo-RBP) with 0.5 μM TTR∗ in the assay mixture and obtain fluorescence readings. This should yield similar results as the reconstituted RBP (see Note 14). 3.6. Identification of Competing Ligands

1. Dispense 50 μl aliquots of assay mixture as described in Section 3.5. 2. Add 1 μl of test compound at a desired concentration. Mix well and incubate at 37◦ C in darkness for 30 min. 3. Add 1 μl at-ROL stock solution. Mix well and incubate at 37◦ C in darkness for another 30 min. 4. Obtain fluorescence readings as described in Section 3.5.

4. Notes 1. It has been determined that measured fluorescence intensity is proportional to optical density up to 0.05 (15). This means that fluorescence emission intensities which are generated from solutions with optical densities above 0.05 should be corrected for a potential inner filter effect. The following equation has been developed to provide a correction factor for data obtained under these conditions (15): Fcorr = Fobs antilog[(ODex + ODem )/2)]. Application of this correction factor is particularly important in binding studies which involve retinoids as they possess rather high optical densities in the UV range. For example, a 1 μM solution of retinol possesses an optical density of ∼0.05 at its absorbance maximum (∼325 nm). Thus, fluorescence data acquired with retinol concentrations above 1 μM should be considered only after applying the appropriate correction factor. In addition, retinoids are extremely hydrophobic and, therefore, insoluble in aqueous solution. This property creates excessive light scatter at concentrations above 1 μM which can obscure the light emanating from the protein source (15).

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2. RBP expression is lost when inoculate with overnight culture. Save pre-induced and post-induced culture to examine RBP expression using SDS-PAGE analysis. 3. Store washed cell pellet at −80◦ C for long-term storage. 4. Save aliquots of all protein samples for SDS-PAGE analysis. 5. Retinol is extremely sensitive to light. All solutions should be protected from light to minimize the photo-degradation. 6. Refolding RBP in the absence of retinol results in poor yield of properly folded RBP. 7. Minimize light exposure to the column and sample fractions. 8. Cell growth is extremely slow and TTR expression is low. Save pre-induced and post-induced culture to examine TTR expression using SDS-PAGE analysis. 9. Cell suspension is stable at this stage for >1 month or store cell pellet at −80◦ C for long-term storage. 10. Pellet is stable and can be stored at −80◦ C. Save small aliquot from each step for SDS-PAGE analysis. 11. Protein concentration should be 5–20 mg/ml. Labeling efficiency will be low if concentration is <2 mg/ml. 12. Labeling efficiency is usually about 2 but should not exceed 2.5. 13. Use small excitation bandpass to minimize photo-bleaching. 14. RBP–TTR interaction can be confirmed with a secondary assay using gel filtration chromatography. Incubate 5 μM apo-RBP with 7.5 μM TTR in the presence or absence of 10 μM at-ROL in darkness for 30 min (use holo-RBP as positive control). Load reaction mixture onto a BioSil SEC125 column equilibrated with PBS (pH 7.4). Recommended flow rate is 0.5 ml/min. Monitor elution profile with a multiwavelength detector. If bound to TTR, the elution peak of at-ROL (OD330 ) and RBP (OD280 ) should co-migrate with the TTR peak (OD280 ).

References 1. Yang, Q., Graham, T.E., Mody, N., Preitner, F., Peroni, O.D., Zabolotny, J.M., Kotani, K., Quadro, L., Kahn, B.B. (2005) Serum retinol binding protein 4 contributes to insulin resistance in obesity and type 2 diabetes. Nature 436, 356–362. 2. Abahusain, M.A., Wright, J., Dickerson, J.W., de Vol, E.B. (1999) Retinol, alphatocopherol and carotenoids in diabetes. Eur. J. Clin. Nutr. 53, 630–635.

3. Basualdo, C.G., Wein, E.E., Basu, T.K. (1997) Vitamin A (retinol) status of first nation adults with non-insulin-dependent diabetes mellitus. J. Am. Coll. Nutr. 16, 39–45. 4. Cho, Y.M., Youn, B.S., Lee, H., Lee, N., Min, S.S., Kwak, S.H., Lee, H.K., Park, K.S. (2006) Plasma retinol-binding protein4 concentrations are elevated in human subjects with impaired glucose tolerance

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5.

6.

7.

8.

9.

10.

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and type 2 diabetes. Diabetes Care 29, 2457–2461. Graham, T.E., Yang, Q., Bluher, M., Hammarstedt, A., Ciaraldi, T.P., Henry, R.R., Wason, C.J., Oberbach, A., Jansson, P.A., Smith, U., et al. (2006) Retinol-binding protein 4 and insulin resistance in lean, obese, and diabetic subjects. N. Engl. J. Med. 354, 2552–2563. Munkhtulga, L., Nakayama, K., Utsumi, N., Yanagisawa, Y., Gotoh, T., Omi, T., Kumada, M., Erdenebulgan, B., Zolzaya, K., Lkhagvasuren, T., et al. (2007) Identification of a regulatory SNP in the retinol binding protein 4 gene associated with type 2 diabetes in Mongolia. Hum. Genet. 120, 879–888. Qi, Q., Yu, Z., Ye, X., Zhao, F., Huang, P., Hu, F.B., Franco, O.H., Wang, J., Li, H., Liu, Y., et al. (2007) Elevated retinol-binding protein 4 levels are associated with metabolic syndrome in Chinese people. J. Clin. Endocrinol. Metab. 92, 4827–4834. Delori, F.C., Dorey, C.K., Staurenghi, G., Arend, O., Goger, D.G., Weiter, J.J. (1995) In vivo fluorescence of the ocular fundus exhibits retinal pigment epithelium lipofuscin characteristics. Invest. Ophthalmol. Vis. Sci. 36, 718–729. Delori, F.C., Staurenghi, G., Arend, O., Dorey, C.K., Goger, D.G., Weiter, J.J. (1995) In vivo measurement of lipofuscin in Stargardt’s disease – fundus flavimaculatus. Invest. Ophthalmol. Vis. Sci. 36, 2327–2331. Schmitz-Valckenberg, S., Bultmann, S., Dreyhaupt, J., Bindewald, A., Holz, F.G., Rohrschneider, K. (2004) Fundus autofluorescence and fundus perimetry in the junctional zone of geographic atrophy in patients with age-related macular degeneration. Invest. Ophthalmol. Vis. Sci. 45, 4470–4476. Cideciyan, A.V., Aleman, T.S., Swider, M., Schwartz, S.B., Steinberg, J.D., Brucker, A.J., Maguire, A.M., Bennett, J., Stone, E.M., Jacobson, S.G. (2004) Mutations in ABCA4 result in accumulation of lipofuscin before slowing of the retinoid cycle: A reappraisal of the human disease sequence. Hum. Mol. Genet. 13, 525–534. Holz, F.G., Bellman, C., Staudt, S., Schutt, F., Volcker, H.E. (2001) Fundus autofluorescence and development of geographic

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20. 21.

22.

23.

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atrophy in age-related macular degeneration. Invest. Ophthalmol. Vis. Sci. 42, 1051–1056. Kanai, M., Raz, A., Goodman, D.S. (1968) Retinol-binding protein: The transport protein for vitamin A in human plasma. J. Clin. Invest. 47, 2025–2044. Blaner, W.S. (1989) Retinol-binding protein: The serum transport protein for vitamin A. Endocr. Rev. 10, 308–316. Lakowicz, J.R. (1999) Principles of Flourescence Spectroscopy, Kluwer Academic/Plenum, New York. Förster, T. (1948) Zwischenmolekulare energiewanderung und fluoreszenz. Ann. Physik. 437, 55–75. Stryer, L. (1978) Fluorescence energy transfer as a spectroscopic ruler. Annu. Rev. Biochem. 47, 819–846. Cogan, U., Kopelman, M., Mokady, S., Shinitzky, M. (1976) Binding affinities of retinol and related compounds to retinol binding proteins. Eur. J. Biochem. 65, 71–78. Noy, N., Xu, Z.J. (1990) Interactions of retinol with binding proteins: Implications for the mechanism of uptake by cells. Biochemistry 29, 3878–3883. Noy, N. (2000) Retinoid-binding proteins: Mediators of retinoid action. Biochem. J. 348(Pt 3), 481–495. Malpeli, G., Folli, C., Berni, R. (1996) Retinoid binding to retinol-binding protein and the interference with the interaction with transthyretin. Biochim. Biophys. Acta 1294, 48–54. Berni, R., Formelli, F. (1992) In vitro interaction of fenretinide with plasma retinolbinding protein and its functional consequences. FEBS Lett. 308, 43–45. Horwitz, J., Heller, J. (1973) Interactions of all-trans, 9-, 11-, and 13-cis-retinal, alltrans-retinyl acetate, and retinoic acid with human retinol-binding protein and prealbumin. J. Biol. Chem. 248, 6317–6324. van Jaarsveld, P.P., Edelhoch, H., Goodman, D.S., Robbins, J. (1973) The interaction of human plasma retinol-binding protein and prealbumin. J. Biol. Chem. 248, 4698–4705. Kopelman, M., Cogan, U., Mokady, S., Shinitzky, M. (1976) The interaction between retinol-binding proteins and prealbumins studied by fluorescence polarization. Biochim. Biophys. Acta 439, 449–460.

Chapter 13 Molecular Biology and Analytical Chemistry Methods Used to Probe the Retinoid Cycle Marcin Golczak, Grzegorz Bereta, Akiko Maeda, and Krzysztof Palczewski Abstract The retinoid (visual) cycle is a complex enzymatic pathway essential for regeneration of the visual chromophore, 11-cis-retinal, a component of rhodopsin that undergoes activation by light in vertebrate eyes. Pathogenic mutations within genes encoding proteins involved in the retinoid cycle lead to abnormalities in retinoid homeostasis and numerous congenital blinding diseases of humans. Thus, elucidation of disease-specific changes in enzymatic activities and retinoid content of the retina can provide important insights into the mechanisms of disease initiation and progression. Here, we use the protein RPE65 as an example to describe generally applicable methods for determining the stability and enzymatic activity of proteins and their mutants involved in retinoid metabolism. Additionally, we introduce a range of analytical techniques involving high-performance liquid chromatography and mass spectrometry to detect and quantify retinoids and their derivatives in eye extracts. Biochemical protocols combined with advanced mass spectrometry should facilitate fundamental biological studies of vision. Key words: RPE65, retinoid isomerization, mass spectrometry, A2E, retinal dimer.

1. Introduction Visual phototransduction relies upon light-induced isomerization of the visual chromophore (11-cis-retinal) coupled to rhodopsin (1). Maintaining continuous vision and preserving the health of photoreceptors require an adequate continuing supply of the chromophore. Unlike invertebrates where 11-cisretinal is regenerated by a photochemical process, vertebrates have evolved a complex enzymatic pathway called the retinoid H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_13, © Springer Science+Business Media, LLC 2010

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cycle to achieve this objective (2). This pathway operates in both the photoreceptor cells and the retinal pigment epithelium (RPE), allowing reformation of 11-cis-retinal from all-trans-retinal by several chemical transformations. The classical vertebrate retinoid cycle contributes primarily to regeneration of rhodopsin in rod cells; however, RPE65-based chromophore production may also be important for cone function (3). The retinoid cycle consists of a complex chain of enzymatic reactions employing over 20 known proteins (Fig. 13.1). Accordingly, it is susceptible to diseases resulting from mutations in genes encoding retinoid cycle proteins that cause impaired vision. In many cases, the disease is caused by loss of catalytic or retinoid-binding function (2, 4, 5). One approach to understanding the etiologies of visual impairments is to examine the enzymatic activity and stability of retinoid cycle proteins and their mutants. Substantial progress has been made in understanding the pathological basis of inactivating mutations in RPE65. Based on screening mutations in the RPE65 gene from patients with Leber’s congenital amaurosis (LCA), autosomal recessive retinitis pigmentosa (arRP), and autosomal dominant/recessive cone–rod dystrophy (CORD), more then 90 different variants were found (6, 7) (summarized in (8, 9)). Many of these were evaluated further by biochemical studies in heterologous expression systems to gain more insight into the disease mechanisms involved. The most common approach used to express RPE65 and its mutants was to generate expression vectors or recombinant adenoviruses to induce transient expression. Although both methods permit robust expression of RPE65 and detection of its enzymatic activity, quantification of 11-cis-retinol production or protein stability requires careful control of multiple variables such as transfection efficiency, viral titer, and protein expression levels. To overcome these obstacles, Bereta et al. employed an advanced bicistronic retroviral-based expression system developed initially by Kitamura and colleagues (10). Insertion of RPE65 and its mutants’ cDNA into a multi-cloning site located upstream of the internal ribosomal entry site (IRES) and an enhanced green fluorescence protein (EGFP) sequence led to expression of both the protein of interest and the EGFP from the same mRNA. This novel arrangement offers two important advantages. First is that expression of the retrovirally transduced cDNA becomes stable after integration. Transduced cells can be visualized due to EGFP co-expression and, notably, they can be sorted or subcloned based on fluorescence intensity to unify and optimize the protein expression level. The last feature is especially important for experiments focused on comparisons of enzymatic properties or stability of wild-type and mutated proteins. To ensure equivalent mRNA levels for wildtype and mutated RPE65, populations of cells characterized by identical fluorescence signals for both cell lines are used. In the

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Fig. 13.1. Schematic representation of the retinoid cycle components and formation of lipofuscin chromophores in roddominant animals (reviewed in (2)). Concomitantly with inactivation of metarhodopsin II, all-trans-retinal is released from the opsin into the lipid membranes of outer disc segments. The first catalytic step in the retinoid cycle involves enzymatic reduction of all-trans-retinal to all-trans-retinol (vitamin A) by members of the short-chain dehydrogenase/reductase (SDR) superfamily. The all-trans-retinol dehydrogenases of both rod and cone outer segments are integral membrane proteins that utilize NADPH as a cofactor. RDH8 as well as ATP-binding cassette transporter 4 (ABCR) play an important role in clearing all-trans-retinal from photoreceptor outer segments (ROS). Delay in this process leads to rapid condensation of retinal with phosphatidylethanolamine (PE) and formation of N-retinylidene-PE (N-ret-PE) that subsequently can be converted into A2E and retinal dimer that can trigger photoreceptor degeneration. Newly formed all-trans-retinol is transported across the interphotoreceptor matrix to retinal pigment epithelium (RPE) cells. There it binds to cellular retinol-binding protein-1 (CRBP1) and is esterified by lecithin:retinol acyl transferase (LRAT) that catalyzes the transfer of an acyl group from the SN1 position of phosphatidylcholine onto retinol. RPE cells store retinyl esters in lipid-droplet-like structures called retinosomes (RESTs). RESTs appear to be metabolically active because retinyl esters can be mobilized in response to light exposure and higher demand for the chromophore. Thermodynamically unfavorable isomerization of planar all-trans-retinoid to the sterically constrained 11-cis conformation is the key enzymatic step in chromophore regeneration. This reaction is catalyzed by RPE-specific 65-kDa protein (RPE65). The final catalytic step in the retinoid cycle is oxidation of 11-cis-retinol carried out by other SCAD family dehydrogenases in the RPE: 11-cis-ROL dehydrogenase type 5 (RDH5) and type 11 (RDH11). The product of this reaction, 11-cis-retinal, is bound by cellular retinal-binding protein (CRBP) that enhances transport of this chromophore through the interphotoreceptor matrix back to photoreceptor rod cells, thereby completing the retinoid cycle.

example shown in Fig. 13.2, a Gly244Val mutant of RPE65 evidenced a lower protein level as determined by immunoblotting. Considering that both cell lines, i.e., the expressing wild-type and mutated protein, had similar levels of RPE65 transcripts, the

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Fig. 13.2. Expression and enzymatic activity of RPE65 and RPE65(Gly244Val)-mutated protein. (a) Graphic representation of pMXs-IG retroviral vector used to generate the NIH3T3 cell line stably expressing RPE65 and its mutant. This vector contains the ampicillin-resistance gene (Amp r), murine leukemia retroviral long terminal repeats (5 and 3 LTR), retroviral package signal (), and multi-cloning site (MCS) for cloning of the gene of interest. Presence of an internal ribosome entry site (IRES) results in expression of the chosen protein and EGFP from a common mRNA. (b) Distribution of EGFP fluorescence intensity in LRAT/RPE65 and LRAT/RPE65(Gly244Val) expressing cell lines after sorting by a FACSAria cell sorter. Similar intensities of EGFP fluorescence indicate comparable levels of RPE65 mRNA in the tested cells. Cells expressing LRAT were prepared with pMXs-IP vector that carries a puromycin resistance gene instead of EGFP. (c) An example of expression levels of RPE65 and its mutant variant analyzed by immunoblotting. With similar mRNA levels, the lower signal from the RPE65(Gly244Val) variant protein indicates its low stability compared to wild-type protein. (d) HPLC separation of retinoids extracted from cultured RPE65-expressing cells incubated with all-trans-retinol. Saponification of the sample significantly increases 11-cis-retinol levels due to hydrolysis of 11-cis-retinyl esters. Peaks were identified based on their elution times and characteristic UV absorbance spectra: (1) retinyl esters, (2) 11-cis-retinol, (3) 13-cisretinol, and (4) all-trans-retinol.

observed difference in protein level clearly suggests lower stability and accelerated degradation of the Gly244Val variant (8). The methodology described here significantly simplifies interpretation of such results and can be applied to any protein of interest. Abnormalities in the retinoid cycle that lead to blinding diseases are often manifested by disruption of retinoid homeostasis within the eye. Thus, detection of abnormalities in retinoid metabolism provides important insights into

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possible mechanisms of disease formation and progression. Several examples of defective retinoid metabolism in the eye are exhibited by animal models of human retinal diseases. For example, Rpe65 –/– mice evidence severely attenuated rod and cone responses and progressive retinal degeneration due to inadequate regeneration of 11-cis-retinals leading to high levels of all-trans-retinyl esters which accumulate in lipid vacuoles (11, 12). Lrat –/– mice manifest undetectable levels of retinyl esters within the RPE and severely attenuated rod and cone visual responses (13, 14) and Rdh5 –/– mice exhibit enhanced accumulation of cis-retinyl esters in their RPE (15). Interestingly, deficits in proteins without enzymatic activity toward retinoids also have been shown to attenuate retinoid metabolism in the eye. Lack of retinal-specific ATP-binding cassette transporter member 4 (ABCA4) causes increased retinal levels in rod outer segments and progressive accumulation of toxic retinal derivatives, e.g., lipofuscin chromophores, within RPE cells (16, 17). Another example is Rlbp1 –/– mice (retinaldehyde-binding protein (CRALBP) knockout mice) that show delayed rates of rhodopsin regeneration and recovery of chromophore following a bleach in addition to accumulation of retinyl esters (18). Retinoid cycle impairment in Rlbp1 –/– mice is manifested by delayed dark adaptation. Quantification of chromophore levels within the retina is especially important because it provides information about the overall performance of the retinoid cycle. This approach has been widely used in combination with electroretinography to investigate potential delays in 11-cis-retinal production and rhodopsin regeneration. Analytical methods for retinoid separation have been gradually developed nearly 100 years since “fat-soluble factor B” (vitamin A) was proven to have an impact on the health and growth of young rats (19). Compounds soluble in organic solvents, including retinol and its esters, were initially separated on alumina- and silica-based stationary phases. A pivotal breakthrough was the development of modern high-performance liquid chromatography (HPLC) techniques in the early 1970s together with commercially available small size stationary phases (20, 21). The relatively mild conditions of HPLC make this technique suitable for separating and analyzing labile light-, heat-, and oxygen-sensitive retinoids. Today, retinoids can be separated under numerous chromatographic conditions (summarized in (22, 23)) optimized for normal and reverse-phase columns. Selection of the most appropriate methodology depends on the chemical properties of the particular retinoid as well as the source of the biological sample. The pool of retinoids present in a vertebrate eye is composed of retinyl esters, retinal, retinol, and their geometric (9-cis-, 11-cis-, 13-cis-, and all-trans-) isomers. Accordingly, the currently preferable method for separating these three classes of compounds, as well as their isomers, is normal-phase

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HPLC wherein hexane/ethyl acetate mixtures are used as the mobile phase (24). This methodology provides unique flexibility in tuning chromatographic conditions to allow identification and quantification of key retinoids, including 11-cis-retinol and 11-cis-retinal. Moreover, use of highly hydrophobic hexane for sample preparation simplifies the tissue extraction procedure to a one-step process without sacrificing overall analytical performance (25). A common method for retinoid detection, identification, and quantification is based on the spectral properties of these compounds. The conjugated polyene chain of retinoids contributes to their relatively strong absorption at UV and visible wavelengths. Thus, absorbance spectra provide information about the number of conjugated double bonds. Even more important, slight differences in wavelengths of maximum absorbance and shapes of the spectra allow precise identification of retinoid isomers. HPLC with UV detection is both economical and easy, providing a limit of retinoid quantification down to ∼2 pmol through photodiode array detection (Fig. 13.3). The great advantage is that the standard curves used for quantification are characterized by excellent linearity over a wide range of concentrations (2–1500 pmol) for all tested retinoids and their isomers (correlation coefficient R2 for syn 11-cis-retinal oxime is 0.9995). But a limitation of this method is the low selectivity of its UV–Vis absorbance, which mandates carefully designed chromatographic conditions

Fig. 13.3. Detection and quantification of syn 11-cis-retinal oxime. (a) Comparison of syn 11-cis-retinal oxime detection and quantification limits obtained by UV/Vis and mass spectrometry. The limit of detection is defined as a signal to noise ratio of 3:1 whereas the quantification limit corresponds to ratio of 10:1. Panels (b) and (c) represent calibration curves for mass spectrometry and UV/Vis spectrometry, respectively.

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and precise identification of the compounds being analyzed. This becomes especially challenging when multiple geometric isomers of retinoids or complex mixtures of retinyl esters with different length fatty acid chains are examined. A complementary technique allowing precise molecular identification and quantification is mass spectrometry, especially when it is combined with high-performance liquid chromatography (LC-MS). Several experimental approaches have been developed over last 15 years to detect and quantify retinoic acid, retinol, and retinyl esters in a variety of tissues. The greatest advantage of LC-MS is its sensitivity that reaches the limits of retinoid detection and quantification, depending on applied methodology and class of instrument, in the 10–50 fmol and 20–200 fmol ranges, respectively. Moreover, instruments that have the capacity

Fig. 13.4. Example of retinal dimer (a) and A2E (b) detection in mice lacking both the ATP-binding cassette transporter 4 and the retinol dehydrogenase 8. These mice develop severe RPE/photoreceptor dystrophy at an early age. Progression of retinal degeneration is correlated with a rapid increase in lipofuscin chromophore accumulation (17). Representative mass spectra and fragmentation patterns for the retinal dimer and A2E are shown on the right side of each panel.

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to perform MSn analyses provide definitive mass and structural identification. These also can be used to enhance the specificity and sensitivity of analyses done in a selected ion or selected reaction monitoring mode. Here we describe methodology that employs mass spectrometry for detection of the chromophores as well as products of retinal conjugation extracted from mouse eyes. We found that application of LC-MS technology is especially important for quantification of 11-cis-retinal in samples containing low amounts of this chromophore. Prominent examples are evaluation of 11-cis-retinal production in retinas of animal models of human ocular diseases treated with gene therapy, determination of retinoid composition in subregions of the retina, or detection of changes in the chromophore levels after light exposure in Nrl−/− mice (26). LC-MS also turns out to be an excellent tool for studying side products of retinoid metabolism, especially cytotoxic lipofuscin chromophores (Fig. 13.4). Correlations of the relative abundance of di-retinoid-pyridium-ethanolamine (A2E) or retinal dimer with age and progressive retinal degeneration may provide important insights into the mechanism(s) of human age-related macular degeneration (AMD) and Stargardt disease (16, 27). Here, we specify LC-MS conditions for the detection and quantification of retinoids optimized for normal-phase HPLC. Experimental conditions are also described for LC-MS-based detection of A2E and retinal dimer in eye extracts.

2. Materials 2.1. RPE65 Heterologous Expression and Activity Assay

1. Primers: RPE65 – GCAGATGAATTCACCATGTCGTCCCAGCC AGCAGG and CGTCTAGCGGCCGCTCAGGGCT GGGCACCATTGG. LRAT – GAGGTGAATTCAGCTACTCAGGGATGAAG AACCCCATGCTG and ACTGACGCGGCCGCATGA AGTTAGCCAGCCATCCATAG. 2. Cell growth medium (GM) consists of Dulbecco’s modified Eagle’s medium, pH 7.2, with 4 mM L-glutamine, 4,500 mg/l glucose, and 110 mg/l sodium pyruvate (HyClone Laboratories, Inc., Logan, UT) supplemented with 10% heat-inactivated fetal bovine serum, 100 units/ml penicillin, and 100 units/ml streptomycin.

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3. 0.067 M phosphate-buffered saline (PBS) composed of NaCl (9 g/l), Na2 HPO4 (0.8 g/l), NaH2 PO4 (0.14 g/l). 4. 2 M CaCl2 solution. 5. 25 mM chloroquine diphosphate salt solution in PBS. 6. 1 M potassium phosphate dibasic (K2 HPO4 ). 7. 5 M sodium chloride (NaCl). 8. 1 M potassium chloride (KCl). 9. 100 mM D dextrose. 10. 1 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), pH 7.05. 11. 5 mg/ml polybrene solution. 12. 4 M potassium hydroxide solution in methanol. 13. Hexane (HPLC grade) (Fisher Scientific, Fair Lawn, NJ). 14. 1 M hydroxylamine, pH 7.4. 15. 4 mM all-trans-retinol solution in N,N-dimethylformamide (DMF). 16. Transfection buffer composed of 50 mM HEPES, pH 7.05, containing 10 mM KCl, 12 mM dextrose, 280 mM NaCl, and 1.5 mM Na2 HPO4 . 2.2. Retinoid Extraction, Detection, and Quantification

1. Ethyl acetate (HPLC grade) (Acros, Morris Plains, NJ). 2. Ethanol (ACS grade) (Pharmco-Aaper, Brookfield, CT). 3. Acetonitrile (HPLC grade) (Fisher Scientific, Fair Lawn, NJ). 4. Isopropanol (HPLC grade) (Fisher Scientific, Fair Lawn, NJ). 5. Trifluoroacetic acid (TFA) (HPLC grade) (Supelco, Bellefonte, PA). 6. Liquid nitrogen. 7. Teflon syringe filter (National Scientific Company, Rockwood, TN). 8. Synthetic, purified standards of 11-cis-retinal oximes, alltrans-13,14-dihydro-retinal oxime, di-retinoid-pyridiumethanolamine (A2E), and retinal dimer. 9. Extraction buffer composed of 50% ethanol in 0.067 M phosphate-buffered saline, pH 7.4, containing 40 mM hydroxylamine. 10. 10% ethyl acetate in hexane. 11. 1% isopropanol in hexane. 12. 10% isopropanol in acetonitrile, 0.1% TFA.

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13. Agilent-Si, 5 μm, 4.5 × 250 mm column (Agilent Technologies, Santa Clara, CA). 14. C18 reverse-phase column, 4.5 × 250 mm (Phenomenex, Torrance, CA). 15. Other materials as in Section 2.1.

3. Methods 3.1. Evaluating Enzymatic Activity of RPE65 and Its Mutants 3.1.1. Stable Transduction and Expression of RPE65 and LRAT in NIH3T3 Cells

Both RPE65 and LRAT are essential for robust production of 11-cis-retinol in cell/tissue cultures. Thus, RPE65 should be expressed in a cell line stably producing LRAT (28). 1. Amplify RPE65 cDNA by PCR introducing EcoRI and NotI restriction sites at the beginning and the end of this sequence, respectively. 2. Clone amplified cDNAs into pMXs-IG retroviral vectors. Confirm the sequence of selected clones by DNA sequencing. 3. Twenty-four hours prior to transfection, seed 1.5 × 106 Phoenix-Eco retroviral producer cells on a 6-cm plate and culture them in 6 ml of GM. Maintain cells at 37◦ C in 5% CO2 . 4. Just prior to transfection, replace media with 6 ml of warm media containing 25 μM chloroquine. 5. Generate a transfection mixture by adding 1 ml of transfection buffer to an equal volume of 250 mM CaCl2 solution containing 15 μg of plasmid DNA. Mix and incubate for 1 min. Add mixture dropwise to the cells and place the cells in 32◦ C, 5%CO2 incubator (see Note 1). 6. Replace the medium with fresh GM at both 8 and 24 h after transfection. 7. Harvest released retrovirus by collecting cell medium 48 h post-transfection; centrifuge at 500×g for 5 min to remove detached cells. Aliquot the retrovirus-containing medium into 0.5-ml portions, freeze in liquid nitrogen, and store at −80◦ C. 8. Twenty-four hours prior to transduction, suspend 2 × 105 of NIH3T3 fibroblasts expressing LRAT in 6 ml of GM and then place the suspension in a 6-cm dish. 9. For transduction, remove 2.5 ml of the medium and mix it with 0.5 ml of viral supernatant. Add polybrene to a final

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concentration of 5 μg/ml and replace the remaining media with the virus mix. Incubate for 24 h at 32◦ C in 5% CO2 before replacing the medium with fresh GM. Split the cells upon reaching confluence and repeat the infection cycle if necessary. 10. Sort the cells based on their expression of EGFP protein by using the FACSAria automated sorter. Prepare cell suspensions containing 5 × 106 cells per 0.5 ml of GM + 5% serum. 3.1.2. Cell Culture Retinoid Isomerization Assay

1. Twenty hours prior to the experiment, plate NIH3T3 cells expressing LRAT and RPE65 in 6-well culture plates at a density of 1 × 106 cells per well. 2. Replace GM with 2 ml of fresh GM containing 10 μM of all-trans-retinol delivered in DMF. Protect cells from white light by covering the plates with aluminum foil. 3. After 16 h of incubation, collect medium and cells separately. Add 2 ml of 4 M KOH in methanol, homogenize the sample in a glass–glass homogenizer, and incubate at 50◦ C for 2 h to hydrolyze retinyl esters. 4. Add 4 ml of hexane and extract retinoids by vigorous shaking for 5 min. 5. Separate the organic and aqueous phases by centrifugation (4000×g, 5 min); carefully collect the hexane phase and place it in a glass test tube (see Note 2). 6. Dry the hexane extract by using a SpeedVac or a stream of inert gas. Redissolve retinoids in 250 μl of hexane and place the mixture in an HPLC vial. The sample now is ready to be analyzed by HPLC under conditions described in Section 3.2.1.

3.2. Sample Preparation and Chromatography Conditions for Retinoid Separation

All procedures described in this and the next section should be carried out in a dark room under a dim red light safe lamp. All chromatographic separations of retinoids described were performed with an Agilent 1100 series binary liquid chromatograph gradient system equipped with a diode array detector and HP Chemstation software. This allowed identification of retinoid isomers by their specific retention times and absorption maxima relative to chemically synthesized standards.

3.2.1. Extraction, Detection, and Quantification of Retinyl Esters, Retinols, and Retinals

Normal-phase chromatography offers better resolution than reverse-phase chromatography for separating retinoid isomers. The former technique is suitable for determining the chromophore levels in eye extracts and quantifying in vitro 11-cisretinol production in retinoid isomerization assays. UV absorption detection offers high analytical specificity because only a

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limited number of compounds extractable from tissues share a spectral absorbance range with retinoids. Retinoid analyses were developed for normal-phase chromatography by using AgilentSi, 5 μm, 4.5 × 250 mm column. Because retinals with a reactive aldehyde group may couple with amine groups of lysine and phosphoethanolamine, hydroxylamine was used to convert retinals into their stable oxime derivatives (syn- and anti-). This strategy ensures efficient extraction and accurate quantification of these reactive compounds (see Notes 3 and 4). 1. Collect a whole mouse eye, put it in an Eppendorf tube, and immediately flash freeze it in liquid nitrogen. Frozen eye lenses are easier to homogenize. 2. Transfer the eye into a glass–glass homogenizer, add 0.5 ml of ice-cold extraction buffer, and homogenize immediately. 3. Incubate the sample for 20 min at room temperature with occasional shaking. 4. Add 2 ml of hexane and extract retinoids. Prepare sample for HPLC as described in Section 3.1.2, points 5 and 6. 5. Set wavelengths at which diode array detector will monitor retinoid elution. These usually are 325 and 360 nm for retinyl esters/retinols and retinals/retinal oximes, respectively. 6. Elute retinoids with an isocratic flow of 10% ethyl acetate in hexane at flow rate of 1.4 ml/min for 30 min to achieve baseline separation of 9-cis-, 11-cis-, 13-cis-, and all-transretinal oximes, as well as the corresponding isomers of retinol. Because of a significantly higher hydrophobicity of retinyl esters, resolution of their isomers and acyl chain variants can be achieved with a lower ethyl acetate concentration (0.5%) in hexane at the same flow rate (see Note 5). Detect retinal oximes at 360 nm. 7. Quantify 11-cis-retinal oximes and 11-cis-retinol based on standard curves that correlate integrated peak areas calculated from chromatograms with known amounts of synthetic standards injected onto the column. 3.2.2. Extraction and HPLC Separation of A2E

1. Mechanically homogenize a mouse eye in 1 ml of acetonitrile with a kinematic polytron homogenizer (PT 1200). 2. Extract the sample twice with 1 ml of acetonitrile. Centrifuge at 13,000×g for 2 min. 3. Collect and transfer the solvent to a glass test tube and dry it down in a SpeedVac. 4. Redissolve the sample in 150 μl of acetonitrile and 0.1% TFA and filter the mixture through a teflon syringe filter.

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5. Load the sample (up to 100 μl) onto a C18 reverse-phase HPLC column, 4.5 × 250 mm and analyze with a linear gradient of acetonitrile in water (80–100%), 0.1% TFA for 30 min. Monitor elution of A2E at 435 nm (see Note 6). 6. Quantify A2E based on known concentrations of pure synthetic A2E standard. 3.2.3. Extraction and Chromatographic Conditions for Retinal Dimer

1. Homogenize a mouse eye as described in Section 3.2.1. 2. Extract retinal dimer with 3 ml of a hexane:ethyl acetate mixture (2:1). 3. Evaporate the solvent; redissolve dried extracts in 150 μl of 10% isopropanol in acetonitrile with 0.1% TFA. 4. Pass the solution through a teflon syringe filter prior to loading the sample on the HPLC. 5. Analyze the sample on a Phenomenex C18 reverse-phase column, 4.5 × 250 mm, with a gradient of isopropanol in acetonitrile (0–25%) in the presence of 0.1% formic acid for 20 min at a flow rate of 1 ml/min. 6. Monitor elution of retinal dimer at 430 nm and quantify it based on the standard curve derived from a synthesized standard.

3.3. LC-MS Analysis of 11-cis-Retinal and Lipofuscin Chromophores

Acquire mass spectra by using a LXQ high-throughput linear ion trap mass spectrometer interfaced with an atmospheric pressure chemical ionization (APCI) source and a series 1100 HPLC system consisting of a vacuum degasser, a binary pump, an autosampler with a cooled sample tray, a thermostatically controlled column compartment, and a diode array detector. The APCI source was chosen for development of LC-MS methodology because of its wide dynamic range and capacity to operate at the high flow rates required for HPLC retinoid separation (29).

3.3.1. The Chromophore

1. Tune the mass spectrometer to ensure optimal operating parameters by using flow injection of the retinoid standard in a mobile phase and a flow rate corresponding to that for its elution from the HPLC column. Positive-ion APCI is more sensitive for retinoid analyses and can be used for all applications. Optimal mass spectrometer parameters for chromophore detection include sheath, aux, and sweep gas flow set at 20, 5, and 0, respectively; capillary temperature and voltage of 275◦ C and 2 V; vaporized temperature 380◦ C; tube lens voltage 70 V; and corona discharge current 5 μA. The base peak at m/z = 300 corresponding to retinal oximes [M + H]+ is recorded by using the selected ion monitoring (SIM) mode.

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2. Separate retinal oximes on an Agilent-Si, 5 μm, 4.5 × 250 mm normal-phase column equilibrated with 1% isopropanol in hexane at a fixed flow rate of 1.4 ml/min. 3. Prepare calibration curves by using external or internal standard methodology. To ensure the highest precision for the external standard quantification technique, spike eye extracts from Rpe65 –/– mice (having undetectable levels of 11-cisretinoids) with known amounts of 11-trans-retinal oximes and extract as described in Section 3.2.1. Construct a calibration curve by plotting the magnitude of the detector response as a function of the external standard concentration (see Note 7). The standard curve should evidence linearity over a 0.1–500 pmol range. Alternatively, to reduce errors generated by potential system instability or variations in source conditions, use commercially available all-trans-13,14-dihydro-retinal oximes or all-trans-3,4dihydro-retinal oximes (m/z = 288 [M + H]+ ) that differ from natural retinal oximes only by saturation of a double bond, as internal standards. Add and mix a fixed amount of internal standard to each sample prior to extraction. After samples are prepared and analyzed, the quantity of the target compound can be determined from the calibration curve that displays the ratio of the target compound and the standard detector responses as a function of the amount of retinal oxime injected on the column. 4. Inject a sample and perform online HPLC separation of retinoids followed by APCI positive ionization mode mass spectrometric detection. 5. Confirm identification of the detected product by referring to the characteristic fragmentation pattern of its parent ion. 6. Determine the areas under peaks of interest. Quantify analytes in relation to internal standards by using one of the methods described above. 3.3.2. Identification of A2E in a Mouse Eye

1. Optimize conditions for A2E detection by using a synthetic standard and the mobile phase required for optimal A2E resolution on a C-18 column. Optimum APCI conditions include sheath, aux, and sweep gas flow fixed at 20, 5, and 0 units, respectively; capillary temperature and voltage of 275◦ C and 46 V; vaporized temperature 450◦ C; tube lens voltage 105 V; and corona discharge current 5 μA. 2. Inject 100 μl of an acetonitrile mouse eye extract onto a reverse-phase column as described in Section 3.2.2 and perform retinoid separation. For the LC-MS application, reduce concentration of TFA in the mobile phase to 0.05%.

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3. Carry out analysis by using the SIM mode at m/z = 592.4 [M + H]+ . 4. Confirm detection of A2E and its isomers by monitoring the unique fragmentation pattern of the parent ion.

3.3.3. Detection of Retinal Dimer by Mass Spectrometry

1. Tune the mass spectrometer by using the purified synthetic retinal dimer standard under chromatographic conditions required for its elution. 2. Separate retinal dimer extracted from a mouse eye according to conditions described in Section 3.2.3. 3. Use the SIM mode to detect the mass at m/z = 551.4 that corresponds to the retinal dimer parent ion [M+H]+ (see Note 8). 4. Confirm detection of retinal dimer by recording the MS2 fragmentation pattern of the parent ion.

4. Notes 1. All buffers for transfecting cells need to be sterilized by filtration (0.2 μm filters) and stored at −80◦ C. The pH of HEPES buffer should be adjusted carefully to 7.05 ± 0.05. This parameter is critical for transfection efficiency. 2. Collect organic (hexane) phase carefully. Even small traces of water in a sample prepared for normal-phase HPLC may destroy a silica column. 3. Retinoids are sensitive to light, heat, and oxygen. So they should be protected from light and stored in organic solvents rather than in dried form. 4. Hydroxylamine solution is unstable so it should always be freshly prepared. Adjust pH of 1 M stock solution to 7.4 by adding concentrated HCl before use. 5. Isopropanol used for retinoid separation in LC-MS experiments provides better sensitivity compared to ethyl acetate utilized under standard conditions. The disadvantage of this approach is that 1% isopropanol in hexane does not permit chromatographic separation of 11-cis- and 13-cis-retinol isomers. 6. We found that TFA is required for chromatography of A2E. Although TFA may reduce mass spectrometer signal intensity, it should not be replaced with formic acid.

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7. Ionization of retinoids does not depend on the geometry of these molecules. Thus, more available all-trans isomers can be used as standards for mass spectrometry-based retinoid quantification (23). 8. Retinal dimer is extremely sensitive to oxidation. Protection from light and excess air is required to prevent formation of a variety of epoxy and furano retinal dimer derivatives.

Acknowledgment This research was supported in part by grants EY009339 and P30 EY11373 from the National Institutes of Health and the Foundation Fighting Blindness. References 1. Palczewski, K. (2006) G protein-coupled receptor rhodopsin. Annu. Rev. Biochem. 75, 743–767. 2. Travis, G.H., Golczak, M., Moise, A.R., Palczewski, K. (2007) Diseases caused by defects in the visual cycle: Retinoids as potential therapeutic agents. Annu. Rev. Pharmacol. Toxicol. 47, 469–512. 3. Jacobson, S.G., Aleman, T.S., Cideciyan, A.V., Heon, E., Golczak, M., Beltran, W.A., Sumaroka, A., Schwartz, S.B., Roman, A.J., Windsor, E.A., Wilson, J.M., Aguirre, G.D., Stone, E.M., Palczewski, K. (2007) Human cone photoreceptor dependence on RPE65 isomerase. Proc. Natl. Acad. Sci. USA 104, 15123–15128. 4. Thompson, D.A., Gal, A. (2003) Vitamin A metabolism in the retinal pigment epithelium: Genes, mutations, and diseases. Prog. Retin. Eye Res. 22, 683–703. 5. McBee, J.K., Palczewski, K., Baehr, W., Pepperberg, D.R. (2001) Confronting complexity: The interlink of phototransduction and retinoid metabolism in the vertebrate retina. Prog. Retin. Eye Res. 20, 469–529. 6. Gu, S.M., Thompson, D.A., Srikumari, C.R., Lorenz, B., Finckh, U., Nicoletti, A., Murthy, K.R., Rathmann, M., Kumaramanickavel, G., Denton, M.J., Gal, A. (1997) Mutations in RPE65 cause autosomal recessive childhood-onset severe retinal dystrophy. Nat. Genet. 17, 194–197. 7. Morimura, H., Fishman, G.A., Grover, S.A., Fulton, A.B., Berson, E.L., Dryja, T.P. (1998) Mutations in the RPE65 gene in patients with autosomal recessive retinitis pigmentosa or Leber congenital

8.

9.

10.

11.

12.

13.

amaurosis. Proc. Natl. Acad. Sci. USA 95, 3088–3093. Bereta, G., Kiser, P.D., Golczak, M., Sun, W., Heon, E., Saperstein, D.A., Palczewski, K. (2008) Impact of retinal disease-associated RPE65 mutations on retinoid isomerization. Biochemistry 47, 9856–9865. Perrault, I., Rozet, J.M., Gerber, S., Ghazi, I., Leowski, C., Ducroq, D., Souied, E., Dufier, J.L., Munnich, A., Kaplan, J. (1999) Leber congenital amaurosis. Mol. Genet. Metab. 68, 200–208. Kitamura, T., Koshino, Y., Shibata, F., Oki, T., Nakajima, H., Nosaka, T., Kumagai, H. (2003) Retrovirus-mediated gene transfer and expression cloning: Powerful tools in functional genomics. Exp. Hematol. 31, 1007–1014. Redmond, T.M., Yu, S., Lee, E., Bok, D., Hamasaki, D., Chen, N., Goletz, P., Ma, J.X., Crouch, R.K., Pfeifer, K. (1998) Rpe65 is necessary for production of 11-cis-vitamin A in the retinal visual cycle. Nat. Genet. 20, 344–351. Imanishi, Y., Batten, M.L., Piston, D.W., Baehr, W., Palczewski, K. (2004) Noninvasive two-photon imaging reveals retinyl ester storage structures in the eye. J. Cell. Biol. 164, 373–383. Batten, M.L., Imanishi, Y., Tu, D., Doan, T., Zhu, L., Pang, J., Glushakova, L., Moise, A.R., Baehr, W., Van Gelder, R.N., Hauswirth, W.W., Rieke, F., Palczewski, K. (2005) Pharmacological and rAAV gene therapy rescue of visual functions in a blind mouse model of Leber congenital amaurosis. PLoS Med. 2, e333.

Chemical Analysis of Retinoid Cycle 14. Batten, M.L., Imanishi, Y., Maeda, T., Tu, D.C., Moise, A.R., Bronson, D., Possin, D., Van Gelder, R.N., Baehr, W., Palczewski, K. (2004) Lecithin-retinol acyltransferase is essential for accumulation of all-trans-retinyl esters in the eye and in the liver. J. Biol. Chem. 279, 10422–10432. 15. Driessen, C.A., Winkens, H.J., Hoffmann, K., Kuhlmann, L.D., Janssen, B.P., Van Vugt, A.H., Van Hooser, J.P., Wieringa, B.E., Deutman, A.F., Palczewski, K., Ruether, K., Janssen, J.J. (2000) Disruption of the 11-cis-retinol dehydrogenase gene leads to accumulation of cisretinols and cis-retinyl esters. Mol. Cell Biol. 20, 4275–4287. 16. Weng, J., Mata, N.L., Azarian, S.M., Tzekov, R.T., Birch, D.G., Travis, G.H. (1999) Insights into the function of Rim protein in photoreceptors and etiology of Stargardt’s disease from the phenotype in abcr knockout mice. Cell 98, 13–23. 17. Maeda, A., Maeda, T., Golczak, M., Palczewski, K. (2008) Retinopathy in mice induced by disrupted all-transretinal clearance. J Biol Chem 283, 26684–26693. 18. Saari, J. C., Nawrot, M., Kennedy, B.N., Garwin, G.G., Hurley, J.B., Huang, J., Possin, D.E., Crabb, J.W. (2001) Visual cycle impairment in cellular retinaldehyde binding protein (CRALBP) knockout mice results in delayed dark adaptation. Neuron 29, 739–748. 19. McCollum, E.V., Davis, M. (1913) The necessity of certain lipins in the diet during growth. J. Biol. Chem. 15, 167–175. 20. Vecchi, J., Vesely, J., Oesterhelt, G. (1973) Applications of high-pressure liquid chromatography and gas chromatography to problems in vitamin A analysis. J. Chromatogr. 83, 447–453. 21. Rotmans, J.P., Kropf, A. (1975) The analysis of retinal isomers by high speed liquid chromatography. Vision Res. 15, 1301–1302.

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22. Kane, M.A., Folias, A.E., Napoli, J.L. (2008) HPLC/UV quantitation of retinal, retinol, and retinyl esters in serum and tissues. Anal. Biochem. 378, 71–79. 23. Kane, M.A., Folias, A.E., Wang, C., Napoli, J.L. (2008) Quantitative profiling of endogenous retinoic acid in vivo and in vitro by tandem mass spectrometry. Anal. Chem. 80, 1702–1708. 24. Van Hooser, J.P., Liang, Y., Maeda, T., Kuksa, V., Jang, G.F., He, Y.G., Rieke, F., Fong, H.K., Detwiler, P.B., Palczewski, K. (2002) Recovery of visual functions in a mouse model of Leber congenital amaurosis. J. Biol. Chem. 277, 19173–19182. 25. Van Hooser, J.P., Garwin, G.G., Saari, J.C. (2000) Analysis of visual cycle in normal and transgenic mice. Methods Enzymol. 316, 565–575. 26. Mears, A.J., Kondo, M., Swain, P.K., Takada, Y., Bush, R.A., Saunders, T.L., Sieving, P.A., Swaroop, A. (2001) Nrl is required for rod photoreceptor development. Nat. Genet. 29, 447–452. 27. Fishkin, N.E., Sparrow, J.R., Allikmets, R., Nakanishi, K. (2005) Isolation and characterization of a retinal pigment epithelial cell fluorophore: An all-trans-retinal dimer conjugate. Proc. Natl. Acad. Sci. USA 102, 7091–7096. 28. Golczak, M., Maeda, A., Bereta, G., Maeda, T., Kiser, P.D., Hunzelmann, S., von Lintig, J., Blaner, W.S., Palczewski, K. (2008) Metabolic basis of visual cycle inhibition by retinoid and nonretinoid compounds in the vertebrate retina. J. Biol. Chem. 283, 9543–9554. 29. van Breemen, R.B., Nikolic, D., Xu, X., Xiong, Y., van Lieshout, M., West, C.E., Schilling, A.B. (1998) Development of a method for quantitation of retinol and retinyl palmitate in human serum using high-performance liquid chromatographyatmospheric pressure chemical ionizationmass spectrometry. J. Chromatogr. A 794, 245–251.

Chapter 14 Visualization of Retinoid Storage and Trafficking by Two-Photon Microscopy Yoshikazu Imanishi and Krzysztof Palczewski Abstract Vertebrate vision is maintained by the retinoid (visual) cycle, a complex enzymatic pathway that operates in the retina to regenerate the visual chromophore, 11-cis-retinal, a prosthetic group of rhodopsin that undergoes activation by light. Many different mutations in genes encoding retinoid cycle proteins can cause a variety of human blinding diseases. Two-photon microscopy is an evolving, non-invasive, and repetitive imaging technology that can be used to monitor biomolecules within the vertebrate retina at a subcellular resolution. This method has the great advantage of portraying endogenous retinoid fluorophores in their native state without the need for artificial staining. Such real-time retinal imaging permits rapid evaluation not only of various stages of retinal disease in live animal models of human retinopathies but also of the outcome from intended pharmacological therapies. Two-photon microscopy offers substantial potential for early detection of age- and disease-related changes in the eye, long before clinical or pathological manifestations become apparent. Key words: Rhodopsin, retinoid isomerization, retinol, two-photon microscopy, RPE, retina, lipid droplet, retinosome. Abbreviations: The abbreviations used are as follows HPLC high pressure liquid chromatography NA numerical aperture REST retinyl ester storage structure (retinosome) RPE retinal pigmented epithelium TPM two-photon microscopy UV ultraviolet light.

1. Introduction Retinoids are signaling molecules essential for a number of biological processes including development, immunity, and vision. One active form of retinoids, 11-cis-retinal, is the chromophore H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_14, © Springer Science+Business Media, LLC 2010

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of rhodopsin and cone visual pigments (1). For each photon absorbed by rhodopsin, a single molecule of 11-cis-retinal is converted to all-trans-retinal. Since retinoids cannot be synthesized in the body, they must be derived from dietary sources. Dietary retinoids are temporarily stored as retinyl esters, primarily in liver, and then used to generate active retinoid species, including 11-cisretinal. To maximize their usage of retinoids, eyes have evolved a unique recycling mechanism for the replenishment of visual pigment chromophores during continuous illumination. Conversion of all-trans back to 11-cis-retinal requires a series of biochemical reactions occurring in the photoreceptors and subsequently in the retinal pigmented epithelium (RPE) (2, 3). The cycle of reactions required for the regeneration of 11-cis-retinal is called the retinoid (or visual) cycle, and deficiencies in its components can lead to a variety of blinding disorders (4, 5). Among the retinoid intermediates of the cycle, fatty acid all-trans-retinyl esters are key substrates for an isomerase RPE65. From all-transretinyl esters, RPE65 generates 11-cis-retinol (6–8), which then is oxidized to 11-cis-retinal by retinol dehydrogenases (9–11). Since only the RPE is capable of generating 11-cis-retinoids, efficient trafficking of retinoids between photoreceptor cells and the RPE is required to complete the retinoid cycle. Currently, studies of the retinoid cycle largely depend on high pressure liquid chromatography (HPLC)-based assays (12). Although HPLC is an excellent tool for quantification and identification of retinoid compounds, it lacks the spatiotemporal resolution essential for understanding their metabolism and trafficking in a subcellular structural context. To study directly the roles of retinoid trafficking in vision, we sought a methodology to monitor retinoid flow in intact mouse eyes at a resolution conferred by fluorescence microscopy (<1 μm). Intrinsic fluorescence of retinols and retinyl esters is often used in analytical chemistry to identify different retinoid species (12). However, the drawback of fluorescent imaging of retinol and retinyl esters, from a biological standpoint, is that the excitation light required for their activation lies in the ultraviolet (UV) range (∼325 nm). Short-wavelength UV light is prone to scatter and get absorbed by biological molecules, resulting in less efficient excitation especially in thick (50–200 μm) biological tissue samples. Also, large doses of UV light can destroy cells and may cause cataracts in the lens. To circumvent these unfavorable properties, we introduced the application of two-photon microscopy (TPM) (see Note 1), in which excitation of a fluorophore is accomplished by nearly simultaneous absorption of two photons in the infrared region (13). Infrared light as used in TPM is considered less toxic and can penetrate tissues better than UV light (14–16). TPM, similar to confocal microscopy, permits monitoring of three-dimensional distributions of fluorescent

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molecules. But in contrast to confocal microscopy, excitation of molecules in TPM is limited within a focal spot of less than 1 fl volume (17). Because of this highly localized two-photon excitation, optical sectioning is possible by scanning specimens, without a confocal pinhole in front of the photon detectors. Both for TPM and other conventional microscopes, high-resolution imaging requires the use of a high numerical aperture (NA) objective lens. Just as important, the efficacy of twophoton excitation is dependent on the fourth power of the NA (18), so the signal-to-noise ratio improves dramatically by using an objective lens with a higher NA. However, high NA lenses generally have short working distances and cannot reach deep inside a specimen. We found that two-photon imaging of the RPE can be accomplished by applying short-pulsed illumination through the sclera and the choroid of mouse eyes (Fig. 14.1A). Under these imaging conditions and with the involved target geometry, the working distance can be kept as short as 50–100 μm to reach the RPE layer. By taking advantage of this short distance, we successfully applied an objective lens with a NA ranging from 0.7 to 1.3 to imaging the RPE of mouse eyes. TPM with excitation at ∼730 nm was applied to RPE cells ex vivo to learn how fatty acid retinyl esters are organized in these cells (Fig. 14.1B, a). The emission signal was collected

Fig. 14.1. Transscleral imaging of mouse RPE. (A) A diagram illustrating the use of transscleral two-photon imaging to study the RPE and retina. Non-pigmented mouse eyes need to be used for this application. The objective lens is located proximal to the surface of the sclera. The black arrow indicates the direction of the excitation light. Because of the short distance between the objective lens and the RPE cells, highresolution imaging is possible with a high NA lens. (B) Imaging of retinyl ester fluorescence in the RPE of isolated mouse eyes. At low-power two-photon excitation (top image), retinyl ester storage structures are visible proximal to plasma membranes. At high-power two-photon excitation (bottom image), less abundant diffusively located retinoids become visible. Nuclei do not show a fluorescence signal, suggesting that retinoids localize to the cytoplasmic area. Top and bottom images were collected from the same group of RPE cells. Scale bar, 20 μm.

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through a custom-made bandpass filter (HQ 465/160, Chroma Technology Corp., Rockingham, VT) to monitor fluorescence in the 385–545 nm range. Previous studies indicated that fatty acid retinyl esters are compartmentalized in small subcellular structures we named retinyl ester storage structures (RESTs, also called retinosomes) (19) that were visualized by excitation at an intensity of ∼3 mW (Fig. 14.1B, a). Additional diffuse fluorescence in the cytoplasmic area was noted after increasing the excitation intensity to ∼30 mW (Fig. 14.1B, b), suggesting that the observed fluorescence originates from retinyl esters. To identify the molecular origin of the observed fluorescence, we compared the fluorescence changes in subcellular structures before (Fig. 14.2A, a) and after (Fig. 14.2A, b) application of all-transretinol to the RPE cells. Not only was an increase of fluorescence observed in the RPE cells (Fig. 14.2A, c) but it also correlated with an increased level of all-trans-retinyl esters found in the specimen (Fig. 14.2A, d). In conjunction with the lack of RPE fluorescence in mice deficient in the formation of retinyl esters (19), we conclude that the fluorescence seen in the RPE mainly originates from retinyl esters. Two-photon retinyl ester imaging can be combined with immunofluorescence confocal microscopy. Localization of alltrans-retinyl esters was stable after fixation with 4% paraformaldehyde for 30 min and subsequent incubation in the presence of

Fig. 14.2. Studies of retinyl ester storage structures (RESTs). (A) (a) RPE cells in an eyecup preparation. (b) RPE cells in an eyecup preparation after application of all-trans-retinol. Newly formed all-trans-retinyl esters localized into the RESTs. (c) Fluorescence in the RPE quantified before and after treatment with all-trans-retinol. Fluorescence increased after the treatment. (d) All-trans-retinyl esters in the eyecups quantified before and after treatment with all-trans-retinol. All-trans-retinyl esters increased after treatment. (B) Two-photon imaging of RESTs reveals colocalization of Adfp. Adfp (green fluorescence) specifically localized to the RESTs (red fluorescence) as shown by yellow color in the merged image. Adapted from Imanishi et al. (19). Scale bars, 20 μm. This research was originally published in the Journal of Cell Biology (19). The Rockefeller University Press.

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0.1% Triton X-100 for over 3 h. Because of this stability, eyecups can be processed for immunofluorescence staining without compromising the arrangement of RESTs. Figure 14.2B indicates the localization of adipose differentiation-related protein (Adfp) (20), a component frequently found on lipid droplets. Adfp (Fig. 14.2B, green color) co-localized with RESTs (Fig. 14.2B, red color) in the RPE cells, suggesting its interaction with retinyl esters. As indicated by the example of Adfp, this type of combined analysis is especially useful for discovering new components of RESTs. The most advantageous capability of TPM is monitoring retinyl ester dynamics in vivo. In this regard, the described application provides an advantage over previously published methods for imaging retinoids. In the 1980s, Kaplan introduced UVexcitation fluorescence microscopy to monitor formation of alltrans-retinol after photobleaching rhodopsin (21). More recently, Koutalos and his colleagues used a more sensitive fluorescence microscope to monitor the in situ formation and diffusion of all-trans-retinol in photoreceptor outer segments (22–24). However, all of these studies were performed on isolated photoreceptor segments or retina slices that lack photoreceptor–RPE contacts. An intact photoreceptor–RPE interface is essential for the proper transport of all-trans-retinol from photoreceptors to RPE cells. These contacts are maintained in either isolated eyes or anesthetized mice used for our studies. In an anesthetized mouse (Fig. 14.3A), transport and subsequent incorporation of retinoid from photoreceptors into the RPE cells can be documented (Fig. 14.3B). Immediately after photoactivation of rhodopsin (Fig. 14.3B, left), RESTs showed relatively weak fluorescence. However, 30 min after photoactivation (Fig. 14.3B, right), RESTs showed strong fluorescence. Retinyl ester levels increased in the RPE due to formation of all-trans-retinol in photoreceptors, trafficking of all-trans-retinol to the RPE, and esterification of all-trans-retinol by lecithin-retinol acyltransferase (19, 25). The increase of the fluorescence showed a time course similar to the increase of retinyl esters in vivo, indicating that the RPE cells were functional during this imaging procedure (19). Two-photon imaging of RPE is not just limited to mouse models but also is applicable to other species including humans. A human RPE layer was fixed after removing the retina from an eyecup, and the RPE layer was dissected out along with the choroid layer. Small pieces (<1 cm2 ) of tissue then were mounted on glass-bottomed 35-mm dishes (MatTek Corp.). RPE cells corresponding to the locations 1–4 shown in Fig. 14.4A were imaged under the same conditions. In the area proximal to the edge of retina (Fig. 14.4B, 1), RPE cells had irregular shapes and punctate fluorescent structures were observed throughout these cells. In equatorial, macular, and foveal regions (Fig. 14.4B,

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Fig. 14.3. Two-photon imaging of RPE cells in vivo. (A) A diagram illustrating twophoton imaging of mouse RPE cells in vivo. One eye of an anesthetized mouse is rotated to image the peripheral part of the RPE layer and the eye is immobilized on a coverglass with cyanoacrylate glue. An infrared short pulse laser beam (in red color) is passed through the sclera and focused on the RPE. (B) RPE cells in the peripheral region imaged by TPM. Retinyl ester fluorescence was imaged at 0 min (left) and 30 min (right) after rhodopsin stimulation by light. Retinyl ester fluorescence increased at 30 min. Images are adapted and modified from Imanishi et al. (25). Scale bar, 20 μm. This research was originally published in the Journal of Biological Chemistry (25). © The American Society for Biochemistry and Molecular Biology.

2–4), these fluorescent structures were arranged in a more orderly fashion proximal to the border of RPE cells. Considerable variation in absolute fluorescence intensities was observed amongst the eye specimens tested, irrespective of the subject’s age. However, the macular RPE always exhibited the highest fluorescence intensities in each eye, whereas the equatorial and foveal RPE showed about threefold weaker fluorescence intensities (Fig. 14.4C). The macular region contains the highest density of photoreceptors (26) so this result is consistent with the high densities of retinoids in the macular region of primate eyes (27). Studies of RESTs should provide a new avenue for understanding the link between lipid storage and the retinoid cycle (19). Methods described in Section 3.4 (Fig. 14.2) will allow identification of novel REST components whereas the method in

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Fig. 14.4. Imaging of human RPE by two-photon microscopy. (A) Schematic drawing of a cross section of human eye. Arrows with numbers 1–4 indicate locations of RPE imaged by TPM. (B) Two-photon imaging of human RPE cells. Retinosome-like structures can be seen proximal to the plasma membrane. (C) Relative fluorescence intensities measured at different regions of four human eyes. Fluorescence intensities in the equator (position 2 in A) and fovea (position 4 in A) were normalized to intensities in the macula. Scale bar, 20 μm.

Section 3.5 (Fig. 14.3) can be used to study the storage and trafficking of retinyl esters in animal models deficient in REST components. One REST component examined in this manner, Adfp protein, was found to be involved in the storage and trafficking of retinyl esters in the RPE (25). Another application with far-reaching implications is the use of TPM for the diagnosis of diseases caused by dysfunction of RPE cells. Involved diseases include, but may not be limited to, retinitis pigmentosa, Leber’s congenital amaurosis, cone–rod dystrophy, and macular degeneration. In mice with deficiencies in the retinoid cycle, obvious changes in either the dynamics or fluorescence intensities of RESTs were observed (19, 28). Thus it will be interesting to apply the TPM method in Section 3.6 (Fig. 14.4) to RPE cells isolated from patients with inherited eye diseases. Such studies will allow us to monitor how the disease affects normal retinoid metabolism of RPE cells. In the future, it would be desirable to develop a two-photon imaging technology for non-invasive imaging of living human eyes. This would expedite following age-dependent changes of retinoid metabolism in the same individual over time, a technology especially useful for studying progressive retina diseases. Imaging technologies are currently under intense development to visualize structures in the living human eye; most notably, photoreceptors can be resolved by introducing adaptive optics into an ophthalmoscope (29). Furthermore, the combination of adaptive optics and TPM can increase both resolution and signal intensity (30). With continuing innovation, we envision that twophoton imaging of the retina will play a pivotal role in studying the retinoid cycle, both in basic and in clinical research.

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2. Materials Unless otherwise noted, all named reagents are available for purchase from either Fisher Scientific (Pittsburgh, PA) or SigmaAldrich (St. Louis, MO). 1. Balb/c mice (Jackson Laboratory, Bar Harbor, ME). Obtain IACUC approval for experimental use of mice. 2. Dissection tools: Spring scissors (catalog no. 15017-10 and 15004-08, Fine Science Tools Inc., Foster City, CA), tweezers (catalog no. 11252-30, Fine Science Tools Inc., Foster City, CA), no. 11 Surgical blades (Becton Dickinson, Franklin Lakes, NJ). 3. A glass-bottomed 35-mm dish (Mattek Corporation, Ashland, MA). 4. Ames medium: Powder of Ames medium (Sigma-Aldrich). Dissolve 8.8 g of powder in 1 l of distilled H2 O. Add 1.9 g of sodium bicarbonate to the medium. Equilibrate with 95% O2 and 5% CO2 . 5. All-trans-retinol: Dissolve all-trans-retinol in Ames medium containing 100 mM (2-hydroxypropyl)-βcyclodextrin with vigorous vortex mixing. 6. Paraformaldehyde (4%): Powder of paraformaldehyde (EM sciences, Hatfield, PA). Prepare 4% solution in 0.1 M phosphate buffer (100 mM sodium phosphate, pH 7.4). 7. PBS: 136 mM NaCl, 11.4 mM sodium phosphate, 0.1% Triton X-100, adjusted to pH 7.4. 8. PBST: PBS with 0.1% Triton X-100. 9. Blocking solution: PBST with 1.5% goat serum. 10. Antibodies: Primary antibody – Guinea pig anti-Adfp (Progen, Heidelberg, Germany). Secondary antibody – Cy2-conjugated donkey anti-guinea pig IgG (Jackson ImmunoResearch Laboratories, Inc., West Grove, PA). Dilute antibodies in PBST at concentrations recommended by the vendors. 11. No. 1 Safelight filter (Eastman Kodak, Rochester, NY). 12. Ketamine and xylazine mixture: Mix to final concentrations of 6 mg/ml ketamine (Bioniche Pharma Inc., Bogart, GA), 0.44 mg/ml xylazine, 10 mM phosphate buffer, and 100 mM NaCl in distilled H2 O. Adjust pH to 7.4. 13. Cyanoacrylate glue from Elmer’s Products Inc. (Columbus, OH). 14. Scotch tape (3 M Corporate, St. Paul, MN). 15. Circular cover glass (44-mm diameter, 0.16-mm thickness) (Carl Zeiss MicroImaging Inc., Thornwood, NY).

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16. A flash unit (Nikon, Melville, NY). 17. Human eyes, 8–12-h postmortem, obtained through Lions Eye Bank (Seattle, WA). Legal requirements for use of human donor retinas and primate retinas should be met (University of Washington Human Subjects, approval on file).

3. Methods 3.1. Configurations of Two-Photon Microscopes and Imaging Procedures

To obtain the data shown in Figs. 14.2 and 14.4, perform TPM with a Zeiss LSM 510 MP-NLO confocal microscope (Carl Zeiss, Germany) with LSM 510 software 3.0. Focus laser pulses of 730 nm from a mode-locked Ti:sapphire laser, Mira-900 (Coherent, Mountain View, CA) on the RPE through the sclera with a Plan-Neofluar 40×/1.3NA objective lens (Carl Zeiss). Alternatively, to obtain the data in Figs. 14.1B and 14.3, perform two-photon excitation microscopy with a Leica TCS SP2 scanning head (Leica) attached to a DM IRBE2 inverted microscope (Leica Microsystems Inc., Bannockburn, IL). Use LCS 3D Software (Leica, Germany) for data acquisition. Deliver laser pulses of 730 nm from a mode-locked Ti:sapphire laser, ChameleonTM -XR (Coherent, Mountain View, CA), through the microscope system to the RPE through the sclera with an HCX PL APO 40× oil immersion objective lens (NA= 1.25, Leica) (see Note 2). For both the microscope systems, maintain the reaction temperature at 36–37◦ C using a temperature-controlled microscope stage (Heating insert P and Tempcontrol 37-2, PeCon, Erbach, Germany). Maintain 40× objective lenses at 37◦ C with an objective heater (PeCon) or an ASI 400 air stream incubator (NEVTEK, Williamsville, VA) (see Note 3). Collect sample fluorescence (385–545 nm) through the objective lens, separated from the excitation light by a dichroic mirror, and filter fluorescence through custom-made filters (HQ 465/160, Chroma Technology Corp., Rockingham, VT) and direct it to a photomultiplier tube detector. Photomultiplier tube detectors should be installed in a non-descanned detection configuration, in which the fluorescence emission is directly delivered to the detectors without passing through the X–Y scanner (see Note 4). Measure the laser beam intensity at the back aperture of the objective lens and keep it at 3–30 mW for ex vivo studies and ∼5 mW for in vivo studies. The imaging resolution should be 0.22–0.24 μm/pixel for ex vivo studies (Figs. 14.1, 14.2, and 14.4) and 0.44 μm/pixel for in vivo studies (Fig. 14.3). Set the line scanning rate at 400 Hz.

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3.2. Two-Photon Imaging of RPE Cells in Mouse Eyeball Preparations

1. After euthanasia, dissect out a mouse eyeball with spring scissors and tweezers (see Note 5). 2. Locate the eyeball at the center of a 35-mm glass-bottomed dish with the sclera in direct contact with the coverslip at the bottom of the dish. Keep the eyeball hydrated with Ames medium. 3. Place the 35-mm dish on the microscope stage. 4. Collect images as described in Section 3.1.

3.3. Two-Photon Imaging of RPE in Mouse Eyecup Preparations

1. After euthanasia, dissect out a mouse eyeball with spring scissors and tweezers. Place the eyeball in a Petri dish and wash with Ames medium. 2. To remove the cornea, cut ∼0.5 mm below the corneal limbus with a number 11 surgical blade and spring scissors. Remove the lens and retina with fine tweezers and spring scissors. Dissect the eye in fresh Ames medium. After dissection, the RPE should be exposed to the Ames medium (see Notes 6 and 7). 3. Locate the eyecup at the center of a glass-bottomed 35-mm dish (MatTek Corporation, MA) so that the sclera is in direct contact with the coverslip at the bottom of the dish. 4. Apply a drop (20–50 μl) of fresh Ames medium onto the RPE. Prevent eyecup from floating by adjusting the volume of added fluid (see Note 8). 5. Image the RPE by TPM as explained in Section 3.1. 6. Remove Ames medium and apply a drop (20–50 μl) of 1.4 mM all-trans-retinol to the RPE. After 3 min, gently remove the solution and wash the eyecup with Ames medium. Collect an image by TPM as described in Section 3.1. 7. Obtain images from the RPE before and after all-transretinol treatment as shown in Fig. 14.2A.

3.4. Two-Photon and Confocal Imaging of the RPE to Discover Protein Components of Retinyl Ester Storage Structures

1. Isolate a mouse eyecup by following the procedures in Section 3.3. 2. Gently apply 1 ml of 4% paraformaldehyde onto the eyecup preparation. 3. Fix the tissue for 30 min. 4. Remove the paraformaldehyde and wash the eyecup with 1 ml of PBST. Wash three times for 5 min each. 5. Remove PBST and incubate the eyecup in 1 ml of blocking solution. Incubate at room temperature for 15 min. 6. Remove blocking solution and add 100 μl of primary antibody diluted in PBST solution. If a sufficient amount of primary antibody is available, increase the volume (see Note 9).

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7. Wash the eyecup with 1 ml of PBST three times for 5 min each. 8. Remove PBST and apply 500 μl of diluted secondary antibody. 9. Incubate for 30–45 min and then wash the eyecup with 1 ml of PBST three times for 5 min each. 10. Locate the eyecup at the center of a glass-bottomed 35-mm dish (MatTek Corporation, MA) so the sclera is in direct contact with the bottom coverslip. 11. Find the RPE cell layer by scanning the eye sample in XYZ directions by two-photon microscopy. 12. Collect an image of retinyl ester storage structures by following the procedure in Section 3.1. 13. Then collect an immunofluorescence image by using the filter, a dichroic mirror, and laser beam appropriate for the fluorescent molecule conjugated with the secondary antibody (see Note 10). 3.5. Two-Photon Imaging of RPE Cells in a Live Mouse

1. Maintain mice in the dark (>1 week) before the experiment (see Note 11). 2. Perform all the procedures under a safelight (>560 nm). 3. Anesthetize a mouse by intraperitoneal injection of the ketamine/xylazine mixture at a dose of 15 μl/g body weight (see Note 12). 4. Maintain the mouse at 36–37◦ C until the anesthesia becomes deep enough for the experiment. 5. Apply a small amount of cyanoacrylate glue between the eye and cover glass (see Fig. 14.3A) (see Note 13). 6. Lay the mouse on the microscope stage maintained at 36–37◦ C. Mount the cover glass on the stage and immobilize it by using scotch tape. 7. Find the RPE cell layer by scanning the eye sample in XYZ directions by two-photon microscopy (see Section 3.1). 8. Obtain an image of the RPE by two-photon microscopy as described in Section 3.1. 9. Expose the eye to intense light flashes to bleach ∼60% of the visual pigment. 10. Image the RPE cells every 1–10 min as described in Section 3.1.

3.6. Two-Photon Imaging of the RPE from a Human Eye

1. Fix a human eyeball in 20 ml of 4% paraformaldehyde for 10 min (see Note 14). 2. Place the eyeball into a Petri dish with PBS.

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3. Remove the lens, cornea, vitreous, and retina from the eye to prepare an eyecup (see Note 15). 4. Fix the eyecup in 20 ml of 4% paraformaldehyde for 1 h. 5. Wash the eyecup with 20 ml of PBS three times for 5 min each. 6. With the eyecup, peel the RPE along with the choroid off the sclera. 7. Cut the RPE/choroid layer into 1 × 1 cm squares; obtain tissue pieces from regions 1–4 as shown in Fig. 14.4A. RPE cells from the macular region are readily discerned because they are more deeply pigmented than cells within the peripheral regions. 8. Place the samples on a 35-mm glass-bottomed dish so that the RPE cells directly contact the surface of the glass and the choroid faces toward the top. 9. Apply two-photon imaging as described in Section 3.1. 10. For quantification of fluorescence, use the “measure” function in ImageJ software. Normalized pixel values for different areas of the retina are compared in Fig. 14.4C. Pixel values should be averaged for entire image fields. Intensities in the nucleus should be used as background and subtracted from measured values.

4. Notes 1. Two-photon microscopy, used throughout this manuscript, is a broad term that includes two-photon excitation laser scanning microscopy and second harmonic imaging microscopy. 2. When using an oil or a water immersion objective lens on an inverted microscope, the diameter of the immersion fluid is a good indicator for estimating the distance between the objective lens and the bottom of the dish. When starting each imaging session, adjust the location of the objective lens as close as possible to the bottom of the dish. 3. The objective lens is a large heat sink. So when used, it needs to be warmed to the physiological temperature of an animal or tissue. 4. Non-descanned detection is highly sensitive and affected by light from computer monitors and other sources. Therefore, we recommend turning off room lighting. Also, do not use the mercury lamp and halogen lamp of the microscope system. We found that the noise level can be reduced

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by covering the computer monitors with red filter sheets (E-color, Rosco Laboratories Inc., Stamford, CT). 5. Pigmented mice are not well suited for imaging through the sclera so albino Balb/c mice were primarily used for our studies. Another commonly used strain, the C57BL/6, tyrosinase mutant mouse (Tyrc-2J ), is not pigmented and is available from Jackson Laboratories (Bar Harbor, ME). 6. In Section 3.3, we found it difficult to penetrate RPE cells with all-trans-retinol if the retina is attached to the RPE. Similar difficulties were encountered in applying several reagents, including FM4-64, BODIPY FL C5 -ceramide, MitoTracker Orange CMTMRos, LysoTracker Green DND-26, DiOC6 , and Nile Red (Invitrogen Corporation, Carlsbad, CA). Thus, it is essential to remove the retina to test the effect of small test molecules on the RPE cell layer. 7. In Sections 3.3 and 3.4, it is easier to access the retina if an eye is cut ∼0.5 mm below the corneal limbus to remove the cornea first. 8. In Sections 3.3 and 3.4, the RPE layer an eyecup preparation is fragile so all solutions should be applied gently. 9. Before proceeding to the procedure in Section 3.4, confirm that the primary antibody works on cryosections of mouse eye when viewed by the same detection method (e.g., Alexa 488 or Cy3-labeled secondary antibodies). 10. In Section 3.4, misalignment of confocal and TPM lasers will result in a shift of two channels of the merged images. Ask the microscope vendor or laser specialist to align the lasers precisely. 11. In Section 3.5, this in vivo imaging method is applicable only to the peripheral part of the retina, because the central part of the eye is embedded in the orbital cavity and inaccessible. In the peripheral part of the retina (Fig. 14.3), we recognize that retinyl ester storage structures are less organized than those in the central retina (Fig. 14.1B). Disorganized fluorescent structures are found at the edge of the human retina as well (Fig. 14.4B). 12. In Section 3.5, individual differences were observed in the susceptibility of mice to the anesthesia. Monitor the condition of these animals carefully during the imaging procedure. 13. In Section 3.5, we noticed that the eyes are embedded deeper in the orbital cavity of older mice. For in vivo imaging, the RPE is optically more accessible in young mice (3–4 weeks) than in older mice (>months).

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14. In Section 3.6, human eyes are a potential source of blood-borne pathogens. Comply with OSHA regulations for handling potential sources of blood-borne pathogens. Upon receipt of these eyes, place them immediately in 4% paraformaldehyde. 15. In Section 3.6, to facilitate removal of the human retina, remove the cornea and lens by cutting 2–3 mm below the corneal limbus.

Acknowledgments This research was supported in part by grants EY009339 and P30 EY11373 from the National Institutes of Health and the Foundation Fighting Blindness. References 1. Palczewski, K. (2006) G protein-coupled receptor rhodopsin. Annu. Rev. Biochem. 75, 743–767. 2. McBee, J.K., Palczewski, K., Baehr, W., Pepperberg, D.R. (2001) Confronting complexity: The interlink of phototransduction and retinoid metabolism in the vertebrate retina. Prog. Retin. Eye Res. 20, 469–529. 3. Lamb, T.D., Pugh, E.N., Jr. (2004) Dark adaptation and the retinoid cycle of vision. Prog. Retin. Eye Res. 23, 307–380. 4. Thompson, D.A., Gal, A. (2003) Vitamin A metabolism in the retinal pigment epithelium: Genes, mutations, and diseases. Prog. Retin. Eye Res. 22, 683–703. 5. Travis, G.H., Golczak, M., Moise, A.R., Palczewski, K. (2007) Diseases caused by defects in the visual cycle: Retinoids as potential therapeutic agents. Annu. Rev. Pharmacol. Toxicol. 47, 469–512. 6. Moiseyev, G., Chen, Y., Takahashi, Y., Wu, B.X., Ma, J.X. (2005) RPE65 is the isomerohydrolase in the retinoid visual cycle. Proc. Natl. Acad. Sci. USA 102, 12413–12418. 7. Jin, M., Li, S., Moghrabi, W.N., Sun, H., Travis, G.H. (2005) Rpe65 is the retinoid isomerase in bovine retinal pigment epithelium. Cell 122, 449–459. 8. Redmond, T.M., Poliakov, E., Yu, S., Tsai, J.Y., Lu, Z., Gentleman, S. (2005) Mutation of key residues of RPE65 abolishes its enzymatic role as isomerohydrolase in the visual cycle. Proc. Natl. Acad. Sci. USA 102, 13658–13663.

9. Yamamoto, H., Simon, A., Eriksson, U., Harris, E., Berson, E.L., Dryja, T.P. (1999) Mutations in the gene encoding 11-cis retinol dehydrogenase cause delayed dark adaptation and fundus albipunctatus. Nat. Genet. 22, 188–191. 10. Driessen, C.A., Winkens, H.J., Hoffmann, K., Kuhlmann, L.D., Janssen, B.P., Van Vugt, A.H., Van Hooser, J.P., Wieringa, B.E., Deutman, A.F., Palczewski, K., Ruether, K., Janssen, J.J. (2000) Disruption of the 11cis-retinol dehydrogenase gene leads to accumulation of cis-retinols and cis-retinyl esters. Mol. Cell. Biol. 20, 4275–4287. 11. Kim, T.S., Maeda, A., Maeda, T., Heinlein, C., Kedishvili, N., Palczewski, K., Nelson, P.S. (2005) Delayed dark adaptation in 11-cis-retinol dehydrogenase-deficient mice: A role of RDH11 in visual processes in vivo. J. Biol. Chem. 280, 8694–8704. 12. Saari, J.C., Garwin, G.G., Haeseleer, F., Jang, G.F., Palczewski, K. (2000) Phase partition and high-performance liquid chromatography assays of retinoid dehydrogenases. Methods Enzymol. 316, 359–371. 13. Denk, W., Strickler, J.H., Webb, W.W. (1990) Two-photon laser scanning fluorescence microscopy. Science 248, 73–76. 14. Denk, W., Svoboda, K. (1997) Photon upmanship: Why multiphoton imaging is more than a gimmick. Neuron 18, 351–357. 15. Svoboda, K., Yasuda, R. (2006) Principles of two-photon excitation microscopy and its applications to neuroscience. Neuron 50, 823–839.

Two-Photon Microscopy in Eye Research 16. Imanishi, Y., Lodowski, K.H., Koutalos, Y. (2007) Two-photon microscopy: Shedding light on the chemistry of vision. Biochemistry 46, 9674–9684. 17. Williams, R.M., Piston, D.W., Webb, W.W. (1994) Two-photon molecular excitation provides intrinsic 3-dimensional resolution for laser-based microscopy and microphotochemistry. FASEB J. 8, 804–813. 18. Diaspro, A. (ed.). (2002) Confocal and TwoPhoton Microscopy: Foundations, Applications, and Advances, Wiley-Liss, New York, NY. 19. Imanishi, Y., Batten, M.L., Piston, D.W., Baehr, W., Palczewski, K. (2004) Noninvasive two-photon imaging reveals retinyl ester storage structures in the eye. J. Cell. Biol. 164, 373–383. 20. Jiang, H.P., Serrero, G. (1992) Isolation and characterization of a full-length cDNA coding for an adipose differentiation-related protein. Proc. Natl. Acad. Sci. USA 89, 7856–7860. 21. Kaplan, M.W. (1985) Distribution and axial diffusion of retinol in bleached rod outer segments of frogs (Rana pipiens). Exp. Eye Res. 40, 721–729. 22. Cornwall, M.C., Tsina, E., Crouch, R.K., Wiggert, B., Chen, C., Koutalos, Y. (2003) Regulation of the visual cycle: Retinol dehydrogenase and retinol fluorescence measurements in vertebrate retina. Adv. Exp. Med. Biol. 533, 353–360. 23. Chen, C., Tsina, E., Cornwall, M.C., Crouch, R.K., Vijayaraghavan, S., Koutalos, Y. (2005) Reduction of all-trans retinal to alltrans retinol in the outer segments of frog

24.

25.

26. 27.

28.

29. 30.

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and mouse rod photoreceptors. Biophys. J. 88, 2278–2287. Wu, Q., Chen, C., Koutalos, Y. (2006) Alltrans retinol in rod photoreceptor outer segments moves unrestrictedly by passive diffusion. Biophys. J. 91, 4678–4689. Imanishi, Y., Sun, W., Maeda, T., Maeda, A., Palczewski, K. (2008) Retinyl ester homeostasis in the adipose differentiation-related protein-deficient retina. J. Biol. Chem. 283, 25091–25102. Rodieck, R.W. (1998) The First Steps in Seeing, Sinauer Associates, Inc., Sunderland, MA. Jacobson, S.G., Aleman, T.S., Cideciyan, A.V., Heon, E., Golczak, M., Beltran, W.A., Sumaroka, A., Schwartz, S.B., Roman, A.J., Windsor, E.A., Wilson, J.M., Aguirre, G.D., Stone, E.M., Palczewski, K. (2007) Human cone photoreceptor dependence on RPE65 isomerase. Proc. Natl. Acad. Sci. USA 104, 15123–15128. Maeda, A., Maeda, T., Imanishi, Y., Golczak, M., Moise, A.R., Palczewski, K. (2006) Aberrant metabolites in mouse models of congenital blinding diseases: Formation and storage of retinyl esters. Biochemistry 45, 4210–4219. Roorda, A., Williams, D.R. (1999) The arrangement of the three cone classes in the living human eye. Nature 397, 520–522. Rueckel, M., Mack-Bucher, J.A., Denk, W. (2006) Adaptive wavefront correction in two-photon microscopy using coherencegated wavefront sensing. Proc. Natl. Acad. Sci. USA 103, 17137–17142.

Chapter 15 Reverse-Phase High-Performance Liquid Chromatography (HPLC) Analysis of Retinol and Retinyl Esters in Mouse Serum and Tissues Youn-Kyung Kim and Loredana Quadro Abstract The ability to measure endogenous metabolites of retinoids (vitamin A and its derivatives) in biological samples is key to understanding the crucial physiological actions of vitamin A. Over the years, many assays and methods have been developed to analyze different retinoids in biological samples. Liquid chromatography is the best analytical technique for routine analysis of these compounds. However, due to their different chemical properties, different retinoid metabolites cannot be accurately separated and quantified in a single chromatographic run. Here, we will describe a reverse-phase HPLC isocratic method that enables extraction, separation, identification, and quantification of all-trans-retinol and different molecular species of retinyl ester with high accuracy, sensitivity, and reliability. Key words: Reverse-phase HPLC, retinol, retinyl ester, quantification, separation, mouse, tissues, retinoids, vitamin A.

1. Introduction Vitamin A is a lipid-soluble hormone that regulates the transcription of a number of genes that are crucial for many important biological functions (1). Adequate levels of retinoids (vitamin A and its derivatives) in serum and tissues are essential to maintain the health of the body (2). Retinoid homeostasis is achieved through a series of complex mechanisms that regulate absorption, storage, transport, and metabolism of this nutrient. Mammals obtain all vitamin A and its derivatives from the diet as preformed dietary vitamin A (retinyl esters, retinol, and very H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_15, © Springer Science+Business Media, LLC 2010

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small amounts of retinoic acid) from animal products or as βcarotene from vegetables and fruits (3). Within the intestinal mucosa, all retinol, regardless of its dietary origin, is enzymatically re-esterified with long-chain fatty acids and, together with other dietary lipids, packaged into chylomicrons and secreted into the lymphatic system (4). Once in the general circulation, chylomicrons undergo lipolysis of the triglycerides giving rise to free fatty acids and smaller lipoprotein particles called chylomicron remnants, still retaining retinyl ester (5, 6). Approximately 75% of retinoids within chylomicron remnants are cleared by the liver, the major site of vitamin A storage and metabolism, while the remaining can be taken up by extrahepatic tissues (7–9). To meet tissue retinoid needs, the liver secretes retinol into the circulation, bound to its sole-specific transport protein retinol-binding protein (RBP; also known as RBP4) (10, 11). Upon recognition of the serum retinol–RBP complex by Stra6, its recently identified specific membrane receptor (12), target tissues acquire retinol which can be subsequently oxidized to retinoic acid, the active form of vitamin A (4). Retinoic acid acts as a ligand for specific nuclear receptors that, in turn, control gene transcription (1). The levels of circulating retinol and retinyl ester reflect the whole-body vitamin A status, which is determined by both the concentration of retinoids within the stores and the recent dietary retinoid intake. Therefore, the ability to measure endogenous retinoid levels in serum and tissues is pivotal to elucidate the regulatory mechanisms that maintain retinoid homeostasis and, ultimately, to overcome many pathological conditions and diseases that have been associated with alterations in retinoid metabolism (13–19). For routine assessment and characterization of retinoids in biological samples, liquid chromatography is the best analytical technique. The different chemical properties of the retinoid metabolites do not allow accurate quantification of retinol, its isomers, retinal, retinyl esters, and retinoic acid in a single chromatographic run (20). In addition, the levels of retinoic acid in biological samples are extremely low, thus requiring sophisticated methods for their accurate quantification (21, 22). Herein, we will describe the extraction, separation, identification, and quantification of all-trans retinol and retinyl esters in murine serum and tissues. The mouse is the most commonly used experimental animal model to study whole-body vitamin A metabolism. However, this method can also be used to measure retinoid concentrations in cell culture system or in human samples. To date, many assays have been developed to analyze retinol and retinyl ester levels in biological samples (23–28). Some of the previously described methods focus on separation of the different molecular species, but either do not offer a rigorous quantification or do not have high sensitivity. Others provide precise identification and robust quantification of several retinoid

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compounds, but do not allow separation of the different molecular species of retinyl ester (20). In this chapter, we describe a reverse-phase high-performance liquid chromatography (HPLC) isocratic method that allows the accurate extraction, separation, identification, and quantification of all-trans-retinol and different molecular species of retinyl ester with a high analytical performance. The sample requirement (from 50 to 200 mg tissue and 80–100 μl serum), the high recovery during extraction (ranging from 75 to 95%), the lower limits of detection (defined as signal/noise ratio of 3:1) of 0.35 and 0.95 pmol for retinol and retinyl palmitate, respectively, and the relatively short run time (35 min) make this method comparable to others recently reported in detail (20). In addition, this method is also suitable for simultaneous analysis and quantification of retinoid and carotenoids from biological samples (29). This methodology, routinely used in our laboratory (30), was originally established by Blaner and colleagues (29).

2. Materials 2.1. Preparation of the Standards

1. Ethanol (ACS grade). 2. Retinol (Sigma). 3. Retinyl acetate (Sigma). 4. Retinyl palmitate (Sigma). 5. Amber vials with cap.

2.2. Retinoid Extraction

1. Fresh or frozen serum or tissues (see Note 1). 2. Internal standard (retinyl acetate). 3. Ethanol (ACS grade). 4. Hexane (HPLC grade). 5. H2 O (HPLC grade). 6. PVDF filter membrane (0.22 μm, 47 mm). 7. All-glass filtration unit. 8. Phosphate buffer saline (PBS), for tissues only. 9. N2 gas, Evap-O-Rac System (Cole-Parmer) . 10. PRO200 Homogenizer (PROscientific), for tissues only. 11. Glass Pasteur pipettes (9 in.) and rubber bulbs. 12. Polypropylene tubes (12 mm × 75 mm). 13. Glass test tubes (13 mm × 100 mm; 16 mm × 100 mm).

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2.3. Retinoid Analysis

1. High-performance liquid chromatography system (see Note 2). 2. Column (see Section 3.4.2, Beckman, Part. no. 235329; see Note 3). 3. Guard column (PerkinElmer, Part. no. 0711-0092). 4. Amber vials (National scientific, cat. no. C4000-2 W; see Note 2). 5. Capacity glass vial inserts (300 μl; National scientific, cat. no. C4010-630; see Note 4). 6. Vial caps with PTFE/Silicon septa (National scientific, cat. no. C4000-54A; see Note 5). 7. Methanol (HPLC grade). 8. Acetonitrile (HPLC grade). 9. Methylenechloride (HPLC grade).

3. Methods Due to the light-sensitive nature of retinoids, all the following experimental procedures should be performed “in the dark” (see Note 6). 3.1. Preparation of the Standards

1. Prepare stock solutions of standards by dissolving each standard into the appropriate solvent as follows: ethanol for retinol and retinyl acetate; hexane for retinyl palmitate. 2. Stock standard solutions should be prepared in amber vials and kept at −20◦ C (see Note 7). 3. Dilute each standard solution up to approximately 1 ng/μl (see Note 8). 4. Measure the absorbance of the diluted standard solutions by spectrophotometer at 325 nm (see Note 9). 5. Calculate the concentration of each standard solution based on the O.D. and the specific extinction coefficient (see Note 10). Concentration (ng/μL) =

OD standard solution × 106 Extinction coefficient × 100

6. Aliquot the diluted standard solutions in small amber vials and keep them at −20◦ C (see Note 8).

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3.2. Determination of the Calibration Curves 3.2.1. Detection Limits

1. Prepare a series of dilutions with different amounts for each standard (retinol, retinyl acetate, and retinyl palmitate) (see Note 11). 2. Inject the dilutions into the HPLC column (see Note 12). 3. Integrate the peak signals detected by UV absorbance at 325 nm and obtain the peak areas. Generate a standard curve by plotting the amount of each standard dilution on the x-axis and the corresponding peak area on the y-axis (Fig. 15.1a).

Fig. 15.1. Examples of limit of detection and standard curve. (a) Representative calibration curve for all-trans-retinol (at-ROL). On the x-axis, retinol mass is expressed in picomoles. On the y-axis the peak area is expressed as absorbance units (mAU). (b) Representative standard curve for retinol:retinyl acetate. On the x-axis are the molar ratios of all-trans-retinol (at-ROL) and retinyl acetate (Rac). On the y-axis the peak area is expressed as absorbance units (mAU). All r2 values are greater than 0.99.

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3.2.2. Standard Curves

1. Prepare two series of standard solutions with different molar ratio of retinol–retinyl acetate and retinyl palmitate–retinyl acetate in HPLC amber vials as follows (see Note 13): Retinol:retinyl acetate (m:m) = 0.1:1, 0.25:1, 0.5:1, 1:1, 2:1 (retinyl acetate concentration should be approximately 1 ng/μl) Retinyl palmitate:retinyl acetate (m:m) = 0.1:1, 0.25:1, 0.5:1, 1:1, 2:1, 3:1, 4:1, 5:1 (retinyl acetate concentration should be approximately 2 ng/μl) 2. Inject the different standard solutions on the HPLC column (see Note 12). 3. Integrate the peak signals detected by UV absorbance at 325 nm and obtain the peak areas. Generate a standard curve by plotting the molar ratio between the retinoid compound of interest and the internal standard on the x-axis and the corresponding peak area on the y-axis (Fig. 15.1b).

3.3. Retinoid Extraction 3.3.1. Serum Extraction (See Note 6)

1. Add 100 μl of serum into a glass test tube. 2. Add 25 μl of the internal standard retinyl acetate and add ethanol so that the ratio between the total volume of ethanol and the total volume of serum used for the extraction is 1:1 (for example, if 150 μl of serum is used, 25 μl of the internal standard and 125 μl of ethanol will be added) (see Note 14). 3. Vortex the tube briefly. 4. Add 4 ml of hexane (see Note 15) and vortex for 30 s two times (see Note 16). 5. Centrifuge at 3,000 rpm for 3 min in a tabletop low-speed centrifuge (see Note 17). 6. By using a glass Pasteur pipette, transfer the upper phase into a new glass test tube containing 500 μl of H2 O. 7. Vortex the new tube briefly. 8. Repeat step 5 and transfer the supernatant into a new glass test tube with a glass Pasteur pipette (see Note 18). 9. Dry the supernatant under a gentle stream of N2 , by using the Evap-O-Rac System (Cole-Parmer). 10. Dissolve the sample in 50 μl of mobile phase (see Note 19) and transfer into a vial for injection on the HPLC column.

3.3.2. Tissue Extraction (See Note 6)

1. The weight of the tissue for extraction will depend on its retinoid content. For the purpose of explaining the

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procedure, we will describe retinoid extraction from liver, for which we recommend to use 100 mg (see Note 20). 2. Place 100 mg of liver into a polypropylene tube containing 2 ml of PBS (see Note 21). 3. Homogenize at medium speed for 10 s (see Note 22). 4. Transfer 200 μl of the homogenate into a glass test tube (see Note 23). 5. Add 100 μl of internal standard retinyl acetate and 100 μl of ethanol (see Note 14). 6. Vortex the tube briefly. 7. Follow the procedure described for serum extraction from step 4 (see Note 24). 3.4. High-Performance Liquid Chromatography (HPLC) Analysis (See Note 25) 3.4.1. Preparation of the Mobile Phase

Prepare the mobile phase according to the protocol below (see Note 26): Acetonitrile

70%

Methanol

15%

Methylenechloride

15%

3.4.2. Chromatography Conditions

3.4.3. Determination of the Retinoid Concentration

Column

Beckman Ultrasphere C18 (5 μm), 4.6 mm × 250 mm

Guard column

C18 (7 μm), 15 mm × 3.2 mm

Flow rate

1.8 ml/min

Run time

35 min

Injection volume

20 μl

PDA detection wavelength

325 nm

Integrate the peak signals detected by UV absorbance at 325 nm and obtain the peak areas for each of the different retinoids separated and identified upon the chromatographic run. The mass of each retinoid compound present under its HPLC peak will be determined from the area under the peak, using the equation of the standard curve generated as described above (see Note 27). A typical liver retinoid HPLC chromatogram is shown in Fig. 15.2.

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Fig. 15.2. Representative chromatogram from reverse-phase HPLC analysis of retinol and retinyl esters in mouse liver. Retinoids were detected by UV absorbance at 325 nm and retinol and retinyl esters (retinyl palmitate, oleate, linoleate, and stearate) were identified by comparing peak integrated areas for unknowns against those of known amounts of purified standards, according to previous reports (29). On the x-axis, the retention time is expressed as minutes. On the y-axis the peak area is expressed as absorbance units (mAU). Peak 1, retinol; peak 2, internal standard retinyl acetate; peak 3, retinyl linoleate; peak 4, retinyl oleate; peak 5, retinyl palmitate; peak 6, retinyl stearate.

4. Notes 1. All samples will be flash-frozen in liquid N2 immediately after dissection and stored at −80◦ C until analysis will be performed. If possible, dissection should be carried “in the dark” (see Note 6) to minimize losses of light-sensitive retinoids. 2. All the HPLC accessories such as amber vials, vial inserts, and caps described in this chapter are compatible with the Dionex Ultimate 3000 series HPLC instrument. Different HPLC systems might require other types of accessories. 3. In our experience, this Beckman column has shown reproducible results (consistent retention times) over time. 4. These glass inserts are held into the vial by a spring that provides a cushion against needle contact. Once the sample analysis is completed, we recommend saving the spring for further assembly of new vials. 5. These vial caps and septa can also be purchased separately (cap w/o septa, National Scientific, cat. no. C400098BLK; septa, National Scientific, cat. no. C4000-60). 6. The extraction of retinoids from serum and tissues must be carried out rapidly “in the dark,” and all the extraction

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steps should be performed on ice or in a cold room. Laboratory windows should be covered with appropriate materials such as aluminum foil or heavy curtains (a room with no windows is the ideal setting to perform this procedure). Artificial lighting should be provided by yellow light bulbs sold in retail stores. Alternatively, use dim light and never expose the samples to direct illumination. 7. Highly concentrated retinoid standard stock solutions may degrade over time, even if stored at −20◦ C. The optimal recommended concentration of stock solutions is approximately 1 mg/ml. 30 mL is the suggested volume for a stock solution. Standard solutions prepared as indicated can be kept for several months at −20◦ C. 8. Diluted standard solutions are kept in amber vials. We recommend preparing small aliquots of 3–4 ml of the diluted standard solutions. Degradation or losses of the compounds can be minimized by flushing the headspace of the vial with N2 gas every time before closing the cap. It is recommended to analyze each aliquot of the diluted standards by HPLC before use, to check its quality. 9. Use 1-cm-width quartz cuvette (1 ml). 10. The extinction coefficient (E11%cm ) depends upon the compound and the solvent in which it is dissolved. The extinction coefficient for retinyl acetate dissolved in ethanol is 1550, for retinol dissolved in ethanol is 1835, and for retinyl palmitate dissolved in ethanol is 975. 11. The detection limit depends upon the detector and the column used. The range of concentrations tested should be chosen based on the retinoids content of the tissue analyzed. See Fig. 15.1a for a typical example of a detection limit curve. 12. How to operate the HPLC system will depend on the type of instrument and the description of this procedure does not pertain to this work. We only recommend monitoring the column pressure and the baseline of the target wavelength prior to loading the samples. Each dilution is run on the HPLC in triplicate. The injection volume varies according to the HPLC system and/or the protocol used. Twenty microliter is the standard injection volume for this protocol. 13. After adding the appropriate volume (according to the molar ratio) of the different retinoid compounds into a glass test tube, dry out each solution under a gentle stream of N2 gas and re-suspended in mobile phase. Vortex well and transfer into the insert of the HPLC amber vial immediately.

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14. The suggested concentration of internal standard is 1 ng/μl. Note that the internal standard is dissolved in ethanol, and therefore its volume should be taken into account when calculating the total ethanol volume required to perform the extraction. 15. Once hexane is added, retinoids are stable. In other words, extraction procedures must be performed rapidly until the addition of hexane. 16. During this step, it is recommended to increase the speed of the vortex slowly so as to avoid spill over of solvent. One should also hold the glass tube from the side. Placing a finger on top of the tube may cause impurities to contaminate the sample, should the solvent overflows while mixing. 17. After this centrifugation step, the sample will consist of two layers: the bottom layer is the aqueous phase and it is slightly cloudy; the top, clear layer is the solvent phase containing retinoids. At the interface between the two layers and/or at the bottom of the tube, white or pinky colored tissue residues can be present. 18. After this centrifugation step, the sample will consist of two layers: the bottom layer contains water and the top layer contains the solvent. Both layers are clear and no residues are visible. Carefully transfer only the upper layer without touching the lower layer. 19. This step should be performed rapidly to avoid sample evaporation. Assemble the HPLC vials (insert vials, cap with septa, etc) ahead. In addition to serum, most tissues can be easily re-suspended in mobile phase. Should this not be the case, the sample appears cloudy and should not be injected on the HPLC column. For example, adipose tissue should be re-suspended in an alternative solvent, such as benzene, mobile phase:benzene (3:2, v:v), acetonitrile. 20. For adipose, we recommend to perform the extraction with less than 50 mg of tissue. For embryos at 14.5 dpc (approximately 200 mg), we suggest using the whole embryo and for adult prostate tissue the whole organ (about 40– 50 mg). 21. Different volumes of PBS may be chosen to homogenize different tissues due to their different retinoid content. For example, 1 ml of PBS is recommended to homogenize a 14.5 dpc embryo, 2 ml of PBS for 50 mg of adipose tissue, and 1 ml of PBS for an adult prostate. 22. To avoid contaminations during the homogenization step, remember to wash the probe carefully with clean PBS between each samples.

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23. Liver is a tissue with a high concentration of retinoids. Therefore, we recommend to perform the extraction with only one-tenth of the homogenate. However, the optimal homogenate volume for the extraction may vary, depending on the retinoid content of the tissue. For example, in the case of retinoid extraction from embryo, we recommend performing the extraction with the entire (or onehalf) volume of the homogenate (1 ml). We also suggest using the whole volume of homogenate to perform the extraction from adipose (2 ml) or from prostate (1 ml). 24. It is not recommended to perform retinoid extraction from tissue homogenates previously stored at −20◦ C, as degradation of retinoids may occur. Freshly prepared tissue homogenates are preferred. 25. When assembling the samples for injection on the HPLC column, we recommended inserting a blank sample (mobile phase only) every 5–6 samples to clean the column from potential impurities. 26. Filter the mobile phase with the glass filter unit under vacuum by using the PVDF filter membrane. This step helps proper mixing of the different solvents and removes potential impurities. Furthermore, the mobile phase should be placed in an ultrasonicator for 30 min to degas it. 27. To obtain the peak areas, integration of the peak signals can be performed automatically through the HPLC system software or manually. Loss during extraction is accounted for by the addition of a known amount of internal standard (retinyl acetate) to the sample prior to extraction. The standard curve generated with retinyl palmitate and retinyl acetate (m/m) will be used to calculate the concentration of the different molecular species of retinyl ester. To obtain the final retinoid concentration, the volume of serum extracted or the percent of homogenate volume extracted vs. the total homogenate volume will also be taken into account.

References 1. Balmer, J.E., Blomhoff, R. (2002) Gene expression regulation by retinoic acid. J. Lipid Res. 43, 1773–1808. 2. Blomhoff, R., Blomhoff, H.K. (2006) Overview of retinoid metabolism and function. J. Neurobiol. 66, 606–630. 3. Sporn, M.B., Roberts, A.B., Goodman, D.S. (1994) The Retinoids, Biology, Chemistry, and Medicine, 2nd ed., Raven Press, New York.

4. Vogel, S., Gamble, M.V., Blaner, W.S. (1999) Biosynthesis, absorption, metabolism and transport of retinoids. In: Nau, H., Blaner, W.S. (eds.), Handbook of Experimental Pharmacology, Retinoids, the Biochemical And Molecular Basis of Vitamin A and Retinoid Action, Springer Verlag Publishing, Heidelberg, Germany, pp. 31–95.

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5. Olivecrona, T., Bengtsson-Olivecrona, G. (1993) Lipoprotein lipase and hepatic lipase. Curr. Opin. Lipidol. 4, 187–196. 6. Goldberg, I.J. (1996) Lipoprotein lipase and lipolysis: Central roles in lipoprotein metabolism and atherogenesis. J. Lipid Res. 37, 693–707. 7. Goodman, D.S., Huang, H.S., Shiratori, T. (1965) Tissue distribution of newly absorbed vitamin A in the rat. J. Lipid Res. 6, 390–396. 8. Cooper, A.D. (1997) Hepatic uptake of chylomicron remnants. J. Lipid Res. 38, 2173–2192. 9. Blaner, W.S., Olson, J.A. (1994) Retinol and retinoic acid metabolism. In: Sporn, M.B., Roberts, A.B., Goodman, D.S. (eds.), The Retinoids, Biology, Chemistry and Medicine, Raven Press, New York, pp. 229–256. 10. Soprano, D.R., Blaner, W.S. (1994) Plasma retinol-binding protein. In: Sporn, M.B., Roberts, A.B., Goodman, D.S. (eds.), The Retinoids, Biology, Chemistry and Medicine, Raven Press, New York, pp. 257–282. 11. Quadro, L., Hamberger, L., Colantuoni, V., Gottesman, M.E., Blaner, W.S. (2003) Understanding the physiological role of retinol-binding protein in vitamin A metabolism using transgenic and knockout mouse models. Mol. Aspect Med. 24, 421–430. 12. Kawaguchi, R., Yu, J., Honda, J., Hu, J., Whitelegge, J., Ping, P., Wiita, P., Bok, D., Sun, H. (2007) A membrane receptor for retinol binding protein mediates cellular uptake of vitamin A. Science 315, 820–825. 13. Yang, Q., Graham, T.E., Mody, N., Preitner, F., Peroni, O.D., Zabolotny, J.M., Kotani, K., Quadro, L., Kahn, B.B. (2005) Serum retinol binding protein 4 contributes to insulin resistance in obesity and type 2 diabetes. Nature 436, 356–362. 14. Ziouzenkova, O., Orasanu, G., Sharlach, M., Akiyama, T.E., Berger, J.P., Viereck, J., Hamilton, J.A., Tang, G., Dolnikowski, G.G., Vogel, S., Duester, G., Plutzky, J. (2007) Retinaldehyde represses adipogenesis and diet-induced obesity. Nat. Med. 13, 695–702. 15. Fields, A.L., Soprano, D.R., Soprano, K.J. (2007) Retinoids in biological control and cancer. J. Cell Biochem. 102, 886–898. 16. Goodman, A.B. (2006) Retinoid receptors, transporters, and metabolizers as therapeutic targets in late onset Alzheimer disease. J. Cell Physiol. 209, 598–603. 17. Golzio, C., Martinovic-Bouriel, J., Thomas, S., Mougou-Zrelli, S., GrattaglianoBessieres, B., Bonniere, M., Delahaye, S., Munnich, A., Encha-Razavi, F., Lyonnet,

18.

19.

20.

21.

22.

23. 24.

25.

26. 27.

S., Vekemans, M., Attie-Bitach, T., Etchevers, H.C. (2007) Matthew-Wood syndrome is caused by truncating mutations in the retinol-binding protein receptor gene STRA6. Am. J. Hum. Genet. 80, 1179–1187. Pasutto, F., Sticht, H., Hammersen, G., Gillessen-Kaesbach, G., Fitzpatrick, D.R., Nürnberg, G., Brasch, F., SchirmerZimmermann, H., Tolmie, J.L., Chitayat, D., Houge, G., Fernández-Martínez, L., Keating, S., Mortier, G., Hennekam, R.C., von der Wense, A., Slavotinek, A., Meinecke, P., Bitoun, P., Becker, C., Nürnberg, P., Reis, A., Rauch, A. (2007) Mutations in STRA6 cause a broad spectrum of malformations including anophthalmia, congenital heart defects, diaphragmatic hernia, alveolar capillary dysplasia, lung hypoplasia, and mental retardation. Am. J. Hum. Genet. 80, 550–560. Clagett-Dame, M., DeLuca, H.F. (2002) The role of vitamin A in mammalian reproduction and embryonic development. Annu. Rev. Nutr. 22, 347–381. Kane, M.A., Folias, A.E., Napoli, J.L. (2008) HPLC/UV quantitation of retinal, retinol, and retinyl esters in serum and tissues. Anal. Biochem. 378, 71–79. Kane, M. A, Chen, N., Sparks, S., Napoli, J.L. (2005) Quantification of endogenous retinoic acid in limited biological samples by LC/MS/MS. Biochem. J. 388, 363–369. Kane, M.A., Folias, A.E., Wang, C., Napoli, J.L. (2008) Quantitative profiling of endogenous retinoic acid in vivo and in vitro by tandem mass spectrometry. Anal. Chem. 80, 1702–1708. Packer, L. (1990) Retinoids: Part A – molecular and metabolic aspects. Methods Enzymol. 189, 3–583. Roberts, A.B., Nichols, M.D., Frolik, C.A., Newton, D.L., Sporn, M.B. (1978) Assay of retinoids in biological samples by reversephase high-pressure liquid chromatography. Cancer Res. 38, 3327–3332. Blaner, W.S., Hendriks, H.F., Brouwer, A., de Leeuw, A.M., Knook, D.L., Goodman, D.S. (1985) Retinoids, retinoid-binding proteins, and retinyl palmitate hydrolase distributions in different types of rat liver cells. J. Lipid Res. 26, 1241–1251. Napoli, J.L., Horst, R.L. (1998) Quantitative analyses of naturally occurring retinoids. Methods Mol. Biol. 89, 29–40. Harrison, E.H., Blaner, W.S., Goodman, D.S., Ross, A.C. (1987) Subcellular localization of retinoids, retinoid-binding proteins, and acyl-CoA:retinol acyltransferase in rat liver. J. Lipid Res. 28, 973–981.

Reverse-Phase HPLC Analysis of Retinol and Retinyl Esters 28. Furr, H.C., Cooper, D.A., Olson, J.A. (1986) Separation of retinyl esters by nonaqueous reversed-phase high-performance liquid chromatography. J. Chromatogr. 378, 45–53. 29. Redlich, C.A., Grauer, J.N., Van Bennekum, A.M., Clever, S.L., Ponn, R.B., Blaner, W.S. (1996) Characterization of carotenoid, vitamin A, and alpha-tocopheral levels in human lung tissue and pulmonary macrophages.

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Am. J. Respir. Crit. Care Med. 154, 1436–1443. 30. Kim, Y.K., Wassef, L., Hamberger, L., Piantedosi, R., Palczewski, K., Blaner, W.B., Quadro, L. (2008) Retinyl ester formation by lecithin:retinol acyltransferase (LRAT) is a key regulator of retinoid homeostasis in mouse embryogenesis. J. Biol. Chem. 283, 5611–5621.

Chapter 16 Detection of Retinoic Acid Catabolism with Reporter Systems and by In Situ Hybridization for CYP26 Enzymes Yasuo Sakai and Ursula C. Dräger Abstract Retinoic acid (RA), an active form of vitamin A, is essential for life in vertebrates, owing to its capacity of influencing expression of a sizable fraction of all genes and proteins. It functions via two modes: (1) as controlling ligand for specific transcription factors in the nucleus it stimulates or inhibits gene expression from RA response elements in gene promoters; (2) in non-genomic pathways it activates kinase-signaling cascades that converge with additional influences to regulate gene expression and mRNA translation. RA performs a critical role in morphogenesis of the developing embryo, which is reflected in spatio-temporally changing expression patterns of RA-synthesizing and RA-degrading enzymes and in its biophysical characteristics as a small diffusible lipid. Because its histological localization cannot be directly visualized for technical reasons, its sites of action in vivo are inferred from the locations of the metabolic enzymes and through use of two kinds of RA reporter systems. Here we explain techniques for use of RA reporter cells and RA reporter mice, and we describe in situ hybridization methods for the three major RA-degrading enzymes: CYP26A1, CYP26B1, and CYP26C1. Comparisons of the different indicators for sites of RA signaling demonstrate that local RA peaks and troughs are important for inferring some but not all locations of RA actions. When integrated within cells of living mice, expression of the RA reporter construct is rarely a simple measure of local RA levels, especially in the developing brain, but it appears to provide cues to an RA involvement in site-specific regulatory networks in combination with other spatial determinants. Key words: P450-linked oxidases, RALDHs, RARβ, CREB, non-canonical RA actions, Rossant RARE-lacZ mice, morphogenetic gradients, pattern formation.

1. Introduction Retinoic acid (RA), a vitamin A derivative, is known as key regulator of cellular growth, differentiation, morphogenesis, and homeostasis in vertebrates (1, 2). Expression of about one-sixth of the H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_16, © Springer Science+Business Media, LLC 2010

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human genome is estimated to be regulated by RA (3). Long before modern molecular techniques were available, experimental embryologists had postulated that the spatial patterning of the developing embryo involves diffusible morphogens, hypothetical compounds which emanate from specific sites and whose diffusion gradients convey positional morphogenetic information onto embryonic fields. In 1982 RA was identified as the first endogenously occurring compound that could perfectly mimic a natural morphogen (4). While over the following 25 years RA’s morphogen function was repeatedly corroborated or disputed, by now a morphogenetic RA gradient is considered essential for establishment of positional information, at least in the embryonic hindbrain; this RA gradient, however, depends critically on RAdegrading enzymes (5). RA is synthesized from food-derived vitamin A by several enzymes, with the last and irreversible step being mainly catalyzed by one of three retinaldehyde dehydrogenases (RALDH1, RALDH2, RALDH3). RA is inactivated primarily by different members of the cytochrome P450 oxidases in the presence of cytochrome P450 reductase (5). The most efficient and best characterized RA-degrading enzymes belong to the CYP26 subgroup: CYP26A1, CYP26B1, and CYP26C1 (6–14). They metabolize RA into oxidative products including 4-oxo-RA, 4-hydroxyRA, 18-hydroxy-RA, and 5,6- or 5,8-epoxy-RA (1, 2, 5, 15). Although it is still a subject of controversy whether some of the products, especially 4-oxo-RA, are active in vivo, observations on Cyp26/Raldh compound null mutants argue for removal of RA activities as the main biological function of the CYP26 enzymes (16). Due to the chemical characteristics of RA as a small amphipathic lipid, it will readily exit from RALDH-expressing cells and form a diffusion gradient in the surrounding tissue. In the early hindbrain such a chemical diffusion gradient, whose shape is potentially labile to various perturbations, is converted into a robust and morphogenetically informative RA gradient through localized RA degradation by CYP26A1, whose expression in turn is regulated by RA and fibroblast growth factor via feedback and feed-forward control loops (5). Morphogenetic RA gradients remain hypothetical so far for technical reasons, as no methods exist for the direct visualization of RA in the tissue. Standard RA detection methods based on high-performance liquid chromatography (HPLC) are poorly suited for questions on developmental pattern formation, because comparative RA measurements require pooling of tissues from exorbitant numbers of embryos. Histological techniques exist for visualization of RA-synthesizing and RAdegrading enzymes, as long as probes for specific enzymes are available, but unknown enzymes would be missed. RA reporter assays, which can detect RA by its biological activity in minute

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tissue samples, contributed significantly to the original identification of RA metabolic enzymes (e.g., (11, 17)). In this chapter we describe methods for detection of RA degradation with help of RA reporter systems and in situ hybridization by focusing on the developing retina and the rostral part of the embryo. All existing RA reporter systems known to us are based on the highly sensitive RA response element (RARE) from the promoter of the RA receptor β (RARβ) gene driving various reporter constructs, either in cell lines or in transgenic animal strains. Here we describe the most frequently used RA reporter mouse strain, which was generated by Rossant et al. (18), and the F9-teratocarcinoma-derived Sil-15 cell line generated by Wagner et al. (19). Both of these systems use the beta-galactosidase (lacZ) gene as reporter, whose product can be easily visualized by 5-bromo-4-chloro-3-indoyl-beta-Dgalactosidase (X-gal). Inspection of the head of an X-gal-reacted RARE-lacZ embryo, as illustrated here for embryonic day 12.5 (E12.5), shows strong labeling of the eye, which appears, however, to be interrupted by a horizontal unstained stripe (arrow, Fig. 16.1a). This indicates that the developing retina must contain three RA subdivisions along its dorso-ventral dimension. These subdivisions are more convincingly visible, when the retinas are dissected free from RARE-lacZ embryos or mice prior to the X-gal reaction (Fig. 16.1b). The RARE-lacZ reporter cell line (19) provides a powerful tool for analysis of the spatial arrangement of retinoid enzymes in minute embryonic tissues. When grown in 96-well plates, the responses of the RARE-lacZ cells can be quantified by serial dilutions of compounds added to the culture supernatants, and following culture and X-gal reaction, the results can be mea-

Fig. 16.1. (a) Head of a 12.5-day-old (E12.5) RARE-lacZ reporter embryo (18) reacted with X-gal. The arrow points to a horizontal gap in the strong eye labeling. (b) Retinas of RARE-lacZ embryos and young pups at 4 ages, reacted with X-gal, and viewed from back or front. For these preparations, the lenses are left in place, because they prevent the retinas from collapsing, they do not interfere with the X-gal reaction, and they are transparent. The horizontal lac-Z-free stripe is located just above the optic disc, through which the optic axons project to the brain.

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Fig. 16.2. (a) Titrations of RARE-lacZ cell (19) responses to the retinoids 3-demethyl RA, 3,7-didehydro RA, 4-oxo RA, 9-cis RA, and all-trans RA (20). (b) Example of a calibration curve for all-trans RA to calculate the concentrations of RA generated by tested tissue samples (20).

sured colorimetrically by an ELISA reader. The cell line responds strongly to the retinoids all-trans RA and 3,7-didehydro RA and only weakly to 13-demethyl RA, 4-oxo RA, and 9-cis RA (Fig. 16.2a). By comparing an all-trans RA standard curve (Fig. 16.2b) with the readings from the test samples, the magnitude of the generated RA can be estimated (20). Because of the tiny tissue quantities required, the RARE-lacZ cells allow measuring RA synthesis within samples dissected from restricted retina locations (Fig. 16.3a). As illustrated in the upper two assays of Fig. 16.3a, the embryonic retina synthesizes RA in its dorsal (D) part, twice as much ventrally (V), but none in a horizontal stripe region (21). The bottom assay (Fig. 16.3a) illustrates how to test, whether this result is due to local lack of RA synthesis or to active RA degradation within the stripe. For this test, both a low amount of RA (here 1 nM) and ketoconazole (40 μM), a general inhibitor of P450-linked oxidases, were added to the tissue samples (the optimal doses of RA and ketoconazole might be different for other tissues). The assay illustrates the presence of RA degradation mediated by a P450-linked oxidase activity in the stripe region (21). With regard to RA-synthesizing enzymes, the RARE-lacZ cells allow, in addition, determinations of enzymatic characteristics. For the illustrated example (Fig. 16.3b), tissue samples are dissected from defined locations along the dorso-ventral axis of several retinas, pooled, homogenized without detergent, and separated on a native isoelectric focusing (IEF) gel (21). The gel is cut into consecutive slices, and each slice is tested for RAsynthesizing activity from added retinaldehyde with the RARElacZ cells (22). Of the two enzymatic possibilities, aldehyde dehydrogenases require addition of nicotinamide adenine dinucleotide

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Fig. 16.3. (a) RARE-lacZ cell assays of defined regions from E13–14 retinas for RA synthesis and degradation (21). Small pieces were dissected from the retinas, as indicated in the sketches on the left, cultured for 15–20 h, and the supernatants were tested for RA contents with the RA reporter cells (19), as shown in the histograms on the right. For the bottom assay, dissected stripe regions were cultured with 1 nM RA added to the medium in the presence or absence of 40 μM ketoconazole, a general inhibitor of P450-linked cytochrome oxidases. (b) Zymography assays for RA-generating activities in six consecutive slivers along the dorso-ventral axis dissected from E13.5 retinas. The dots along the ordinates of the zymography traces represent single gel slices cut along a native isoelectric focusing (IEF) gel and tested for enzymatic activities, which can synthesize RA from 80 nM retinaldehyde in the presence of 2.4 mM NAD. In the dorsal samples, the IEF-zymography traces detect an enzymatic activity focused at pH 7.6, which is known to characterize RALDH1, and in the ventral samples an activity focused at pH 5.7 represents RALDH3 (17, 22).

(NAD) to the test wells, whereas aldehyde oxidases function with enzyme-bound cofactors. Most detectable RA synthetic activities in embryos and adults turn out to be NAD dependent, consistent with aldehyde dehydrogenases as the major RA-synthesizing enzymes in vertebrate tissues (23). Since all aldehyde dehydrogenases have similar molecular weights (55–56 kD), they are best distinguished by charge, which is the reason for separation by IEF. The IEF traces illustrated (Fig. 16.3b) reveal the presence of RALDH1 by its pI of 7.6 in the dorsal retina and of RALDH3 ventrally at pI 5.7. Within the horizontal stripe that contains the P450-linked oxidase activity, no RA-synthesizing enzyme is detectable. Because the RARE-lacZ cells do not allow identification of which particular P450-linked oxidase is expressed in the embryonic retina, in situ hybridizations for the best characterized subgroup, the CYP26A1, CYP26B1, and CYP26C1 enzymes, are illustrated here (Fig. 16.4). These specimens were prepared following a modification of the protocol by Henrique et al. (24,

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Fig. 16.4. In situ hybridizations for the three CYP26 enzymes (25). Cyp26a1 expression begins to form a broad stripe across the retina at E11.5, and Cyp26c1 expression forms a narrower one around E14, resulting in the patterns shown here for E15 (a). These expression patterns persist until juvenile ages. In newborn retinas, double labeling for the RA reporter and the RA-degrading enzymes reveals that Cyp26c1 occupies the center of the lacZ-free stripe and Cyp26a1 more or less fills it (b). Cyp26b1 is not expressed in the retina, as illustrated here by the unlabeled eye (arrow) in the whole-mounted head of an E12.5 embryo next to positive controls, the emerging whisker follicles (c).

25), and the mRNA localizations are visualized with the digoxigenin (DIG) labeling kit of Boehringer (for details, see below). At E11.5 Cyp26a1 begins to form a broad horizontal stripe across the embryonic retina, and Cyp26c1 expression starts around E14.0 as a narrower stripe in the same location (Fig. 16.4a). These expression patterns of Cyp26a1 and Cyp26c1 persist until postnatal stages. In newborn retinas of RARE-lacZ mice, double labeling for the RA reporter and the two RA-degrading enzymes reveals that Cyp26c1 occupies the center of the lacZ-free stripe and Cyp26a1 more or less fills it (Fig. 16.4b). Cyp26b1 is not expressed in the developing or mature retina of the mouse, at least not under normal conditions (25, 26). In the whole mount of an E12.5 head (Fig. 16.4c), the unlabeled eye (arrow) contrasts with the heavily labeled emerging whisker follicles as positive controls. RA reporter animals, in particular the RARE-lacZ transgenic strain (18) described here, have been valued highly in different RA-signaling studies on the developing organism done in many laboratories. In the embryonic retina, the comparisons of the responses in the RARE-lacZ cells (Fig. 16.2), the RARE-lacZ mice (Fig. 16.1), and the enzyme expression patterns (Figs. 16.3b and 16.4) demonstrate a good match: at the RALDH-expressing retina sites, both the cells and reporter mice indicate the presence of RA signaling, and at CYP26-expressing locations the two assays show lack of RA signaling. However, this congruence between the three signs for local RA signaling cannot be generalized. The three indicators agree in very early embryos and at selected sites such as the retina, but not at many other locations (27). A discrepancy is especially apparent in the brain, where strong RARE-lacZ labeling in the reporter mice rarely points to local expression of RA-synthesizing enzymes, and lack of reporter labeling does not necessarily indicate local RA degradation, at

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least as far as we have been able to determine (27). By comparison with the embryonic retina, overall RA synthesis within the brain is exceedingly low: in adult rats almost 90% of total brain RA is supplied by the blood circulation (28). Some experimental observations on convergence and discrepancies between local indicators for RA signaling are summarized graphically in Fig. 16.5; here the results from serial titrations of sample supernatants onto the RARE-lacZ cells are illustrated by single representative wells of the 96-well plates. The preparations from the older RARElacZ embryo (Fig. 16.5c) and the adult (Fig. 16.5d) are sections through the head and brain labeled for X-gal. A technique for preparation of X-gal-labeled serial sections is described below (see Section 3).

Fig. 16.5. Comparisons of the three indicators for RA signaling: (1) expression of RA-synthesizing enzymes, (2) RARElacZ cell responses, and (3) labeling patterns in RARE-lacZ mice. For the youngest embryos at E8.5 (a), “eye” designates the anterior neural ridge that contains the anlage for the future eye, and “brain” indicates the rostral neural plate, which will fold up to form the brain. The RARE-lacZ cell responses are shown here simplified as single representative wells from serial dilutions of culture supernatants in 96-well plates. Whereas the first two criteria, (1) expression of RALDHs and (2) RARE-lacZ cell responses, agree consistently and quantitatively, the RA signaling in the RARE-lacZ mice might concur or it might be discordant. Discrepancies are especially glaring in the brain (27).

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No global explanation exists for the discrepancies in observations between the RARE-lacZ cells and RARE-lacZ mice, but the extraordinary characteristics of the reporter strain (18) are compelling motivations for appreciating the disagreements as the most relevant information ahead of its time, which can be gained from RA reporter use. Regarding the RARE-lacZ cell responses, the only experimental variable is the amount of RA titrated into the culture wells, but the reporter animals promise to give cues to unexplained complexities of RA signaling in vivo, which are particularly prominent for the brain (29, 30). The mouse line is a very intriguing research tool: the specimens are easily and rapidly prepared, the results are visible as high-resolution, Golgi-like patterns, the labeling is extremely reproducible, and during embryonic and early postnatal brain development the patterns undergo changes over hours or days, which occur with clockwise precision. For instance, along the developing optic tract, the lacZ patterns in the brain trace out the future pathway for the RALDH-rich optic axons several days before the axons arrive (27), and in the developing cerebral cortex, intense lacZ expression precedes the select, de novo appearance of RALDH3-positive neurons in the postnatal medial cortex (31). When viewed over time, the changing reporter patterns appear to indicate events, in which RA is either already known to play a role or they depict well-known developmental processes, for which an RA involvement has yet to be tested. The main problem with the RARE-lacZ mice is a difficulty common to many transgenic strains: the reporter expression tends to become silenced by epigenetic mechanisms (27). How to deal with this problem is explained below (see Note 2). While an abundance of investigations on different RA reporter animals leaves no doubt that the local patterns of RA reporter expression require RA, and thus represent true RA signaling, for the brain a modification of RA signaling by unknown additional factors was confirmed repeatedly (e.g., (27, 32, 33)). Although RA is primarily known to function via the classical genomic pathway that involves binding of RA-liganded receptors to RAREs in gene promoters in the nucleus, a very large number of studies describe extra-nuclear, non-genomic RA actions via cellular signaling pathways, which converge with other factors to regulate both gene transcription as well as mRNA translation (e.g., (34–48)). This applies conspicuously to neurons: in most instances where specific RA-regulated gene expression was analyzed in neuronal cell cultures, non-genomic mechanisms were identified (e.g., (36, 38, 42, 46)). In a large fraction of the studied examples, the non-genomic RA-signaling cascades converge onto the cAMP response element-binding (CREB) transcription factor; RA and CREB signaling pathways share the coactivator CREB-binding protein (CBP) (49); and the promoter of the RARβ, on which the reporter assays are based, contains

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a functional CREB response element (CRE) (50). It was suggested that a convergence of RA and CREB signaling on RARβ results in synergistic enhancement (34), a positive loop that may contribute to the interpretation of unexplained RA actions in the functional brain (51, 52). In conclusion, both RA-synthesizing and RA-degrading enzymes bestow the essential conditions for RA signaling to occur in vertebrates. Whereas RA reporter cells provide the most sensitive tools for measuring RA levels, the RA reporter animals are invaluable for detecting the spatio-temporal locations of RA actions in the context of the living organism, where any form of RA signaling is bound to involve interactions with numerous additional factors and feedback loops, which for the most part still remain to be characterized.

2. Materials 2.1. RA Reporter Systems

1. The RARE-lacZ mice, which were generated in 1991 by Rossant et al. (18), are maintained in many individual laboratories, and they can also be purchased now from the Jackson laboratory (JAX). 2. The address of Dr. Michael Wagner, who made the RARElacZ Sil-15 cell line (19), is Department of Anatomy and Cell Biology, State University of New York, Health Sciences Center, 450 Clarkson Avenue, Brooklyn, NY 11203, USA.

2.1.1. Stock Solutions for LacZ Reactions

1. 0.5 M K3 Fe(CN)6 (potassium ferricyanate) in distilled water, store at room temperature (RT) shaded from light; 2. 0.5 M K4 Fe(CN)6 (potassium ferrocyanate) in distilled water, store at RT shaded from light; 3. 20 mg/ml 5-bromo-4-chloro-3-indoyl-beta-D-galactosidase (X-gal) in dimethylformamide (DMF), store at −20◦ C shaded from light; 4. 1 M MgCl2 in distilled water, store at RT.

2.1.2. Culture of RARE-LacZ Cells (19)

1. Gelatin-coated tissue culture flasks and 96-well flat-bottom plates prepared by covering with 0.2% aqueous gelatin solution for 2 h at RT and 2x washes in sterile PBS (pH 7.4); 2. L15–CO2 medium (Speciality Media) with 10% fetal calf serum and 0.8 mg/ml G418 (Gibco) to select for transfected F9 RARE-lacZ cells; 3. Cell fixative: 1% glutaraldehyde, 0.1 mM MgCl2 in 0.1 M phosphate buffer, pH 7.0;

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4. Development buffer: a fresh mixture of 0.2% X-gal in PBS with 1 mM MgCl2 , 0.3 mM potassium ferrocyanate, and 0.3 mM potassium ferricyanate; 5. A 0.1 M stock solution of all-trans RA (Sigma-Aldrich) in dimethyl sulfoxide (DMSO), prepared under dim yellow light and stored frozen at −80◦ C; 6. Ketoconazole (Sigma-Aldrich). 2.1.3. Reagents for RARE-LacZ Mice

1. In order to allow for sufficient time, it is better to perform complex dissections of live tissues in any available, even outdated, serum-containing tissue culture medium, rather than PBS; 2. Fixative for tissue whole mounts: 0.2% glutaraldehyde in 0.1 M PBS with 1 mM MgCl2 ; 3. Fixative for perfusion: 2% glutaraldehyde in 0.1 M PBS with 1 mM MgCl2 ; 4. 4% paraformaldehyde (PFA) in PBS stored in aliquots at −20◦ C (see Note 1).

2.2. In Situ Hybridization 2.2.1. Preparation of RNA Probes

The probes are prepared following the protocol provided with Boehringer’s digoxigenin (DIG) labeling kit: 1. Linear plasmid DNA for reaction templates: cut plasmid by restriction enzyme (3 -protruding or blunt) and adjust to 1 μg/ml in TE; stored at −20◦ C; 2. DIG labeling kit: 10x reaction buffer, DIG-RNA labeling mixture, RNase inhibitor, stored at −20◦ C; 3. RNA polymerases: T3, T7, SP6 stored at −20◦ C; 4. RNase-free water (not DEPC treated); 5. DNase I (RNase free) stored at −20◦ C; 6. Gycogen (RNase free): 20 mg/ml stored at −20◦ C; 7. 4 M LiCl stored at RT; 8. TE/SDS: freshly prepare 1:1 TE/10% SDS.

2.2.2. Whole-Mount In Situ Hybridization

This is a modification of the procedure described by Henrique et al. (24), which is available on the web site of the Rossant laboratory. 1. 4% paraformaldehyde (PFA) in PBS, stored in aliquots at −20◦ C (see Note 1); 2. PTW: 0.1% Tween-20 in PBS, stored at RT; 3. Proteinase K: 10 mg/ml in purified water, stored in aliquots at −20◦ C;

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4. Glutaraldehyde stored at 4◦ C; 5. Hybridization buffer: 25 ml of formamide, 3.25 ml of 20× SSC (pH 5.0), 0.5 ml of 0.5 M EDTA (pH 8.0), 125 μl of yeast RNA (20 mg/ml), 1 ml of Tween-20 (10%), 2.5 ml of CHPS (10%), 100 μl of heparin (50 mg/ml), 17.5 ml of H2 O (DEPC treated); store at −20◦ C; 6. DIG labeling kit (Boehringer) stored at −20◦ C; 7. MABT: 100 mM maleic acid, 150 mM NaCl, pH 7.5, 0.5% Tween-20; stored at RT; 8. Boehringer blocking reagent (BBR): 10% (w/v) BBR in MABT (pre-warmed at 65◦ C) stored in aliquots at −20◦ C; 9. Heat-treated sheep serum stored in aliquots at −20◦ C; 10. AP-anti-DIG antibody (Boehringer) stored at 4◦ C; 11. BM purple (Boehringer) stored at 4◦ C.

3. Methods 3.1. RA Reporter Systems 3.1.1. RARE-LacZ Cell Assays

1. Grow the RARE-lacZ cells in L15–5% CO2 tissue culture medium adhered to gelatin-coated culture flasks. Dissociate the cells by standard trypsination. Pipette the cells in 75 μl L15 medium per well into gelatin-coated 96-well plates and grow to confluency. 2. Dissect small pieces of live tissue under dim yellow light and culture overnight in small volumes of L15 medium. The tissue sizes and medium volumes ought to be adjusted to maintain the tissue healthy over the culture period. Collect the supernatants, pipette them into the 96-well plate containing the RARE-lacZ cells, and titrate them out in serial dilutions. 3. Culture the plates for 12–15 h (overnight) in the incubator. Wash briefly with PBS and fix in 1% glutaraldehyde. After 15 min, wash four times with PBS. Pipette 50 μl of freshly prepared development buffer (1 mg/ml X-gal, 0.3 mM potassium ferrocyanate, 0.3 mM potassium ferricyanate, 1 mM MgCl2 ) per well and incubate the plates at 37◦ C. Development of the color reaction takes from minutes to hours, depending on the RA contents of the supernatants, but the reaction can be continued for days. The color intensities can be quantified in an ELISA reader. 4. To achieve quantification beyond inter-sample comparisons, a standard dilution of all-trans RA (see Fig. 16.3b) needs

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to be run in parallel with the tested samples. The reason for this is variability in the RARE-lacZ cell responses. Despite growth of the cells in G418 selection medium, the cells can lose responsiveness over time, usually after about 25 passages. It is advisable to keep most of the cells frozen in aliquots that can be thawed individually. 5. To test for the presence of RA-degrading enzymes in the cultured tissues, a small amount of RA is added to the culture medium and its degradation by ketoconazole is assayed with the RARE-lacZ cells. For the experiments shown in Fig. 16.3a, the degradation of 1 nM RA by 40 μM ketoconazole was tested. Because the existing IEF-zymography assay with the RARE-lacZ cells was only developed to distinguish RA-synthesizing but not RA-degrading enzymes, it is not described here further, in addition to the details given in the legend of Fig. 16.3b (22). For detection of RA-degrading enzymes, it is necessary to prepare microsomal fractions, which can be done from intermediate-sized embryonic tissue samples, if appropriate equipment is available, such as an airfuge for ultracentrifugation of small tissue volumes (20). 3.1.2. RARE-LacZ Reporter Mice

Whole embryos or tissues of the RARE-lacZ reporter mice are dissected in chilled, serum-containing tissue culture medium, transferred to tubes of appropriate sizes, and fixed in 0.2% glutaraldehyde with 1 mM MgCl2 on a gentle shaker. After a variable time (5–40 min) dependent on the tissue sizes or consistencies, the glutaraldehyde is aspirated and exchanged with the lacZ staining solution (1 mg/ml X-gal, 0.3 mM potassium ferrocyanate, 0.3 mM potassium ferricyanate, 1 mM MgCl2 ). To avoid sticking of the tissue samples to the walls of tubes, dishes, or pipettes, a brief rinse of all surfaces with serum-containing medium is recommended. When a desired staining intensity is achieved, the tissues are briefly post-fixed with 4% paraformaldehyde. The lacZ enzyme activity detectable with the X-gal reaction is rather insensitive to low levels of glutaraldehyde, but it is rapidly terminated by 4% paraformaldehyde. The 0.2% glutaraldehyde used for fixation of whole mounts is insufficient for transcardial perfusion, where it does not result in any hardening of the tissues. In order to prepare serial sections from older embryos and postnatal RARE-lacZ mice, we briefly perfuse transcardially under a stereomicroscope with PBS containing 2% glutaraldehyde and 1 mM MgCl2 . This glutaraldehyde concentration results in very efficient fixation, but it will also cause a slow inactivation of the X-gal detectable lacZ activity, unless the tissues are processed right away. The embryonic heads (Fig. 16.5c) or postnatal brains (Fig. 16.5d) are rapidly embedded in low melting point

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agarose and sliced at 100–150 μm on a vibratome. The sections are serially collected in 24-well plates in MgCl2 -containing PBS placed on wet ice. The plate is reacted with X-gal until an optimal contrast is achieved, and the reaction is terminated with 4% paraformaldehyde. A common problem with RARE-lacZ mice is an epigenetic inactivation of the lacZ gene, which happens to a variable extent between reporter mice, as well as over the lifetime in individual mice, a chance occurrence that is unavoidable even in a well-maintained RARE-lacZ mouse colony (27). In order to avoid wasting time with sectioning of poor responder mice, it is advisable to punch out a piece of the spinal cord from the embryos, or to dissect retinas from postnatal mice following perfusion, and to test with X-gal: from a good responder these tissues show a color reaction in a few seconds (see Note 2). 3.2. In Situ Hybridization 3.2.1. Preparation of RNA Probes

1. Incubate the following reaction mixture at 37◦ C: 2 μl of 10x reaction buffer, 2 μl of DIG RNA labeling mixture, 1 μl of linear plasmid DNA, 1 μl of RNase inhibitor, 2 μl of RNA polymerase, 12 μl of RNase-free water; 2. Incubate for 60 min and check 1 μl of the reaction mixture on an agarose gel; 3. Add 2 μl of DNase I and incubate for 15 min at 37◦ C; 4. Add 1 μl of glycogen and mix well; 5. Add 1/10 volume of 4 M LiCl; 6. Add 2.5–3.0 volumes of chilled ethanol; 7. Mix well by vortex and incubate for 30 min at −80◦ C; 8. Centrifuge at 12,000 g for 15 min at 4◦ C; 9. Decant ethanol and wash the pellet with 100 μl of 70% ethanol; 10. Centrifuge again for 5 min; 11. Remove ethanol and dry the pellet; 12. Resuspend in 50 μl of TE/SDS, mix well, and store at −80◦ C.

3.2.2. Whole-Mount In Situ Hybridization

1. Dissect out tissues in chilled culture medium (10% fetal calf serum in DMEM). The neural retinas have to be dissected without the lenses; embryonic heads are cut off from the bodies and opened along the sagittal midline; 2. Fix in 4% paraformaldehyde in PBS at 4◦ C by rocking overnight; 3. Wash twice in PTW for 10 min;

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4. Wash with 50% methanol/PTW, then 100% methanol twice for 10 min. The samples can be stored at this point at −20◦ C for a few months. Day 1 5. Rehydrate samples through 50% methanol/PTW for 10 min and wash twice with PTW for 10 min; 6. Treat with 10 μg/ml proteinase K in PTW for 5–15 min (see Note 3); 7. Rinse briefly with PTW and post-fix for 20 min in 4% paraformaldehyde + 0.1% glutaraldehyde in PTW; 8. Rinse and wash once for 10 min with PTW. Transfer samples to screw cap 2 ml tubes; 9. Rinse once with 1:1 PTW/hybridization mix. Let samples sink; 10. Rinse with 0.5 ml hybridization mix. Let samples sink; 11. Replace with 0.5 ml hybridization mix and incubate horizontally for 1 h at 70◦ C; 12. Add 0.5 ml pre-warmed hybridization mix with 0.1∼1 μg/ml DIG-labeled RNA. Immediately place at 70◦ C; 13. Incubate overnight horizontally at 70◦ C. Rock once after 20–30 min (see Note 4). Day 2 14. Rinse twice with pre-warmed (70◦ C) hybridization mix (see Note 5); 15. Wash twice for 30 min at 70◦ C with 1 ml pre-warmed hybridization mix; 16. Wash for 20 min at 70◦ C with 1 ml of pre-warmed 50% hybridization mix/MABT; 17. Rinse three times with 1 ml of MABT; 18. Wash twice for 30 min with 1 ml MABT; 19. Incubate with 1 ml of MABT + 2% Boehringer blocking reagent (BBR) for 1 h at RT; 20. Incubate with 1 ml of MABT + 2% BBR + 20% heat-treated sheep serum for 1 h at RT; 21. Incubate with 1 ml of MABT + 2% BBR + 20% serum + 1/2000 dilution of AP-anti-DIG antibody (Boehringer) overnight at 4◦ C by rocking. Day 3 22. Rinse three times with 1 ml of MABT;

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23. Wash three times for 1 h with 1.5 ml of MABT by rocking; 24. Wash with 1.5 ml of MABT O/N at 4◦ C by rocking (see Note 6). Day 4 25. Replace with 1.5 ml of BM purple (Boehringer) to develop a color reaction at 37◦ C or RT; 26. To stop the reaction, rinse and wash twice with PTW for 10 min, then refix with 4% PFA; 27. For taking photographs, transfer samples into Tissue-Tek cryostat-embedding medium diluted 1:1 with PBS.

4. Notes 1. Although it is commonly recommended that paraformaldehyde (PFA) needs to be prepared freshly, we have used frozen aliquots of PFA solution routinely and without any detectable problems. 2. The lacZ gene in the RARE-lacZ mice tends to undergo epigenetic inactivation, a rather unpredictable process that is progressive both with the age of individual mice and between generations. The inactivation is heritable, between generations and ontogenetically, as visible in different-sized X-gal-positive cell clones (27). When nothing is done, the RARE-lacZ mouse colony will over time turn into nonresponders, in which all mice are still positive for lacZ by PCR, but the enzyme is rarely expressed. Although the epigenetic silencing ought to be reversible, we have not been able to achieve this, despite elaborate attempts. The only measure is to select for super-responders as breeders of a new colony. A reasonably efficient method is to out-cross all RARE-lacZ mice, which are usually maintained as homozygotes, to CD1 mice, the background strain. The newborn litters from each breeding pair are killed and one retina of every pup is tested with X-gal. The male and female from the breeding pairs with the highest proportion of positive pups are then selected to found a restored colony, and all other mice are killed. 3. Incubation time for proteinase K depends on the sizes or stages of the samples. Begin with 10 min of reaction time and titrate the activity of proteinase K. 4. To avoid damage of the samples or attachment to the tube walls, do not move the tubes during hybridization.

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5. It is very important to avoid cooling down the samples during the manipulations. 6. The final washing step is critical, and it is better to spend too much than too little time. References 1. Ross, S.A., McCaffery, P., Dräger, U.C., De Luca, L.M. (2000) Retinoids in embryonal development. Physiol. Rev. 80, 1021–1054. 2. Niederreither, K., Dollé, P. (2008) Retinoic acid in development: Towards an integrated view. Nat. Rev. Genet. 9, 541–553. 3. Cawley, S., Bekiranov, S., Ng, H.H., Kapranov, P., Sekinger, E.A., Kampa, D., Piccolboni, A., Sementchenko, V., Cheng, J., Williams, A.J., Wheeler, R., Wong, B., Drenkow, J., Yamanaka, M., Patel, S., Brubaker, S., Tammana, H., Helt, G., Struhl, K., Gingeras, T.R. (2004) Unbiased mapping of transcription factor binding sites along human chromosomes 21 and 22 points to widespread regulation of noncoding RNAs. Cell 116, 499–509. 4. Tickle, C., Alberts, B., Wolpert, L., Lee, J. (1982) Local application of retinoic acid to the limb bud mimics the action of the polarizing region. Nature 296, 564–566. 5. White, R.J., Schilling, T.F. (2008) How degrading: Cyp26s in hindbrain development. Dev. Dyn. 237, 2775–2790. 6. White, J.A., Guo, Y.D., Baetz, K., BeckettJones, B., Bonasoro, J., Hsu, K.E., Dilworth, F.J., Jones, G., Petkovich, M. (1996) Identification of the retinoic acid-inducible all-trans-retinoic acid 4-hydroxylase. J. Biol. Chem. 271, 29922–29927. 7. Fujii, H., Sato, T., Kaneko, S., Gotoh, O., Fujii-Kuriyama, Y., Osawa, K., Kato, S., Hamada, H. (1997) Metabolic inactivation of retinoic acid by a novel P450 differentially expressed in developing mouse embryo. EMBO J. 16, 4163–4173. 8. Ray, W.J., Bain, G., Yao, M., Gottlieb, D.I. (1997) CYP26, a novel mammalian cytochrome P450, is induced by retinoic acid and defines a new family. J. Biol. Chem. 272, 18702–18708. 9. MacLean, G., Abu-Abed, S., Dolle, P., Tahayato, A., Chambon, P., Petkovich, M. (2001) Cloning of a novel retinoic-acid metabolizing cytochrome P450, Cyp26B1, and comparative expression analysis with Cyp26A1 during early murine development. Mech. Dev. 107, 195–201. 10. Tahayato, A., Dolle, P., Petkovich, M. (2003) Cyp26C1 encodes a novel retinoic acid-

11.

12.

13.

14.

15.

16.

17.

metabolizing enzyme expressed in the hindbrain, inner ear, first branchial arch and tooth buds during murine development. Gene Express. Patterns 3, 449–454. Sakai, Y., Meno, C., Fujii, H., Nishino, J., Shiratori, H., Saijoh, Y., Rossant, J., Hamada, H. (2001) The retinoic acidinactivating enzyme CYP26 is essential for establishing an uneven distribution of retinoic acid along the anterio-posterior axis within the mouse embryo. Genes Dev. 15, 213–225. Abu-Abed, S., Dolle, P., Metzger, D., Beckett, B., Chambon, P., Petkovich, M. (2001) The retinoic acid-metabolizing enzyme, CYP26A1, is essential for normal hindbrain patterning, vertebral identity, and development of posterior structures. Genes Dev. 15, 226–240. Yashiro, K., Zhao, X., Uehara, M., Yamashita, K., Nishijima, M., Nishino, J., Saijoh, Y., Sakai, Y., Hamada, H. (2004) Regulation of retinoic acid distribution is required for proximo-distal patterning and outgrowth of the developing mouse limb. Dev. Cell 6, 411–422. Uehara, M., Yashiro, K., Mamiya, S., Nishino, J., Chambon, P., Dolle, P., Sakai, Y. (2007) CYP26A1 and CYP26C1 cooperatively regulate anterior-posterior patterning of the developing brain and the production of migratory cranial neural crest cells in the mouse. Dev. Biol. 302, 399–411. Lutz, J.D., Dixit, V., Yeung, C.K., Dickmann, L.J., Zelter, A., Thatcher, J.E., Nelson, W.L., Isoherranen, N. (2009) Expression and functional characterization of cytochrome P450 26A1, a retinoic acid hydroxylase. Biochem. Pharmacol. 15, 258–268. Niederreither, K., Abu-Abed, S., Schuhbaur, B., Petkovich, M., Chambon, P., Dolle, P. (2002) Genetic evidence that oxidative derivatives of retinoic acid are not involved in retinoid signaling during mouse development. Nat. Genet. 31, 84–88. McCaffery, P., Lee, M.-O., Wagner, M.A., Sladek, N.E., Dräger, U.C. (1992) Asymmetrical retinoic acid synthesis in the

Detection of Retinoic Acid Catabolism with Reporter Systems

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

dorso-ventral axis of the retina. Development 115, 371–382. Rossant, J., Zirngibl, R., Cado, D., Shago, M., Giguère, V. (1991) Expression of a retinoic acid response element-hsplacZ transgene defines specific domains of transcriptional activity during mouse embryogenesis. Genes Dev. 5, 1333–1344. Wagner, M., Han, B., Jessell, T.M. (1992) Regional differences in retinoid release from embryonic neural tissue detected by an in vitro reporter assay. Development 116, 55–66. Yamamoto, M., Dräger, U.C., McCaffery, P. (1998) A novel assay for retinoic acid catabolic enzymes shows high expression in the developing hindbrain. Dev. Brain Res. 107, 103–111. McCaffery, P., Wagner, E., O Neil, J., Petkovich, M., Dräger, U.C. (1999) Dorsal and ventral retinal territories defined by retinoic acid synthesis, break-down and nuclear receptor expression. Mech. Dev. 82, 119–30. Corrections 85, 203–214. McCaffery, P., Dräger, U.C. (1997) A sensitive bioassay for enzymes that synthesize retinoic acid. Brain Res. Protocols 1, 232–236. McCaffery, P., Dräger, U.C. (1995) Retinoic acid synthesizing enzymes in the embryonic and adult vertebrate. Adv. Exp. Med. Biol. 372, 173–183. Henrique, D., Adam, J., Myat, A., Chitnis, A., Lewis, J., Ish-Horowicz, D. (1995) Expression of a Delta homologue in prospective neurons in the chick. Nature 375, 787–790. Sakai, Y., Luo, T., McCaffery, P., Hamada, H., Dräger, U.C. (2004) CYP26A1 and CYP26C1 cooperate in degrading retinoic acid within the equatorial retina during later eye development. Dev. Biol. 276, 143–157. Luo, T., Sakai, Y., Wagner, E., Dräger, U.C. (2006) Retinoids, eye development and maturation of visual function. J. Neurobiol. 66, 677–686. Luo, T., Wagner, E., Grün, F., Dräger, U.C. (2004) Retinoic acid signaling in the brain marks formation of optic projections, maturation of the dorsal telencephalon, and function of limbic sites. J. Comp. Neurol. 470, 297–316. Kurlandsky, S.B., Gamble, M.V., Ramakrishnan, R., Blaner, W.S. (1995) Plasma delivery of retinoic acid to tissues in the rat. J. Biol. Chem. 270, 17850–17857. Dräger, U.C. (2006) Retinoic acid signaling in the functioning brain. Science STKE 324, pe10.

293

30. Lane, M.A., Bailey, S.J. (2005) Role of retinoid signalling in the adult brain. Prog. Neurobiol. 75, 275–293. 31. Luo, T., Wagner, E., Crandall, J.E., Dräger, U.C. (2004) A retinoic-acid critical period in the early postnatal mouse brain. Biol. Psychiat. 56, 971–980. 32. Liao, W.L., Wang, H.F., Tsai, H.C., Chambon, P., Wagner, M., Kakizuka, A., Liu, F.C. (2005) Retinoid signaling competence and RARbeta-mediated gene regulation in the developing mammalian telencephalon. Dev. Dyn. 232, 887–900. 33. Molotkova, N., Molotkov, A., Duester, G. (2007) Role of retinoic acid during forebrain development begins late when Raldh3 generates retinoic acid in the ventral subventricular zone. Dev. Biol. 303, 601–610. 34. Aggarwal, S., Kim, S.W., Cheon, K., Tabassam, F.H., Yoon, J.H., Koo, J.S. (2006) Nonclassical action of retinoic acid on the activation of the cAMP response elementbinding protein in normal human bronchial epithelial cells. Mol. Biol. Cell 17, 566–575. 35. Alique, M., Lucio-Cazana, F.J., Moreno, V., Xu, Q., Konta, T., Nakayama, K., Furusu, A., Sepulveda, J.C., Kitamura, M. (2007) Upregulation of cyclooxygenases by retinoic acid in rat mesangial cells. Pharmacology 79, 57–64. 36. Canon, E., Cosgaya, J.M., Scsucova, S., Aranda, A. (2004) Rapid effects of retinoic acid on CREB and ERK phosphorylation in neuronal cells. Mol. Biol. Cell. 15, 5583– 5592. 37. Dey, N., De, P.K., Wang, M., Zhang, H., Dobrota, E.A., Robertson, K.A., Durden, D.L. (2007) CSK controls retinoic acid receptor (RAR) signaling: A RAR-cSRC signaling axis is required for neuritogenic differentiation. Mol. Cell. Biol. 27, 4179–4197. 38. Fernandes, N.D., Sun, Y., Price, B.D. (2007) Activation of ATM s kinase activity by retinoic acid is required for CREB-dependent differentiation of neuroblastoma cells. J. Biol. Chem. 282, 16577–16584. 39. Hughes, P.J., Zhao, Y., Chandraratna, R.A., Brown, G. (2006) Retinoid-mediated stimulation of steroid sulfatase activity in myeloid leukemic cell lines requires RARalpha and RXR and involves the phosphoinositide 3-kinase and ERK-MAP kinase pathways. J. Cell. Biochem. 97, 327–350. 40. Kim, S.W., Hong, J.S., Ryu, S.H., Chung, W.C., Yoon, J.H., Koo, J.S. (2007) Regulation of mucin gene expression by CREB via a nonclassical retinoic acid signaling pathway. Mol. Cell Biol. 27, 6933–6947.

294

Sakai and Dräger

41. Lal, L., Li, Y., Smith, J., Sassano, A., Uddin, S., Parmar, S., Tallman, M.S., Minucci, S., Hay, N., Platanias, L.C. (2005) Activation of the p70 S6 kinase by all-trans-retinoic acid in acute promyelocytic leukemia cells. Blood 105, 1669–1677. 42. Lee, J.H., Kim, K.T. (2004) Induction of cyclin-dependent kinase 5 and its activator p35 through the extracellular-signalregulated kinase and protein kinase A pathways during retinoic-acid mediated neuronal differentiation in human neuroblastoma SKN-BE(2)C cells. J. Neurochem. 91, 634–647. 43. Liao, Y.P., Ho, S.Y., Liou, J.C. (2004) Nongenomic regulation of transmitter release by retinoic acid at developing motoneurons in Xenopus cell culture. J. Cell Sci. 117, 2917–2924. 44. Liou, J.C., Ho, S.Y., Shen, M.R., Liao, Y.P., Chiu, W.T., Kang, K.H. (2005) A rapid, nongenomic pathway facilitates the synaptic transmission induced by retinoic acid at the developing synapse. J. Cell Sci. 118, 4721–4730. 45. Lopez-Andreo, M.J., Torrecillas, A., ConesaZamora, P., Corbalan-Garcia, S., GomezFernandez, J.C. (2005) Retinoic acid as a modulator of the activity of protein kinase Calpha. Biochemistry 44, 11353–11360. 46. Masia, S., Alvarez, S., de Lera, A.R., Barettino, D. (2007) Rapid, non-genomic actions of retinoic acid on phosphatidyl-Inositol-3-

47.

48.

49.

50.

51.

52.

kinase signaling pathway mediated by the retinoic acid receptor. Mol. Endocrinol. 21, 2391–2402. Ochoa, W.F., Torrecillas, A., Fita, I., Verdaguer, N., Corbalan-Garcia, S., GomezFernandez, J.C. (2003) Retinoic acid binds to the C2-domain of protein kinase C(alpha). Biochemistry 42, 8774–8779. Poon, M.M., Chen, L. (2008) Retinoic acidgated sequence-specific translational control by RARalpha. Proc. Natl. Acad. Sci. USA 105, 20303–20308. Rosenfeld, M.G., Lunyak, V., Glass, C.K. (2006) Sensors and signals: A coactivator/corepressor/epigenetic code for integrating signal-dependent programs of transcriptional response. Genes Dev. 20, 1405– 1428. Kruyt, F.A., Folkers, G., van den Brink, C.E., van der Saag, P.T. (1992) A cyclic AMP response element is involved in retinoic aciddependent RAR beta 2 promoter activation. Nucleic Acids Res. 20, 6393–6399. Dräger, U.C., Luo, T., Wagner, E. (2008) Retinoic acid function in central visual pathways. In: Chalupa, L.M., Williams, R. (eds.), Eye, Retina and the Visual Systems of the Mouse, MIT Press, Cambridge, MA, pp. 363–376. Luo, T., Wagner, E., Dräger, U.C. (2009) Integrating retinoic acid signaling with brain function. Dev. Psychol. 45, 139–150.

Chapter 17 Diet in Vitamin A Research A. Catharine Ross Abstract A properly formulated diet is an essential underpinning for all in vivo research. This chapter focuses on the use of diet in retinoid research from two perspectives: human research, in which diet is usually variable and analysis of dietary intake is paramount to interpreting the study’s results, and animal (rodent) research, in which diet is imposed as a factor in the experimental design, and the diet consumed is usually monotonous. Many standard rodent diets are nonpurified and the amount of vitamin A in the diet is unknown. Moreover, it is likely to be much higher than expected from the label. By using a wellformulated purified diet with an exact amount of vitamin A, retinoid status in rodents can be closely controlled to create specific physiological conditions that represent the wide range of vitamin A status present in human populations. Key words: Diet, dietary assessment, retinol, retinoic acid, vitamin A deficiency, vitamin A supplementation. Abbreviations: AIN American Institute of Nutrition IOM Institute of Medicine NRC National Research Council RA retinoic acid RDA recommended dietary allowance.

1. Introduction Diet provides all vertebrates with the macronutrients needed for energy production and tissue anabolism, with minerals, such as calcium and phosphorus, that serve a structural role, and with numerous micronutrients that play an essential role as cofactors in metabolism and as regulators of metabolic functions. Thus, a properly formulated diet is essential for practically all in vivo H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_17, © Springer Science+Business Media, LLC 2010

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research. In the early 1900s, McCollum and Davis and Osborne and Mendel demonstrated the nutritional requirement for “fatsoluble A” and deduced several of the vitamin’s most important effects. It is now well established that all vertebrates require vitamin A for adequate growth, cell and tissue differentiation, vision, development and function of the immune system, and survival. Although the amount of vitamin A (retinol) required for these functions, in the range of micrograms per day (1), is a trace component of the diet, its biological impact is wide ranging. This chapter focuses on the practical use of diet in retinoid research from two perspectives: human diets and animal (rodent) diets. In human research on vitamin A, vitamin A intake is seldom controlled, although this has been done in a few studies. However, the assessment of vitamin A intake is important in population-based and epidemiological research. Humans consume a variety of foods and thus obtain their vitamin A in multiple forms – preformed retinol and provitamin A carotenoids – and the variable consumption of different foods contributes significantly to the problem of analyzing how much vitamin A has been consumed. In contrast, for animal studies, diet is a controllable factor and the diet is usually the same over time. The amount of vitamin A present in the diet establishes the animal’s vitamin A status, which, in turn, can determine outcomes, such as levels of gene expression and rates of metabolic processes. However, many standard animal diets are nonpurified and the vitamin A content can be variable. Some of the issues in planning studies and selecting appropriate methods are considered here.

2. Materials As described below, controlling diet and for assessing diet are fundamentally different and thus the materials needed are also different. Assessing diet intake requires a method for recording exact intake (food intake diary), or the frequency of consumption of certain foods (food frequency questionnaire, FFQ), and access to an appropriately detailed food composition database, as further discussed in Section 2.1 and its subsections. For animal diets, discussed in Section 3.1 and its subsections, a specific formulation must be decided on and the diet either prepared or purchased according to the formulation from a commercial supplier. As discussed below, investigators first need to determine the needs of their study, then formulate an appropriate approach to the diet to be used.

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2.1. Human Studies: Controlling and Assessing the Diet 2.1.1. Controlling the Diet

In human studies, the intake of vitamin A has seldom been strictly controlled. Some recent studies have controlled intake to some extent and these studies provide guidance into the planning that is required (see Note 1). Regarding vitamin A, Haskell et al. (2) fed diets low in vitamin A to adult male Bangladeshi volunteers (with initially low vitamin A status evidenced by plasma retinol of 0.51–1.22 μmol/l at the beginning of the study), then added graded supplements of retinol to produce differences in the total body vitamin A pool size. All meals were provided at the study’s research center for a period of 129 days. The basal diet consisted primarily of rice and lentils with small amounts of curried meats (mutton, chicken, and fish), vegetables (cabbage, cauliflower, white squash, and white potato), and fruit (banana), all with low vitamin A content. In another stable isotope dilution study that was conducted in the United States to evaluate intestinal and postintestinal β-carotene conversion, Tang et al. (3) fed controlled diets to older men and women while subjects resided in their research facility at Tufts University for the first 10 days of the study. For days 11–57 of the study the subjects returned to their homes. They were provided with instructions from the study dietitian to consume only fruits and vegetables from a list of low-carotene foods and were counseled to abstain from multivitamins, minerals, nutritional supplements, fortified cereals, and fish liver oil. Recently, Ahmad et al. (4) conducted a controlled feeding study in 36 healthy Bangladeshi men (20–30 years of age), designed to assess whether total body vitamin A pool size, determined by a deuterium-labeled retinol dilution technique, predicted immune response to immunization. This study used a 2-month residential period in which a low-vitamin A basal diet was fed, modeled on the traditional Bangladeshi diet that provided the equivalent of 40 μg retinol/day. These studies illustrate the logistical details required to carefully control vitamin A intake in humans.

2.1.2. Assessing the Diet

By far the majority of human studies have relied on methods for assessing vitamin A or carotenoid intake in community-dwelling subjects. Subjects either have consumed their usual diet without any control or have been instructed to follow dietary advice in terms of avoiding, limiting, or consuming certain foods. A number of epidemiological investigations have investigated the intake of dietary vitamin A intake, either as preformed retinol or

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provitamin A carotenoids, or both, in studies of cross-sectional, case–control, or prospective designs (5–8). Two main approaches to the collection of dietary information have been developed: (i) dietary recordings followed by nutrient analysis or recalls and (ii) food frequency questionnaire, FFQ, often focused on the frequency of intake of certain foods high in vitamin A or carotenoids. Either assessment method is complicated by the multiple forms of vitamin A present in the foods in most diets. Preformed vitamin A is not only found as retinol or retinyl esters in foods of animal origin – typically milk, cheese, and liver – but also present in some nutritional supplements and fortified foods such as enriched breakfast cereals. Provitamin A carotenoids are present in numerous vegetables, being highest in green leafy and yellow vegetables, and certain fruits, such as mangoes and oranges. To assess total vitamin A intake, retinol and carotenoids must be calculated separately (1), followed by conversion to a basic unit.

3. Methods 3.1. Units

The units for expressing vitamin A in foods and supplements have changed over time and can be confusing. Thus, before collecting dietary information, is it necessary to have a plan for converting information to a standard unit. Nutritional units currently in use include the international unit (IU), the retinol equivalent (RE), and for human diets only, the Retinol Activity Equivalent (RAE) (1). One IU (also equal to 1 USP unit) is equivalent to 0.3 μg all-trans-retinol (molecular weight 286.6), 0.55 g retinyl palmitate (molecular weight 525), and 0.6 μg of β-carotene. The current unit for human research, established by the Institute of Medicine (IOM) in 2001, is the retinol activity equivalent (RAE), Table 17.1. This table presents the nutritional equivalency among retinol, β-carotene in supplements (e.g., in oily solution from which it is readily absorbed), β-carotene in foods (embedded in food matrix and therefore less bioavailable), and provitamin A carotenoids other than β-carotene (α-carotene or β-cryptoxanthin in foods). The IOM established the new RAE unit because research on carotenoid bioavailability in human subjects had established that carotenoids present in foods are converted into retinol less efficiently than was previously thought and less efficiently than from supplements (1). In analyzing dietary data and comparing study results, careful attention must be given to the form of vitamin A in foods consumed as well as the units in which vitamin A is reported in tables of food composition (see Note 2).

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Table 17.1 Comparison of the 1989 National Research Council and 2001 Institute of Medicine Interconversion of Vitamin A and Carotenoid Units NRC (1989)

IOM (2001)

1 retinol equivalent (μg RE)

1 retinol equivalent (μg RE)

= 1 μg of all-trans-retinol

= 1 μg of all-trans-retinol

= 2 μg of supplemental all-trans-β carotene

= 2 μg of supplemental all-trans-β carotene

= 6 μg dietary β-carotene

= 12 μg dietary β-carotene

= 12 μg other dietary provitamin A carotenoids

= 24 μg other dietary provitamin A carotenoids

See Note 2: 1 μg retinol = 3.33 vitamin A activity from retinol (47); 10 IU β-carotene = 3.33 IU retinol (47). From (1).

3.2. Methods for Dietary Assessment in Human Studies

Instruments that can be used to screen for intakes of fruit and vegetable, percent energy from fat, fiber, added sugar, dairy, calcium, and red meat are available from the National Cancer Institute (9). The fruit and vegetable questionnaire is available as either a quantitative (with portion size questions) or a nonquantitative (without portion size) screening tool. Other questionnaires are also commonly used (10, 11). Obtaining accurate information on the amount of the foods consumed is very problematic, as people are seldom aware of the weight or volume of the foods they consume and often confuse the standard serving size with their usual “helping size” which may be larger. Diet records and recalls can be improved by the use of measuring cups, scoops, food models, or photographs to illustrate portion size. Mixed dishes are also problematic, as the recipe must essentially be deconstructed and analyzed to obtain good estimates of the item’s nutrient contents. All together, recording and recall methods are more difficult than might be assumed. Strengths and weaknesses of dietary recall methods have been reviewed (10, 11). Factors contributing to the imprecision of dietary records or recalls include a tendency for subjects to forget or underreport the foods they have consumed; variations in nutrient intake over time that may not be recalled in the recording period; and limitations in the food composition databases that must be used to translate food consumption into nutrient intake. For vitamin A, certain factors affect the quality of food records. It is known that vitamin A intake in humans has wider day-to-day variations than for some other nutrients; thus many days of food records were needed to attain a good correlation between FFQ data and amounts of usual vitamin A intake (11). A second approach to estimating nutrient intakes is to assess how often certain foods are consumed using a food frequency

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questionnaire, FFQ. Commonly used FFQs have been designed to assess diet and disease risk in large surveys and in multi-ethnic studies (11). Information from FFQs may be used to establish qualitative trends, such as dietary patterns (12), or they may be used in a quantitative manner by approximating not only how often certain foods were consumed but also approximately how much was consumed. Inquires are made about the frequency of intake (daily, weekly, etc.) and are generally focused on those types of foods that are likely to contribute a substantial amount to the total intake of the particular nutrient of interest. In the case of vitamin A, FFQ queries should focus on the intake of green leafy and yellow vegetables, tomatoes and tomato-based products, oranges and orange juice, mangoes, eggs, milk (and whether the milk is vitamin A enriched), certain fortified breakfast cereals that may contain vitamin A, fish, and meat, especially liver. Some FFQs have been simplified to include a shorter list of foods or food groups richest in the nutrient of interest, with the information gained being less precise than from a more detailed or quantitative FFQ. 3.3. Supplement Use as a Factor in Dietary Vitamin A Intake

Vitamin–mineral supplement use is an important component of vitamin A intake in developed countries. Dietary intake and vitamin–mineral supplement use were determined in the HawaiiLos Angeles Multi-Ethnic Cohort study, which includes 215,823 adults who were aged 45 years at baseline in 1993–1996. Murphy et al. (13) concluded that 48% of men and 56% of women reported using a multivitamin supplement at least once weekly for the past year. For vitamin A, the percentage of the population with an adequate intake (comparable to the RDA) increased by 16% for men and 14% for women when supplements were included along with intake from foods, while the prevalence of vitamin A intakes greater than the upper level (UL, 3,000 μg of retinol/day for adults) was 15.6% in men and 15.7% in women. An analysis of data from the 2002 Feeding Infants and Toddlers Study (14) also showed 30% higher vitamin A intakes in supplement users compared to nonusers. Thus, supplement use needs to be factored in to accurately assess vitamin A intake in human studies (10).

3.4. Potential Uses of Dietary Information in Molecular Research

In general, dietary assessment involving vitamin A has not yet received much attention in molecular biological research. However, for other nutrients, such as iron and folate acid, interactions of diet and genotype are now well documented (15–17). As genetic factors modifying vitamin A metabolism are identified, research may turn to determining the interactions of genotype and dietary vitamin A (see Note 3).

3.5. Diets for Research in Animals

In small animals used for research, diet can be used to create conditions that cannot be studied in humans. For vitamin A,

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this includes studies of vitamin A status ranging from deficiency to toxicity, developmental studies, physiological studies of nutrient metabolism in genetic models (expressing transgenes or null deletions), pharmacological studies with natural and synthetic retinoids, and a variety of studies related to preclinical testing. The diets fed to most rodents housed in research facilities are nonpurified diets. In contrast, most research diets are, or should be, purified. The characteristics of these types of diets will be discussed first, and then some recommendations will be proposed for diets appropriate for different types of animal studies. Diets and nutrient recommendations for animals (see Note 4) are almost always expressed in amounts (mass) per weight of diet (e.g., g/kg or g%), not in an amount per day as for humans. 3.5.1. Nonpurified Diets for Animal Studies

The primary ingredients in nonpurified diets come from natural sources. Most nonpurified diets are comprised of a mixture of grains (corn, wheat, barley, sorghum, alfalfa, soybean meal, as examples) and other products (animal or vegetable fats). Feed manufacturers offer a range of such diets, formulated for the growth and reproductive needs of particular species. These diets are suitable for feeding production animals and companion animals as regular feeds and are the typical diet of research animals unless special diets are indicated. Nonpurified diets are formulated to provide at least a minimal amount of all essential nutrients – protein, fat, fiber, and vitamins and minerals – which must fall within certain ranges. They are sometimes classified as open or closed formula. For open formula diets, the composition is made available to the potential user and the diet must be formulated as specified. For closed formula diets, although specified tolerances must be maintained the exact composition of the diet is known only to the manufacturer (18). For research requiring standardization of vitamin A, grainbased diets are not sufficiently uniform over time, nor is the actual vitamin A content of the diet known, other than that it meets a minimal standard. The amount of carotenoids present in grains such as corn and grasses such as alfalfa can vary by strain, season, or geographical origin. Rodents are generally very efficient at converting these carotenoids to retinol within the intestine and thus variations in the amount of provitamin A in the diet can be expected to lead to differences in the vitamin A content of the animal’s vitamin A-storing tissues. Manufacturers may add vitamin A as retinol to nonpurified diets to meet a certain level. A chapter on “Label Review” of the Feed Inspector’s Manual, Association of American Feed Control Officials Inspection and Sampling Committee (19), refers to feed labeling for vitamin A as follows: “Vitamin A, other than precursors of vitamin A, in International Units per pound.” This implies that precursors of vitamin A (carotenoids) in the feed are not counted on

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the product label. In additional, manufacturing allowances can also affect vitamin A content. Manufacturers are allowed to add more than the stated amount of certain nutrients to compensate for “shelf life” (possible loss or deterioration prior to consumption). However, if the diet is carefully stored and is used quickly, the amount of the nutrient is unlikely to have decayed substantially and the amount the animal ingests could be higher than the amount calculated from the product label and the quantity of diet consumed. Manufacturers may also increase the level of micronutrients in diets designed to be autoclaved. A summary of 13 studies on the retention of vitamins after steam autoclaving reported a range of 23–95% retention for vitamin A, with >80% retention in 8 of the studies (20). Thus, the amount lost during autoclaving does not appear to be high. Diets used in transgenic mouse facilities are likely to be of the autoclavable type (21), and thus animals housed in such facilities may be ingesting even higher amounts of vitamin A (and other micronutrients) from nonpurified diets labeled “autoclavable” than even from regular nonpurified diets, which are already high in vitamin A. Overall, the types and amounts of vitamin A present in nonpurified diets are essentially unknown and the use of such diets in research, except for general maintenance of rodent colonies, should be discouraged. 1. Determine if the type of diet fed by the animal facility is appropriate for the study. (If not, see Section 3.6 on purified diets.) 2. Record the diet’s manufacturer, product number, and whether it is described as autoclavable. If the formula is open, maintain a record of the formula for future reference. 3. Include an “Animals” section in the Materials and Methods section of publications resulting from the study, including the animal species, strain, sex, age, diet, and housing conditions. 3.5.2. Purified Diets in Animal Studies

Purified diets, also known as semisynthetic diets, are made of refined ingredients, including isolated proteins, refined sugars and oils, and purified sources of vitamins and minerals. They are formulated to minimize nutrient variability. Purified, fixed-formula diets are typically prepared with a particular type of protein, casein (see Note 5), or whey protein isolated from milk, soy protein, or another protein; a particular oil or fat (corn oil, soybean oil, canola oil, etc., or a known blend); certain carbohydrates (dextrose (glucose), maltose, sucrose, corn starch, or another carbohydrate, and cellulose); and all of the essential vitamins and minerals added in purified form and in exact amounts. Complete mixing is paramount (see Notes 6, 7, and 8). Many researchers

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prefer to purchase custom diets from diet manufacturers who are experienced in the handling of the ingredients and can finish the diet, such as by pelleting it, which is helpful for delivering the diet to the animals using standard caging. Powdered diets can also be fed but require special glass cups and holders to prevent spillage. Purified diets are meant to be stored carefully (e.g., refrigerated or frozen in closed bags or bins to prevent oxidation and light exposure) and used soon after purchase; the amounts of the nutrients they contain should be exactly those specified on the formula sheet. 3.6. AIN-76 and AIN-93 Diets as Classical Purified Diets

Since the 1970s, the American Institute for Nutrition (AIN), now the American Society for Nutrition, has sponsored the testing of purified diets for rodent research. The AIN-76 diet (22) and the slightly modified AIN-76A diet (23) were used extensively by nutritional scientists and others, around the world for two decades. Having a common reproducible diet has greatly aided comparisons among studies. Wise (24) has discussed several practical issues in preparing this diet, including the order of addition of ingredients, which should be understood by researchers planning to prepare animal diets in their laboratory (see Notes 6, 7, and 8). Two new formulations were published in 1993, based on new science and testing in rats for growth and reproduction. AIN-93G (G for growth) was designed for feeding to young animals during rapid growth and for pregnancy and lactation, and AIN-93 M (M for maintenance), for which the optimal protein intake is lower, for feeding to mature animals. As described by Reeves (25), the criteria used for the AIN-93 formulations were the following: (i) the diets can be made from purified ingredients; (ii) they conform to or exceed the nutrient requirements suggested by the NRC, 1978 and 1995 (26, 27); (iii) they can be made from readily available components at a reasonable cost; (iv) the compositions are consistent and reproducible; and (v) the diets can be used over a wide range of applications. Some major differences were made in the formulation of AIN-93G diet, compared with AIN76A diet, to increase the amount of the essential n−3 fatty acid linolenic acid; substitute cornstarch for sucrose to reduce dental caries; reduce the amount of phosphorus to help eliminate the problem of kidney calcification in female rats, which had become apparent with AIN-76; substitute L-cystine for DL-methionine as the amino acid supplement for casein, known to be deficient in the sulfur amino acids; lower the manganese concentration; increase the amounts of vitamin E, vitamin K, and vitamin B12; and add the trace minerals molybdenum, silicon, fluoride, nickel, boron, lithium, and vanadium to the mineral mix. For the AIN-93 M maintenance diet, the amount of fat is reduced to 40 g/kg (4% from 7% in the AIN-93G formula), and the amount of casein to

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140 g/kg from 200 g/kg in the AIN-93G diet, because a lower protein diet was beneficial for maintenance. The energy distribution and ingredients in the AIN-93G diet are listed in Table 17.2. The AIN-93 diets contain 4,000 IU (1,200 μg) retinol/kg of diet, with retinol added to the vitamin mix in the form of gelatinstabilized retinyl palmitate or acetate for greater diet stability.

Table 17.2 Composition of diets used for vitamin A depletion and long-term maintenance of selected vitamin A status in rats AIN-93G growing rodent diet (25) Energy distribution

Modifications to control the level of vitamin Aa Vitamin A deficient

Low marginal

Marginal

Adequate (used as control)

Supplemented

Protein (kcal %)b

20.3

20.3

20.3

20.3

20.3

20.3

Carbohydrate (kcal %)

63.9

63.9

63.9

63.9

63.9

63.9

Fat (kcal %)

15.8

15.8

15.8

15.8

15.8

15.8

Energy density (kcal/g) Final vitamin A concentration:μg retinol/g diet

4.00

4.00

4.00

4.00

4.00

4.00

0

0.35

0.73

4

25, 50, or 100

Coding colors c

(None)

“White”

“Purple”

“Green”

“Pink”

“Gold”

FD&C red dye #40

0

0

0.025

0.05

0.025

FD&C blue dye #1

0

0

0.025

FD&C yellow dye #5

0

0

0.025 0.025

0.025

Ingredients Casein, lactic (g/kg)d

200

200

200

200

200

L-Cysteine (g/kg)

3

3

3

3

3

Corn starch

397.5

397.5

397.5

397.5

397

Maltodextrin

132

132

132

132

132

Sucrose

100

100

100

100

100

Fiber: cellulose, BW220

50

50

50

50

50

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Table 17.2 (Continued) AIN-93G growing rodent diet (25)

Modifications to control the level of vitamin Aa

Soybean oile

70

70

70

70

70

Antioxidant: tertbutylhydroquinone

0.014

0.014

0.014

0.014

0.014

Mineral mix (S10022G)f

35

35

35

35

35

35

Vitamin A-free vitamin mix (V13002)g

0

10

10

10

10

10

Std. vitamin mix (V10037)

10

Vitamin A (retinyl palmitate) concentrate, 500,000 USP units/gh, i

0

0.0027

0.0049

0.0278

0.174, 0.348, or 0.695

Choline bitartrate

2.5

2.5

2.5

2.5

2.5

Total (g)

1000

1000

1000

1000

1000

a Any reputable diet manufacturer can produce the AIN diet or modifications thereof. We have purchased the diets

indicated from Research Diets, Inc. The ingredient product numbers are those of this manufacturer. b Maintenance formula is modified to 14% protein, 73% carbohydrate, and 4% fat (g%) for mature animals. c Food-grade dyes are used to visually code the diets. d Standard casein is used. Vitamin-free (vitamin tested) casein and alcohol-stripped casein are more expensive and were

not found to be necessary since their retinol content is extremely low. e Tocopherol-stripped soybean oil is not necessary unless the diet also must be limited in tocopherol. Soybean oil is

not a source of vitamin A or carotene. f The complete mineral mix is the same as that reported by Reeves (25). Thirty-five grams of this mix is added per kilogram of diet. g The vitamin A-free vitamin mix, per kilogram of mix, contains the following: vitamin D3 (100,000 IU/g), 1.0 g; vitamin E acetate (500 IU/g), 15 g; vitamin K as phylloquinone, 0.075 g; biotin (1%), 2.0 g; cyanocobalamin (0.1%), 2.5 g; folic acid, 0.2 g; nicotinic acid, 3.0 g; pantothenate, calcium, 1.6 g; pyridoxine-HCl, 0.7 g; riboflavin, 0.6 g; thiamin HCl, 0.6 g; powdered sucrose 972.7 g; total: 1 kg. 10 g of this mix is added per kg of diet. [Modified from (25).] h 1 USP unit = 1 IU = 0.55 μg of retinyl palmitate/g of the concentrate. i Diets labeled marginal, adequate, and supplemented diet (25 μg/g diet) were fed to rats for up to 20 months in a long-term aging study. See (30, 31) for plasma and liver vitamin A levels in young, middle-aged, and old rats.

3.7. Liquid Diet with Ethanol

Certain purified diets cannot be prepared in solid form. A commonly used liquid diet used in research on alcohol, and vitamin A–alcohol interactions, is the Lieber–DeCarli diet containing ethanol, 1 kcal/g (36% of kcal) of liquid diet (28). The ingredients for this diet can be purchased as a solid mix, minus

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ethanol, and blended in the laboratory to contain 36% of calories (or a modified level) from ethanol. The vitamin A content is 6,000 IU/l (1,800 μg/l). Given that the AIN-93 solid diet has an energy density of ∼4 kcal/g and contains 1.2 μg retinol/g, equal to 0.3 μg retinol/kcal, the Lieber–DeCarli diet with 1.8 μg retinol/kcal provides more vitamin A per calorie. 3.8. Custom Modifications of Dietary Vitamin A to Control Vitamin A Status

Custom diets are purified diets tailored to the needs of the users. Vitamin A-deficient diets and diets with different, specified levels of vitamin A fall into this category. In the 1980s, our laboratory began studies of vitamin A depletion and repletion, and later of a range of vitamin A status, using diets first based on AIN-76 and later on AIN-93G, with AIN-93 M used for mature animals in a long-term study of aging (29–31). Table 17.2 provides a summary of the AIN-93G diet and its modification to obtain five different “levels” of vitamin A status in rats, ranging from “low marginal” to “supplemented.” We increased the concentration of vitamin A in the adequate diet to 4 μg/g (from 1.2 in the AIN-93 formula) after finding that plasma retinol and liver retinyl esters were relatively low if the animals were fed AIN-93 diet for several months. Rats fed the vitamin A-adequate diet with 4 μg retinol/g had liver total retinol concentrations of 362 nmol/g (104 μg/g) at 3 months of age (31), within the range of liver total retinol considered adequate in humans (32).

3.9. Methods for Induction of Vitamin A Deficiency

The time required to induce vitamin A deficiency depends on the level of preexisting vitamin A storage and the animal’s rate of growth. 1. To obtain a reproducible time course in the development of vitamin A deficiency in rats (applicable also to mice), begin by feeding vitamin A-deficient diet (Table 17.2) to the lactating mothers of nursling pups; this significantly reduces the transfer of vitamin A from mother to pups (33, 34). 2. Wean the pups (3 weeks of age) onto the same vitamin Adeficient diet or onto a vitamin A-containing diet according to the study’s design. 3. Weigh the animals weekly. Vitamin A-deficient animals will start to show reduced weight gain, although this tends to follow rather than precede biochemical depletion. 4. To assess the progression of vitamin A deficiency, one must measure plasma retinol. A first measurement is suggested at 5–6 weeks and a second at 7–8 weeks. If the study is to continue beyond 8 weeks, physical signs may become apparent (see Note 9). Blood can be collected from the tail, retro-orbital sinus, or heart, according to the investiga-

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tor’s approved animal protocol. Serum or plasma retinol is determined by extraction and chromatography (an HPLC method with UV detection is preferred (35)). 5. Animals purchased as weanlings from animal suppliers cannot be expected to follow this time course of depletion, due to their accumulation of vitamin A before shipping (see Note 10). 6. Mice are well known to be difficult to deplete of vitamin A (see Note 11). A deficient state with impaired immune response was produced in mice by vitamin A-deficient diet (36) in a manner similar to the rat studies, above. In studies of development, Morriss-Kay and Sokolova (37) noted a low incidence of mild effects is in the first litter of mice fed a vitamin A-deficient diet, with a higher incidence of more severe effects observed in the second litter, reflecting a greater degree of maternal deficiency during the second pregnancy. Clagett-Dame and coworkers developed a dietary strategy to produce vitamin A deficiency at specific times later in pregnancy (38). 7. Female mice or rats are first made deficient in vitamin A, then mated with vitamin A-adequate males. 8. Pregnant females are fed vitamin A-deficient diet supplemented with tRA/g (12 μg/g of diet, approximately equal to 230 μg tRA/rat/day, or an equivalent oral supplement daily, due to the rapid turnover of tRA) to assure normal fetal development to mid-pregnancy. 9. The tRA is withdrawn at predetermined times that depend on the developmental outcomes to be assessed. This approach (38) enabled the investigators to achieve tight control over the timing of the deficiency state of the animal, owing to the rapid turnover of RA. They observed gross abnormalities, including defects in eye development, in rat embryos at day E12.5. 10. For mice, we have fed the same vitamin A-deficient and vitamin A-adequate diets shown in Table 17.2, with similar effects on plasma and liver vitamin A as in rats. However, external signs of vitamin A deficiency (body condition) were not as readily apparent in mice as in rats. 3.10. Diets Differing in Macronutrient Content

The diets shown in Table 17.2 are all equal in energy density. When protein, carbohydrate, or fat contents are altered, the energy density of the diet is also altered. Thus the amounts of the vitamin and mineral mixes also must be adjusted if a constant intake per kcal is to be maintained. The diets formulated by Clinton and Visek (39) provide an excellent example of the

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correct approach to purified diets that differ in protein or fat, while maintaining micronutrients at a constant level per kilocalorie. These or similar diets could be modified for studies of vitamin A metabolism related to obesity, metabolic syndrome, etc., to assure that micronutrient intakes are comparable across all diet groups and do not differ due to changes in the macronutrient composition of the diet. 3.11. Practical Considerations for Diet Preparation and Storage

Nonpurified diets are formulated with a low moisture level to improve stability. The diet is typically extruded in the form of hard pellets, which simplifies the feeding of animals and generally reduces waste. Diet is often added to the cage unit in amounts that will last several days. By contrast, purified diets are more labile and should be stored in a closed container in a cold room, kept dark, and protected from oxidation. Diets with a very high fat content should be kept frozen. These diets should be fed in amounts that animals will consume in a day or two and then replaced as needed.

3.12. Care of Incisors

Purified diets of the AIN type can be prepared in pelleted form. However, some diet formulas, like very low-fat diets, are difficult to form into pellets and thus must be fed in powdered form using a glass cup that fits inside the animal’s cage. Since the incisors of rodents grow continuously, care should be taken to observe the teeth of animals fed soft diets and to trim the teeth as necessary (40).

4. Notes 1. Controlling the diet in human studies is very expensive and also poses logistical challenges. All food, and often lodging, must be provided, and subjects are usually compensated at the completion of the study for their successful participation. Logistically, a human feeding study requires using a Metabolic Kitchen (General Clinical Research Center, Clinical and Translational Science facility, or the like) and having highly trained staff, including a research dietitian, food preparation specialists skilled in preparing standardized meals, a study coordinator, and dining room staff to oversee subjects during meals. Controlled feeding studies require approval by an Institutional Review Board and, depending on the intervention planned, may require a Data Safety Monitoring Board. The web site of the National Association of Bionutritionists has information on “WellControlled Diet Studies in Humans: A Practical Guide to Design and Management,” which addresses many practical

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problems, covering design, implement, and management of human diet research studies (41). 2. It is important to determine that the units of vitamin A obtained from different sources are comparable, or to convert them to a common unit. The RAE (see Section 3.1) is the current unit for human vitamin A intake. Note that β-carotene in supplements (oily solution) is converted to retinol three times more efficiently than is β-carotene from foods (present in food matrix), and the conversions shown in Table 17.1 reflect this. 3. Examples include that Berson (42) has noted that nutritional approaches have been effective in treating certain diseases of the retina, for example, the night blindness associated with Sorsby fundus dystrophy can be reversed over the short term with vitamin A and has concluded that “risk-factor analyses of well-defined populations followed over time with food frequency questionnaires in conjunction with careful assessments of visual function may reveal other dietary constituents that can modify the course of degenerative diseases of the retina.” The proven benefit of antioxidant supplementation, including carotenoids, for age-related macular degeneration in the Age-Related Eye Disease Study (AREDS) also suggests that the interaction of diet or supplementation and genetic risk factors should be examined more closely (43). 4. Nutrient recommendations for animals [such as issued by the National Research Council (27)] are expressed in amounts (mass) per weight of diet (e.g., g/kg or g%), whereas in contrast intakes for humans are expressed in amount per day. To estimate how much of a given nutrient an animal will consume or has consumed, it is necessary to measure food intake by providing animals with a weighed portion of food and weighing back the unconsumed portion (include a full 24-h day or two full days, as rodents are night eaters). If approximate intakes are determined in a preliminary study, the amount of diet needed for a larger study can be estimated. Keenan et al. (44) have argued for feeding rodents a calorically restricted diet, equal to 70–75% of ad libitum intake, to prevent the development of overweight, diabetes, tumors, and reduced survival, in sedentary rodents. Such a strategy may be useful in longterm studies of vitamin A in rodents. 5. We have not found it necessary to use alcohol-extracted casein (which is more expensive than regular casein), as the vitamin A content of non-extracted casein is extremely low. 6. If a purified diet is to be mixed in-house, first determine if there is a large diet-mixing facility that can be used. The mineral and the vitamin “premix” contains very

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small amounts of certain minerals, such as copper, trace elements, and vitamins such as cyanocobalamin, which are present in minute amounts. Distributing these minor components evenly in the bulk diet is critical. Dry ingredients should be handled separately from fat-soluble ingredients (see Note 6). The major component of the premix is a dry component (powdered sucrose, dextrose, cornstarch, etc.) to act as a reservoir/binder for the vitamins and minerals. The dry vitamins and minerals are to be weighed carefully, using an analytical balance, and added to a small portion (e.g., 10%) of the powdered sucrose, then mixed thoroughly. A kitchen-type food mixer with a wire whisk blade is useful; use a slow speed and use a spatula to aid in mixing. Do not use a blender that will aerate the mix. After all the dry ingredients are well mixed with the small portion of the dry component, gradually add the rest of the dry component in several small additions and blend thoroughly after each addition. Store the mix in a well-covered container (to prevent exposure to light and oxygen) at 4◦ C or lower. 7. Fat-soluble vitamins, especially vitamin A and vitamin E, pose special problems for blending well into the diet. If the vitamin A is in the form of an oil (retinyl palmitate), it is recommended to make a concentrated premix using a portion, e.g., 5%, of oil to be added to the diet. The appropriate amount of vitamin A in oil is then added to the remaining portion of the oil, mixed, and the oil is then added to the dry ingredients of the diet. Vitamin A can also be obtained in a concentrated gelatin-stabilized form. This too is added to the oil and blended thoroughly prior to mixing the oil with the dry ingredients. We prefer to add oily solutions into the mix and into the final diet by weight rather than by using cylinders or pipettes, due to incomplete drainage of oily solutions from these containers. 8. To avoid spillage, do not prepare more than one-quarter of the amount of diet that can be blended at once, e.g., if the size of the mixing bowl is 12 kg, no more than 3 kg of diet should be mixed at a time. Spillage before the diet has completely mixed will cause deviations from the desired formula. 9. We have found that growing rats fed the vitamin Adeficient diet have become biochemically depleted of vitamin A (liver total retinol concentration of <5 μg/g tissue; plasma retinol <0.3 μM) by 7 weeks of age for males and 8 weeks of age for females. External signs of vitamin A deficiency become apparent approximately a week later for both sexes. Signs include slowing or cessation

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of growth, somewhat rough fur, and occasionally a red crust (porphyrins from dried tears) around the eyes and/or nares. A staggered gait and locomotor impairment are seen rather late in the course of vitamin A deficiency. Neuromuscular dysfunction has been attributed to changes in striatal neurons, implicating vitamin A in the maintenance of basal ganglia motor function in the adult rat brain (45). Internally, nearly all of the animal’s visceral fat is depleted and thus the adrenal glands can be easily seen in the remnants of perirenal fat. The thymus may be smaller and less white in color. 10. If weanling rats or mice are purchased from suppliers and then fed a vitamin A-deficient diet, the time course of depletion is likely to be longer and not easily predictable, as the young animals will have received more vitamin A from their dams, and they most likely will have begun to consume crumbs of their mother’s nonpurified diet before weaning. Thus these animals will have accumulated vitamin A in their tissues prior to the time of shipment. 11. One reason for the difficulty in depleting mice of vitamin A may be related to the practice of animal research facilities, especially those housing transgenic animals, to feed an autoclavable nonpurified diet with extra vitamin A (see above). Another reason could be related to the efficiency of recycling retinoids. Both mice and rats practice coprophagy (in which feces are consumed), which may serve to conserve retinol even after a vitamin A-deficient diet has been imposed. Young mice are said to practice “vigorous coprophagy” (46), which may increase the recycling and delay the onset of deficiency. The type of housing used for mice (typically shoebox-type cages with multiple animals per cage) may enable more recycling of feces, although this is a conjecture. Hanging wire cages, sometimes used for rats, reduce but do not totally eliminate coprophagy, but this type of caging is seldom used for mice.

Acknowledgments Support: NIH grants CA-90214; DK-41479. References 1. Institute of Medicine. (2001) Dietary Reference Intakes for Vitamin A, Vitamin K, Arsenic, Boron, Chromium, Copper, Iodine, Iron, Manganese, Molybdenum, Nickel,

Silicon, Vanadium, and Zinc, National Academy Press, Washington. 2. Haskell, M.J., Mazumder, R.N., Peerson, J.M., Jones, A.D., Wahed, M.A.,

312

3.

4.

5.

6.

7.

8.

9.

10.

11.

12.

13.

Ross Mahalanabis, D., Brown, K.H. (1999) Use of the deuterated-retinol-dilution technique to assess total-body vitamin A stores of adult volunteers consuming different amounts of vitamin A. Am. J. Clin. Nutr. 70, 874–780. Tang, G.W., Qin, J., Dolnikowski, G.G., Russell, R.M. (2003) Short-term (intestinal) and long-term (postintestinal) conversion of beta-carotene to retinol in adults as assessed by a stable-isotope reference method. Am. J. Clin. Nutr. 78, 259–266. Ahmad, S.M., Haskell, M.J., Raqib, R., Stephensen, C.B. (2008) Men with low vitamin A stores respond adequately to primary yellow fever and secondary tetanus toxoid vaccination. J. Nutr. 138, 2276–2283. Hargreaves, M.K., Baquet, C., Gamshadzahi, A. (1989) Diet, nutritional status, and cancer risk in American blacks. Nutr. Cancer 12, 1–28. Fontham, E.T.H., Pickle, L.W., Haenszel, W., Correa, P., Lin, Y., Falk, R.T. (1988) Dietary vitamins A and C and lung cancer risk in Louisiana. Cancer 62, 2267–2273. Paganini-Hill, A., Chao, A., Ross, R.K., Henderson, B.E. (1987) Vitamin A, β-carotene, and the risk of cancer: A prospective study. JNCI. 79, 443–448. Ross, A.C. (1994) Vitamin A and cancer. In: Carroll, K.K., Kritchevsky, D. (eds.), Nutrition and Disease Update, Cancer, AOCS Press, Champaign, IL, pp. 27–109. National Cancer Institute. (2009) Short Dietary Assessment Methods. http:// riskfactor.cancer.gov/diet/ and http:// riskfactor.cancer.gov/diet/screeners/ (last accessed April 22, 2010). Smiciklas-Wright, H., Mitchell, D.C., Ledikwe, J.H. (2007) Dietary intake assessment: Methods for adults. In: Berdanier, C.D., Dwyer,J.T., Feldman, E.B. (eds.), Handbook of Nutrition and Food, 2nd ed, CRC Press, Boca Raton, FL, pp. 493–508. Reeves, R.S., Pace, P.W. (2007) Use of food frequency questionnaires in minority populations. In: Berdanier, C.D., Dwyer, J.T., Feldman, E.B. (eds.), Handbook of Nutrition and Food, CRC Press, Boca Raton, FL, pp. 509–528. Haskell, M.J., Lembcke, J.L., Salazar, M., Green, M.H., Peerson, J.M., Brown, K.H. (2003) Population-based plasma kinetics of an oral dose of [2H4]retinyl acetate among preschool-aged, Peruvian children. Am. J. Clin. Nutr. 77, 681–686. Murphy, S.P., White, K.K., Park, S.Y., Sharma, S. (2007) Multivitaminmultimineral supplements’ effect on total

14.

15.

16.

17. 18. 19.

20.

21.

22.

23.

24. 25. 26.

nutrient intake. Am. J. Clin. Nutr. 85(suppl), 280S–284S. Briefel, R., Hanson, C., Fox, M.K., Novak, T., Ziegler, P. (2006) Feeding infants and toddlers study: Do vitamin and mineral supplements contribute to nutrient adequacy or excess among US infants and toddlers? J. Am. Diet. Assoc. 106, S52–S65. Mithen, R. (2007) Effect of genotype on micronutrient absorption and metabolism: A review of iron, copper, iodine and selenium, and folates. Int. J. Vitam. Nutr. Res. 77, 205–216. Powers, H.J. (2005) Interaction among folate, riboflavin, genotype, and cancer, with reference to colorectal and cervical cancer. J Nutr. 135(Suppl), 2960S–2966S. Friso, S., Choi, S.W. (2005) Gene-nutrient interactions in one-carbon metabolism. Curr. Drug Metab. 6, 37–46. Newberne, P.M., Fox, J.G. (1980) Nutritional adequacy and quality control of rodent diets. Lab. Anim. Sci. 30, 352–365. Feed Inspector s Manual. (2000) Chapter 4, Label Review, 2nd ed., Association of American Feed Control Officials Inspection and Sampling Committee. http://www.aafco.org/Portals/0/Public/ COMPLETE_INSPECTORS_MANUAL. pdf (last accessed April 22, 2010). Tobin, G., Stevens, K.A., Russell, R.M. (2007) Nutrition. In: Fox, J.G., Barthold, S.W., Davisson, M.T., Newcomer, C.E., Quimby, F.W., Smith, A.L. (eds.), The Mouse in Biomedical Research, Academic, New York, pp. 321–384. Lipman, N.S. (2007) Design and analysis of research facilities for mice. In: Fox, J.G., Barthold, S.W., Davisson, M.T., Newcomer, C.E., Quimby, F.W., Smith, A.L. (eds.), The Mouse in Biomedical Research, 2nd ed. Academic, New York. American Institute of Nutrition. (1977) Report of the American Institute of Nutrition ad hoc committee on standards for nutritional studies. J. Nutr. 107, 1340–1348. American Institute of Nutrition. (1980) Second report of the ad hoc committee on standards for nutritional studies. J. Nutr. 110, 1726. Wise, A. (ed.). (1991) Diet Formulation and Experimental Design for Laboratory Animal Studies, CRC Press, Boca Raton, FL. Reeves, P.G. (1997) Components of the AIN-93 diets as improvements in the AIN76A diet. J. Nutr. 127, 838S–841S. National Research Council. (1978) Nutrient Requirements of Laboratory Animals, 3rd ed. National Academies Press, Washington, DC.

Diet in Vitamin A Research 27. National Research Council. (1995) Nutrient Requirements of Laboratory Animals, 4th ed. National Academies Press, Washington, DC. 28. Lieber, C.S., DeCarli, L.M. (1982) The feeding of alcohol in liquid diets: Two decades of applications and 1982 update. Alcohol Clin. Exp. Res . 6, 523–531. 29. Zolfaghari, R., Ross, A.C. (1995) Chronic vitamin A intake affects the expression of mRNA for apolipoprotein A-I, but not for nuclear retinoid receptors, in liver of young and aging Lewis rats. Arch. Biochem. Biophys. 323, 258–264. 30. Dawson, H.D., Li, N.Q., DeCicco, K.L., Nibert, J.A., Ross, A.C. (1999) Chronic marginal vitamin A status reduces natural killer cell number and function in aging Lewis rats. J. Nutr. 129, 1510–1517. 31. Dawson, H.D., Yamamoto, J., Zolfaghari, R., Rosales, F., Dietz, J., Shimada, T., Li, N.Q., Ross, A.C. (2000) Regulation of hepatic vitamin A storage in a rat model of controlled vitamin A status during aging. J. Nutr. 130, 1280–1286. 32. Olson, J.A. (1984) Serum level of vitamin A and carotenoids as reflectors of nutritional status. J. Natl. Cancer Inst. 73, 1439–1444. 33. Davila, M.E., Norris, L., Cleary, M.P., Ross, A.C. (1985) Vitamin A during lactation: Relationship of maternal diet to milk vitamin A content and to the vitamin A status of lactating rats and their pups. J. Nutr. 115, 1033–1041. 34. Akohoue, S.A., Green, J.B., Green, M.H. (2006) Dietary vitamin A has both chronic and acute effects of vitamin A indices in lactating rats and their offspring. J. Nutr. 136, 128–132. 35. Ross, A.C. (1986) Separation and quantitation of retinyl esters and retinol by highperformance liquid chromatography. Meth. Enzymol. 123, 68–74. 36. Smith, S.M., Levy, N.L., Hayes, C.E. (1987) Impaired immunity in vitamin A deficient mice. J. Nutr. 117, 857–865.

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37. Morriss-Kay, G.M., Sokolova, N. (1996) Embryonic development and pattern formation. FASEB J. 10, 961–968. 38. Clagett-Dame, M., DeLuca, H.F. (2002) The role of vitamin A in mammalian reproduction and embryonic development. Annu. Rev. Nutr. 22, 347–381. 39. Clinton, S.K., Imrey, P.B., Alster, J.M., Simon, J., Truex, C.R., Visek, W.J. (1984) The combined effects of dietary protein and fat on 7,12-dimethylbenz(a)anthraceneinduced breast cancer in rats. J. Nutr. 114, 1213–1223. 40. Donnelly, T.M., Brown, C.J. (2009) Rodent Husbandry and Care. http://www.veterinary partner.com/Content.plx?P=A&A=2497& S=4 (last accessed April 22, 2010). 41. National Organization of Bionutritionists. Well-Controlled Diet Studies in Humans: A Practical Guide to Design and Management. http://www.nabconnection.org/ (last accessed April 22, 2010). 42. Berson, E.L. (2000) Nutrition and retinal degenerations. Int. Ophthalmol. Clin. 40, 93–111. 43. Coleman, H., Chew, E. (2007) Nutritional supplementation in age-related macular degeneration. Curr. Opin. Ophthalmol. 18, 220–223. 44. Keenan, K.P., Laroque, P., Dixit, R. (1998) Need for dietary control by caloric restriction in rodent toxicology and carcinogenicity studies. J. Toxicol. Environ. Health B Crit. Rev. 1, 135–148. 45. Cartaa, M., Stancampianoa, R., Troncib, E., Colluc, M., Usiellod, A., Morellib, M., Faddaa, F. (2006) Vitamin A deficiency induces motor impairments and striatal cholinergic dysfunction in rats. Neuroscience 139, 1163–1172. 46. Brown, C.J., Donnelly, T.M. (2004) Rodent husbandry and care. Vet. Clin. North Am. Exot. Anim. Prac. 7, 201–225. 47. World Health Organization. (1966) WHO Expert Committee on Biological Standardization Eighteenth Report. Technical report series, No. 329, Geneva.

Chapter 18 Experimental Approaches to the Study of A2E, a Bisretinoid Lipofuscin Chromophore of Retinal Pigment Epithelium Janet R. Sparrow, So Ra Kim, and Yalin Wu Abstract Bisretinoid lipofuscin compounds that accumulate in retinal pigment epithelial (RPE) cells are implicated in the pathogenesis of some forms of macular degeneration. In the development of approaches to the amelioration of retinal disorders characterized by enhanced RPE lipofuscin formation, attention is being given to therapies that reduce the production of these damaging pigments. An understanding of the biosynthetic pathways by which these molecules form is essential to the development of these therapies. Thus methods for studying the biosynthesis of these compounds are presented. A tissue culture model is also described whereby a human RPE cell line that is otherwise devoid of bisretinoid lipofuscin compounds is employed and synthesized A2E is delivered to the cells. This approach allows for a population of RPE cells that have accumulated the lipofuscin fluorophore A2E in addition to A2E-free cells. Key words: A2E, A2PE, all-trans-retinal, all-trans-retinal dimer, bisretinoids, retinal pigment epithelium, visual cycle, vitamin A.

1. Introduction Vitamin A aldehyde-derived compounds accumulate as lipofuscin in retinal pigment epithelial (RPE) cells with time and have been linked to some forms of macular degeneration, including juvenileonset recessive Stargardt disease, dominant Stargardt-like maculopathy, Best vitelliform macular degeneration, and age-related macular degeneration. These bisretinoid constituents of lipofuscin are unique to retinal pigment epithelial cells and include the compounds A2E, a C13–C14 Z-isomer of A2E (isoA2E), and other minor cis-isomers (Fig. 18.1) (1, 2). A2E is a pyridinium H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_18, © Springer Science+Business Media, LLC 2010

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Fig. 18.1. Structures and absorbance maxima (λmax ). (a) Structures and λmax of A2E and the biosynthetic precursors dihydropyridinium-A2PE and A2PE. The structure of A2E is that of a pyridinium bisretinoid. (b) Structures and λmax of the all-trans-retinal dimer series of compounds: all-trans-retinal dimer-PE, all-trans-retinal dimer-E, and all-transretinal dimer. (c) Structure and λmax of all-trans-retinol. Absorbance maxima of bischromophores in (a) and (b) can be assigned to the shorter and longer side arms. All-trans-retinol (c) has an absorbance maximum at 325 nm that reflects an extended conjugation system consisting of a polyene chain of four double bonds with a fifth conjugated olefin in the β-ionone ring. For the bischromophore dihydropyridinium-A2PE, an absorbance maximum of ∼330 nm is generated by the same conjugation system along the short arm. Similarly, absorbance maxima of A2E and A2PE originate from the shorter and longer side arms extending from the pyridinium ring. Note that the short arm of all-trans-retinal dimerPE and all-trans-retinal dimer-E has only four double bonds (three along the polyene chain and one in the β-ionone ring); accordingly, the absorbances generated from the short arms of these compounds are blue-shifted relative to the absorbances generated from the short arm of A2E, dihydropyridinium-A2PE, and A2PE. For all-trans-retinal dimer-PE and all-trans-retinal dimer-E, the absorbance generated from the long arm exhibits a bathochromic shift (red shift) due to protonation of the imine functional group (–C=N–).

bisretinoid conjugate named because it can be synthesized in the laboratory from vitamin A aldehyde and ethanolamine in a 2:1 ratio. In addition to A2E and its isomers, other bisretinoid constituents of RPE lipofuscin are A2-dihydropyridinephosphatidylethanolamine (A2-DHP-PE) (3) and those pigments generated via the condensation of two all-trans-retinal including all-trans-retinal dimer and the related conjugates all-trans-retinal dimer-phosphatidylethanolamine (all-trans-retinal dimer-PE) and all-trans-retinal dimer-ethanolamine (all-trans-retinal dimer-E). The levels of all of these bisretinoid fluorophores are particularly elevated in Stargardt disease and it is notable that the fold increase in atRAL dimer-PE is greater than that of A2E (4, 5). For all of these chromophores of RPE lipofuscin, absorbance maxima in the visible range of the spectrum are conferred by the extended conjugation systems located along the vitamin A aldehyde-derived

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side arms (in particular the long arm) of these molecules together with double bonds within the six-membered rings (A2E: λmax 338, 439; iso-A2E: λmax 337, 428; all-trans-retinal dimer: λmax 290, 432; all-trans-retinal dimer-E and all-trans-retinal dimer-PE: λmax 290, 510) (Fig. 18.1). The additional red shift to 510 nm in the case of all-trans-retinal dimer-PE and all-trans-retinal dimerE occurs due to protonation of the Schiff base nitrogen (4, 5) (Fig. 18.1). Although A2E/isoA2E and the all-trans-retinal dimer series of compounds are all bisretinoid compounds, differences in the properties of these molecules are of particular interest. For example, A2E/isoA2E are pyridinium salts containing a quaternary amine nitrogen that confers a permanent positive charge on the head group. A2E cannot be deprotonated or reprotonated (6). Conversely, all-trans-retinal dimer-PE and all-transretinal dimer-E are dimers of all-trans-retinal attached to phosphatidylethanolamine or ethanolamine, respectively, via a Schiff base linkage that is protonated (4, 5). By recording changes in the absorbance of these compounds, we demonstrated that the protonation state of the imine nitrogen is pH dependent (5). Several considerations (5) indicate that when housed in lysosomes (pH ∼ 5) all-trans-retinal dimer-PE and all-trans-retinal dimer-E exist largely in the protonated Schiff base form. Yet, since deprotonation followed by Schiff base hydrolysis of all-trans-retinal dimerPE/E generates unconjugated all-trans-retinal dimer and given that we detect both protonated all-trans-retinal dimer-PE/E and unprotonated unconjugated all-trans-retinal dimer in extracts of RPE lipofuscin (4, 5), conditions in the lysosome likely permit both protonated and unprotonated forms. Importantly, the relative levels of these pigments can be altered by a change in lysosomal pH. Adverse effects of lipofuscin pigments on the RPE cell are likely attributable, in large part, to two properties of the bisretinoid compounds: amphiphilic structures that confer detergent-like activity and photooxidative processes that are initiated by these compounds (7, 8). It is well known that A2E when irradiated at an excitation maximum in the blue region of the spectrum serves as a photosensitizer generating various reactive forms of oxygen with singlet oxygen adding to A2E at carbon– carbon double bonds along the side arms of the molecule (9–13). The reactive species generated within photooxidized A2E include endoperoxides, epoxides, and furanoid moieties that have been identified in hydrophobic extracts of human RPE and Abca4–/– mouse eyecups and that likely account for the adverse effects of A2E photoreactivity (11). It is notable that unconjugated alltrans-retinal dimer is an even more efficient generator of singlet oxygen than is A2E and the all-trans-retinal dimer series of compounds are also more efficient quenchers of singlet oxygen (5).

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Differences in electron distribution within the molecules likely account for the greater susceptibility to oxidation exhibited by all-trans-retinal dimer-PE/E, and the photooxidative processes associated with these compounds are likely to be significant to an understanding of the damaging effects of lipofuscin on RPE cells. We previously proposed that A2E forms in vivo via reactions between the membrane phospholipid phosphatidylethanolamine (PE) and all-trans-retinal that forms upon light-mediated isomerization of 11-cis-retinal (6, 14, 15) (Fig. 18.2). We confirmed by mass spectrometry that the compound formed by this reaction is N-retinylidene-phosphatidylethanolamine (NRPE) (15) (Figs. 18.2 and 18.3), the Schiff base conjugate that is

Fig. 18.2. Biosynthetic pathway of A2E. All-trans-retinal generated by light-mediated isomerization of 11-cis-retinal reacts with phosphatidylethanolamine (PE) to generate the Schiff base adduct N-retinylidene-phosphatidylethanolamine (NRPE). After [1,6]-proton tautomerization, reaction with a second molecule of all-trans-retinal and 6π-electrocyclization, a phosphatidyl-dihydropyridinium bisretinoid (dihydropyridinium-A2PE) is formed. Dihydropyridinium-A2PE would undergo a 1,3-H shift and hydrogen atom elimination to give rise to A2-DHP-PE or eliminate two hydrogens to form A2PE, a phosphatidyl-pyridinium bisretinoid. Following aromatic autooxidation, A2PE is generated. Hydrolysis of the phosphate ester of A2PE by phospholipase D yields A2PE. Under room light, A2E and isoA2E reach photoequilibrium at a ratio of 4:1.

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Fig. 18.3. HPLC monitoring of biomimetic synthesis of A2PE and A2E. (a) Synthesis of A2E with all-trans-retinal and ethanolamine as starting materials. Incubation for 7 days. In this reaction mixture ethanolamine was substituted for phosphatidylethanolamine; the latter would be the reactant in vivo. Dihydropyridinium-A2E (A2E-H2 ) is detected as an intermediate of this biosynthetic reaction. Oxidation of the latter intermediate yields A2E. (b) Biomimetic synthesis of lipofuscin pigments using all-trans-retinal and phosphatidylethanolamine as starting materials. Incubation for 7 days. Compounds generated within the reaction mixture include N-retinylidene-phosphatidylethanolamine (NRPE), A2-dihydropyridine-phosphatidylethanolamine (A2-DHP-PE), and A2PE, known intermediates of the A2E biosynthetic pathway. Also generated are compounds of the alltrans-retinal dimer series: all-trans-retinal dimer (atRALdi) and all-trans-retinal dimerPE (atRALdi-PE). (c) A2E is generated from A2PE by phospholipase D (PLD)-mediated hydrolysis. A2PE was incubated in the absence (left) and presence (right) of the lysosomal enzyme PLD for 3 h. Reverse-phase C18 (a, b) and C4 (inset in b, c) columns and monitoring at 490 (a, b) and 430 nm (c).

likely the ligand for ABCA4/ABCR, the photoreceptor-specific ATP-binding cassette transporter (16–21). Mutations in the gene encoding ABCA4/ABCR are responsible for recessive

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Stargardt disease (22). We also showed that as proposed (6), reaction with a second molecule of all-trans-retinal leads to the formation of a phosphatidyl-dihydropyridinium molecule (dihydropyridinium-A2PE) (Figs. 18.1, 18.2, and 18.3) (23). Specifically by quantum chemical simulation we calculated that the UV–visible absorbance spectrum of dihydropyridinium-A2PE exhibits maxima at ∼494 and 344 nm and in an A2E biomimetic reaction mixture we identified a compound exhibiting similar absorbance maxima (λmax 490, 330 nm) and having the expected mass (dihydropyridinium-A2E was used as a model of dihydropyridinium-A2PE in these experiments) (23). Oxidative aromatization of dihydropyridinium-A2PE with the elimination of two hydrogens yields A2PE, a phosphatidyl-pyridinium bisretinoid. Mass spectrometry served to corroborate the structure of A2PE and also established that this compound was the immediate precursor of A2E that is detected in photoreceptor outer segments. More recently we have described an additional pathway by which the elimination of one hydrogen from dihydropyridinium-A2PE rather than two can lead to formation of the stable uncharged dihydropyridine compound A2-DHP-PE (Fig. 18.2). Experiments demonstrating the release of A2E upon phospholipase D-mediated removal of the phosphatidic acid of A2PE demonstrated that the biosynthetic pathway by which A2E forms includes the hydrolytic activity of RPE lysosomes (14, 15, 24) (Fig. 18.3). A2E is housed in the lysosomal compartment of the RPE cell (25), but further degradation of A2E does not occur probably because the unusual structure of this pigment leaves it unrecognizable by the wide variety of acid hydrolases that constitute the enzymes of lysosomes. This would also be the case for the all-trans-retinal dimer series of compounds.

2. Materials 2.1. Biomimetic Synthesis of A2E with Detection of the Intermediate DihydropyridiniumA2PE and the Immediate Precursor A2PE

1. All-trans-retinal, 1,2 dipalmitoyl-sn-glycero-3-phosphoethanolamine, ethanolamine, and phospholipase D (PLD) from Streptomyces chromofuscus (2,000 units/mg) are obtained from Sigma-Aldrich. 2. HPLC-grade acetonitrile, trifluoroacetic acid, chloroform, and methanol are obtained from Fisher Scientific, Fair Lawn, NJ. 3. MOPS buffer (1X): 4.18 g 3-(N-morpholino) propanesulfonic acid (MOPS); 0.68 g sodium acetate; 2 ml 0.5 M

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EDTA; add H2 O to 1,000 ml and dissolve. Store in the dark at 4◦ C. Not autoclavable. R 4. An Atlantis dC18 column (3 μm, 4.6 × 150 mm, Waters, R C4 column (5 μm, 3.9 × 150 mm, USA) and a Delta Pak Waters, USA) are employed.

2.2. Constructing a Cell Culture Model

The tissue culture model which we developed and which is described later provides for an experimental design not possible in vivo: a population of RPE cells that have accumulated the lipofuscin fluorophore A2E and a population of RPE cells devoid of A2E. This culture model allows RPE cells to accumulate A2E in vitro; levels of A2E accrued can be compared to that present in the eye (10, 26). 1. An adult human RPE cell line (ARPE-19 cells) is obtained from American Type Culture Collection, Manassas, VA (see Note 7). 2. Dulbecco’s modified Eagle’s medium (DMEM) with Lglutamine is obtained from Fisher Scientific (Pittsburg, PA); MEM nonessential amino acid solution, fetal bovine serum, and gentamicin sulfate are obtained from Invitrogen (Carlsbad, CA). 3. Eight-well plastic chamber slides (Lab-Tek) are obtained from Nunc, (Naperville, IL) or clear culture inserts (0.4 μm pore, 12 mm diameter inserts, polyester membrane, Transwell) available from Corning Costar, Corning, NY). 4. LysoTracker Red DND-99, MitoTracker Red CM-H2Xros, and Dead Red nuclear stain are purchased from Invitrogen/Molecular Probes, Carlsbad, CA.

3. Methods 3.1. Biomimetic Synthesis of A2E with Detection of the Intermediate DihydropyridiniumA2PE and the Immediate Precursor A2PE

The biomimetic synthesis of A2E can be studied using alltrans-retinal and phosphatidylethanolamine as starting materials, these compounds being the precursors of A2E in vivo. Alternatively, since condensation of ethanolamine with all-transretinal is more facile and thus affords more reaction product, ethanolamine can be used in place of phosphatidylethanolamine. Substitution with ethanolamine is also practical since the phospholipid moiety of phosphatidylethanolamine does not contribute to absorbances above 250 nm. With phosphatidylethanolamine as starting material, the dihydro intermediate is a phosphatidyldihydropyridinium bisretinoid (dihydropyridinium-A2PE); with ethanolamine as starting material, the corresponding intermediate is a dihydropyridinium bisretinoid (dihydropyridinium-A2E).

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1. A mixture of all-trans-retinal (300 μg, 2 equivalents) and dipalmitoylphosphatidylethanolamine (364 μg, 2 equivalents) or ethanolamine (32 μg, 1 equivalent) in ethanol (3 ml) is stirred in the presence of acetic acid (0.3 μl) at room temperature in a capped vial in the dark for 3–7 days. To slow the rate of synthesis, the concentrations of starting materials can be decreased (e.g., 1/10). 2. Separate reaction mixtures should be incubated in parallel if independent analysis at various time points (e.g., 1, 4, and 8 h and 1, 3, and 7 days) is desired. 3. After incubating, 10 μl of the reaction mixture is injected into the HPLC using reverse-phase C18 column and elution with the following gradients of acetonitrile in water (containing 0.1% TFA): for A2PE-H2 detection, 75–90% (0–30 min; flow); 90–100% (30–40 min); 100% (40–80 min) with a flow rate of 0.5 ml/min; for dihydropyridinum-A2E detection, 85–100% (15 min); 100% (15–20 min) with a flow rate of 0.8 ml/min. Monitor by photodiode array at 440 and 490 nm. 4. The hydrophobic long alkyl chain in the stationary phase of the C18 column is designed to capture small molecules; thus with the C18 column, good chromatographic separation is obtained for A2E, isoA2E, all-trans-retinal dimer, alltrans-retinal dimer-PE, and all-trans-retinal dimer-E. A2PE is more strongly retained on the C18 column but is effectively eluted using a C4 column. The order of compound elution is based on hydrophobicity. All of these compounds are detected by photodiode array monitoring at 430 nm; however, 500 nm detection should be used for quantitation of all-trans-retinal dimer-PE and all-trans-retinal dimer-E (see Note 1). 5. Based on retention times, co-elution with authentic standards and UV–visible absorbance spectra, the following compounds can be identified: (i) the starting material, all-trans-retinal; (ii) compounds formed transitionally, N-retinylidene-PE, dihydropyridinium-A2PE (the corresponding intermediate when starting materials are all-transretinal and ethanolamine is dihydropyridinium-A2E); (iii) A2-dihydropyridine-phosphatidylethanolamine (A2-DHPPE) (3); (iv) A2PE; (v) A2E, the product released by PLD-mediated phosphate hydrolysis of A2PE (see Note 2) (Fig. 18.3). Compounds of the all-trans-retinal dimer series also form in reaction mixtures of all-trans-retinal and phosphatidylethanolamine. 6. Software designed to integrate chromatographic peaks is used to determine peak area. From a standard curve con-

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structed using known concentrations of A2E, picomoles of A2E can also be computed (see Note 3). 7. For experiments comparing synthesis under normal air and deoxygenated conditions, the starting materials are combined and the vessel is purged with argon before capping. In an environment of reduced oxygen, the yield of dihydropyridinium-A2PE is augmented and oxidative aromatization of dihydropyridinium-A2PE to generate A2PE is delayed (see Note 4) (23). 8. To generate A2E from A2PE, the hydrophobic fraction containing A2PE is reconstituted in dimethyl sulfoxide (DMSO), and 15 μl is added to 285 μl of a mixture (3:7) of 40 mM MOPS buffer (pH 6.5) containing 300 units/ml PLD. The final concentration of A2PE is 100 μM. The mixture is incubated for 3 h at 37◦ C, extracted with chloroform/methanol (2:1, v/v) and 0.1% TFA, dried under argon, and re-dissolved in 300 μl of 50% methanolic chloroform. The injection volume is 10 μl. A2PE and A2E are detected by HPLC using a reverse-phase C4 column, monitoring at 430 nm, and elution with the following gradients of acetonitrile in water (containing 0.1% TFA): 75% (5 min; flow rate, 1 ml/min), 75–100% (5 min; flow rate, 1.5 ml/min), and 100% (10 min; flow rate, 1.5 ml/min). 9. A2E is purified by silica gel column chromatography with elution by methanol:chloroform (5:95) and further elution with methanol:chloroform:trifluoroacetic acid (8:92:0.001). Pure samples are obtained by HPLC purification using a C18 column with a gradient of water and methanol (85–96%) and 0.1% trifluoroacetic acid (see Notes 5 and 6). A2E should be stored at −80◦ C in an amber vial wrapped in foil. 3.2. Constructing a Cell Culture Model

1. ARPE-19 cells are plated and grown to confluence in DMEM with 10% fetal bovine serum, 2 mM glutamine, 0.1 mM MEM nonessential amino acid, and 10 μg/ml gentamicin sulfate. After reaching confluence, serum is reduced to 5%. 2. The cells are incubated with 10 μM A2E delivered in culture medium. Control cultures are not treated or are incubated with an equivalent concentration of DMSO only (see Note 8). 3. The accumulation of A2E can be monitored as the acquisition of yellow granules under bright field microscopy or by visualizing a golden yellow autofluorescence which is detectable by epifluorescence microscopy using a bandpass excitation filter centered on 440 nm and a 475 nm long pass

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filter. For HPLC measurements of A2E levels in the cultures, cells can be harvested by scraping and aspiration; amounts are then expressed as nanograms/105 cells (see Note 9). 4. When this approach is followed, A2E accumulates in the lysosomal compartment of the cells (26). This can be demonstrated using LysoTracker dyes such as LysoTracker Red DND-99. Cells are incubated with the latter dye in culture media at a concentration of 50 nM for 2 h. The cultures are washed, fixed with 2% paraformaldehyde in PBS, and examined using a confocal laser scanning system. Exclusion from mitochondria can be demonstrated in parallel assays using a fluorescent mitochondrial marker (MitoTracker Red CM-H2Xros, 500 nM). 5. Companion cultures should be examined for the presence of nuclei-stained membrane-compromised cells (see Note 8) using fluorescent membrane-impermeant dyes such as Dead Red. Cultures are incubated with Dead Red (1/500 dilution in HEPES-buffered Hanks’ balanced salt solution) for 15 min, after which the cells are washed and fixed in 4% glutaraldehyde for 1 h. Cultures are examined under a fluorescence microscope with excitation at λ 545 ± 15 nm and visualization at 535 ± 25 nm (26).

4. Notes 1. It should be noted that A2PE, A2-DHP-PE, and dihydropyridinium-A2PE are actually compound mixtures with phosphatidic acid moieties that vary in fatty acid composition. Thus by HPLC, these pigments are detected as broad peaks. By mass spectrometry these chromophores present with multiple mass to charge (m/z) ratios. For instance, we have shown that the A2PE region of the FABMS profile consists of a complex cluster of peaks (including m/z 1,280.9, 1,294.8, 1,310.9, 1,322.8, 1,338.8), the particularly intense signals at m/z 1,294.8 and 1,322.8 being attributable to PE with fatty acids docosahexaenoic acid (22:6)/palmitic acid (16:0) and docosahexaenoic acid/stearic acid (18:0), the fatty acids that are especially common in outer segments (15). 2. The molecular weights of the compounds are A2E, 592; isoA2E, 592; dihydropyridinium-A2PE, 1226; A2PE, 1224; all-trans-retinal dimer, 550; all-trans-retinal dimer-E, 594; all-trans-retinal dimer-PE, 1225; and A2-dihydropyridinephosphatidylethanolamine (A2-DHP-PE), 1225. Note that the molecular weights given for all of the phosphatidyl-

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bisretinoid compounds (A2PE, dihydropyridinium-A2PE, A2-DHP-PE, and all-trans-retinal dimer-PE) are based on synthesis with dipalmitoyl-PE. 3. The extinction coefficients of A2E at the following wavelengths are 439 nm, εM 36,900; 336 nm, ε M 25,600 and of isoA2E are 426 nm, ε M 31,000; 335 nm, εM 27,000. 4. UV–visible absorbance maxima of dihydropyridiniumA2PE/dihydropyridinium-A2E (494, 344 nm) were calculated by time-dependent density functional theory (23). The calculated absorbance maxima agreed well with the absorbance maxima obtained experimentally under HPLC conditions. 5. The elemental composition of A2E is C42 H58 NO, molecular weight 592. Although A2E is often described as N-retinyl-N-retinylidene-ethanolamine, this structural assignment is incorrect as it does not connote the pyridinium head group, an important structural feature of A2E. The quaternary amine nitrogen of A2E carries a permanent positive charge. The counterion of this pyridinium salt is likely chloride in vivo and trifluoroacetate in the case of synthetic samples. 6. A2E and isoA2E are in photoequilibrium at a ratio of approximately 4:1. Thus samples of A2E contain approximately 15% isoA2E. 7. ARPE-19 cells are employed because they are devoid of endogenous lipofuscin (26). Human fetal RPE also does not contain endogenous lipofuscin compounds and thus we also use these cells in our assays (6). Human fetal eyes can be obtained from Advanced Bioscience Resources (Alameda, CA) with Institutional Review Board approval. 8. Higher concentrations of A2E in culture media cause membrane damage (26, 27). Besides being a photoreactive compound, A2E is an amphiphilic compound exhibiting detergent-like activity. Cultures of RPE that have accumulated A2E should be left quiescent for a minimum of 7 days before use; this period allows for the attrition of any membrane-damaged cells. Detergent-like effects of A2E could contribute to several of the effects of A2E that have been reported in the literature (28–30), particularly when cells are incubated at high concentrations (e.g., 50 and 100 μM) even for short periods of time (e.g., 6 h). 9. The mechanism by which A2E accumulates in cultured RPE cells has not been demonstrated. However, A2E is not a lysosomotropic compound as is the case with alkyl amines that cross the cell membrane in a deprotonated state and become trapped in the acidic environment of lysosomes after

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protonation. Rather, as noted above, the quaternary amine nitrogen of A2E carries a permanent positive charge and unlike lysosomotropic compounds does not deprotonate and reprotonate. As such A2E and isoA2E neither would deprotonate/protonate in response to a change in pH nor would A2E/isoA2E generate a change in pH by deprotonation/protonation.

Acknowledgments This work was supported by National Institutes of Health Grant EY12951, the Kaplen Foundation, a gift from Dr. Gertrude Neumark Rothschild, and a grant from Research to Prevent Blindness to the Department of Ophthalmology. JRS is the recipient of a Research to Prevent Blindness Senior Investigator Award. References 1. Sparrow, J.R., Boulton, M. (2005) RPE lipofuscin and its role in retinal photobiology. Exp. Eye Res. 80, 595–606. 2. Sparrow, J.R. (2007) RPE lipofuscin: Formation, properties and relevance to retinal degeneration. In: Tombran-Tink, J., Barnstable, C.J., (eds.), Retinal Degenerations: Biology, Diagnostics and Therapeutics, Humana Press, Totowa, NJ. 3. Wu, Y., Fishkin, N.E., Pande, A., Pande, J., Sparrow, J.R. (2009) Novel lipofuscin bisretinoids prominent in human retina and in a model of recessive Stargardt disease. J. Biol. Chem. 284, 20155–20166. 4. Fishkin, N., Sparrow, J.R., Allikmets, R., Nakanishi, K. (2005) Isolation and characterization of a retinal pigment epithelial cell fluorophore: An all-trans-retinal dimer conjugate. Proc. Natl. Acad. Sci. USA 102, 7091–7096. 5. Kim, S.R., Jang, Y.P., Jockusch, S., Fishkin, N.E., Turro, N.J., Sparrow, J.R. (2007) The all-trans-retinal dimer series of lipofuscin pigments in retinal pigment epithelial cells in a recessive Stargardt disease model. Proc. Natl. Acad. Sci. USA 104, 19273–19278. 6. Parish, C.A., Hashimoto, M., Nakanishi, K., Dillon, J., Sparrow, J.R. (1998) Isolation and one-step preparation of A2E and iso-A2E, fluorophores from human retinal pigment epithelium. Proc. Natl. Acad. Sci. USA 95, 14609–14613.

7. Sparrow, J.. (2007) Lipofuscin of the retinal pigment epithelium. In: Holz, F.G., Schmitz-Valckenberg, S., Spaide, R.F., Bird, A.C. (eds.), Atlas of Autofluorescence Imaging, Springer, Heidelberg. 8. Sparrow, J.R., Kim, S.R., Jang, Y.P., Zhou, J. (2008) The lipofuscin of retinal pigment epithelial cells: Learning from mouse models of retinal disease. In: Chalupa, L.M. (ed.), Eye, Retina, and Visual System of the Mouse, MIT Press, Cambridge, MA, pp. 539–546. 9. Sparrow, J.R., Zhou, J., Ben-Shabat, S., Vollmer, H., Itagaki, Y., Nakanishi, K. (2002) Involvement of oxidative mechanisms in blue light induced damage to A2Eladen RPE. Invest. Ophthalmol. Vis. Sci. 43, 1222–1227. 10. Sparrow, J.R., Nakanishi, K., Parish, C.A. (2000) The lipofuscin fluorophore A2E mediates blue light-induced damage to retinal pigmented epithelial cells. Invest. Ophthalmol. Vis. Sci. 41, 1981–1989. 11. Jang, Y.P., Matsuda, H., Itagaki, Y., Nakanishi, K., Sparrow, J.R. (2005) Characterization of peroxy-A2E and furan-A2E photooxidation products and detection in human and mouse retinal pigment epithelial cells lipofuscin. J. Biol. Chem. 280, 39732–39739. 12. Ben-Shabat, S., Itagaki, Y., Jockusch, S., Sparrow, J.R., Turro, N.J., Nakanishi, K. (2002) Formation of a nona-oxirane from A2E, a lipofuscin fluorophore related to mac-

Experimental Approaches to the Study of A2E

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

ular degeneration, and evidence of singlet oxygen involvement. Angew. Chem. Int. Ed. 41, 814–817. Kim, S.R., Jockusch, S., Itagaki, Y., Turro, N.J., Sparrow, J.R. (2008) Mechanisms involved in A2E oxidation. Exp. Eye Res. 86, 975–982. Ben-Shabat, S., Parish, C.A., Vollmer, H.R., Itagaki, Y., Fishkin, N., Nakanishi, K., Sparrow, J.R. (2002) Biosynthetic studies of A2E, a major fluorophore of RPE lipofuscin. J. Biol. Chem. 277, 7183–7190. Liu, J., Itagaki, Y., Ben-Shabat, S., Nakanishi, K., Sparrow, J.R. (2000) The biosynthesis of A2E, a fluorophore of aging retina, involves the formation of the precursor, A2-PE, in the photoreceptor outer segment membrane. J. Biol. Chem. 275, 29354–29360. Illing, M., Molday, L.L., Molday, R.S. (1997) The 220-kDa rim protein of retinal rod outer segments is a member of the ABC transporter superfamily. J. Biol. Chem. 272, 10303–10310. Sun, H., Molday, R.S., Nathans, J. (1999) Retinal stimulates ATP hydrolysis by purified and reconstituted ABCR, the photoreceptorspecific ATP-binding cassette transporter responsible for Stargardt disease. J. Biol. Chem. 274, 8269–8281. Molday, L.L., Rabin, A.R., Molday, R.S. (2000) ABCR expression in foveal cone photoreceptors and its role in Stargardt macular dystrophy. Nat. Genet. 25, 257–258. Ahn, J., Wong, J.T., Molday, R.S. (2000) The effect of lipid environment and retinoids on the ATPase activity of ABCR, the photoreceptor ABC transporter responsible for Stargardt macular dystrophy. J. Biol. Chem. 275, 20399–20405. Weng, J., Mata, N.L., Azarian, S.M., Tzekov, R.T., Birch, D.G., Travis, G.H. (1999) Insights into the function of Rim protein in photoreceptors and etiology of Stargardt s disease from the phenotype in abcr knockout mice. Cell 98, 13–23. Sun, H., Nathans, J. (2001) ABCR, the ATP-binding cassette transporter responsible for Stargardt macular dystrophy, is an efficient target of all-trans retinal-mediated photo-oxidative damage in vitro: Implications for retinal disease. J. Biol. Chem. 276, 11766–11774. Allikmets, R., Singh, N., Sun, H., Shroyer, N.F., Hutchinson, A., Chidambaram, A.,

23.

24.

25.

26.

27.

28.

29.

30.

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Gerrard, B., Baird, L., Stauffer, D., Peiffer, A., Rattner, A., Smallwood, P., Li, Y., Anderson, K.L., Lewis, R.A., Nathans, J., Leppert, M., Dean, M., Lupski, J.R. (1997) A photoreceptor cell-specific ATP-binding transporter gene (ABCR) is mutated in recessive Stargardt macular dystrophy. Nat. Genet. 15, 236–246. Kim, S.R., He, J., Yanase, E., Jang, Y.P., Berova, N., Sparrow, J.R., Nakanishi, K. (2007) Characterization of dihydro-A2PE: An intermediate in the A2E biosynthetic pathway. Biochemistry 46, 10122–10129. Sparrow, J.R., Kim, S.R., Cuervo, A.M., Bandhyopadhyayand, U. (2008) A2E, a pigment of RPE lipofuscin is generated from the precursor A2PE by a lysosomal enzyme activity. Adv. Exp. Med. Biol. 613, 393–398. Ng, K.P., Gugiu, B.G., Renganathan, K., Davies, M.W., Gu, X., Crabb, J.S., Kim, S.R., Rozanowska, M.B., Bonilha, V.L., Rayborn, M.E., Salomon, R.G., Sparrow, J.R., Boulton, M.E., Hollyfield, J.G., Crabb, J.W. (2008) Retinal pigment epithelium lipofuscin proteomics. Mol. Cell Proteomics 7, 1397–1405. Sparrow, J.R., Parish, C.A., Hashimoto, M., Nakanishi, K. (1999) A2E, a lipofuscin fluorophore, in human retinal pigmented epithelial cells in culture. Invest. Ophthalmol. Vis. Sci. 40, 2988–2995. Sparrow, J.R., Fishkin, N., Zhou, J., Cai, B., Jang, Y.P., Krane, S., Itagaki, Y., Nakanishi, K. (2003) A2E, a byproduct of the visual cycle. Vision Res. 43, 2983–2990. De, S., Sakmar, T.P. (2002) Interaction of A2E with model membranes. Implications to the pathogenesis of age-related macular degeneration. J. Gen. Physiol. 120, 147–157. Vives-Bauza, C., Anand, M., Shirazi, A.K., Magrane, J., Gao, J., Vollmer-Snarr, H.R., Manfredi, G., Finnemann, S.C. (2008) The age lipid A2E and mitochondrial dysfunction synergistically impair phagocytosis by retinal pigment epithelial cells. J. Biochem. Chem. 283, 24770–24780. Holz, F.G., Schutt, F., Kopitz, J., Eldred, G.E., Kruse, F.E., Volcker, H.E., Cantz, M. (1999) Inhibition of lysosomal degradative functions in RPE cells by a retinoid component of lipofuscin. Invest. Ophthalmol. Vis. Sci. 40, 737–743.

Chapter 19 Analysis of the Retinoid Isomerase Activities in the Retinal Pigment Epithelium and Retina Gabriel H. Travis, Joanna Kaylor, and Quan Yuan Abstract Light sensitivity in the vertebrate retina is mediated by the opsin visual pigments inside rod and cone photoreceptor cells. These pigments consist of a G protein-coupled receptor and the photo-sensitive ligand, 11-cis-retinaldehyde (11-cis-RAL). Absorption of a photon by an opsin pigment induces isomerization of the 11-cis-RAL chromophore to all-trans-retinaldehyde (all-trans-RAL), rendering the pigment insensitive to light. The bleached opsin regains light sensitivity by recombining with another 11-cis-RAL. The vertebrate eye contains a biochemical mechanism for regenerating 11-cis-RAL chromophore from all-trans-RAL, called the visual cycle. The visual cycle takes place within cells of the retinal pigment epithelium (RPE). A second visual cycle also appears to be present in Müller glial cells of the retina. A critical step in the regeneration of 11-cis-RAL chromophore is thermal re-isomerization to the 11-cis configuration of an all-trans-retinyl ester (all-trans-RE) or an all-trans-retinol (all-trans-ROL). In RPE cells, this step is carried out by an enzyme called Rpe65 isomerase. This chapter provides methods for assaying Rpe65 isomerase. Although Rpe65 utilizes an all-trans-RE such as all-trans-retinyl palmitate (alltrans-RP) as substrate, it can be assayed in RPE homogenates by providing all-trans-ROL substrate and allowing the endogenous lecithin:retinol acyl transferase (LRAT) to synthesize all-trans-REs using fatty acids from phosphatidylcholine in the membranes. Alternatively, all-trans-RP can be provided directly as substrate, although this requires the isomerase reaction to be carried out in the presence of detergent, since fatty-acyl esters of all-trans-ROL are insoluble. Methods are provided in this chapter for assaying Rpe65 in RPE homogenates with both all-trans-ROL and all-trans-RP substrates. A second visual cycle appears to be present in the retinas of cone-dominant species such as chicken. This retinal pathway may augment the RPE to provide 11-cis-RAL to cone photoreceptors under conditions of bright light where the rate of opsin photoisomerization is high. The isomerase in this pathway (isomerase-2) utilizes alltrans-ROL and palmitoyl coenzyme A (palm CoA) as substrates to synthesize 11-cis-retinyl palmitate (11-cis-RP). Isomerase-2 appears to be present in Müller cells but has not yet been identified. Methods are provided in this chapter for assaying isomerase-2 in chicken retina homogenates. Key words: Cellular retinaldehyde-binding protein, chicken, chromophore, cone, enzyme assay, isomerase, lecithin retinol acyl transferase, Müller cell, photoreceptor, retina, retinal pigment epithelium, retinol, retinyl ester, rhodopsin, rod, Rpe65, visual cycle.

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1. Introduction Rods and cones are two types of photoreceptor cells present in the vertebrate retina. These cells contain a light-sensitive membranous process called the outer segment (OS). OS membranes are packed with rhodopsin or cone-opsin visual pigments. Opsins are members of the G protein-coupled receptor superfamily. The light-absorbing ligand or chromophore in most vertebrate opsins is 11-cis-retinaldehyde (11-cis-RAL), which is covalently coupled to a Lys residue in the protein. Absorption of a photon by an opsin pigment induces isomerization of the 11-cis-RAL to all-trans-retinaldehyde (all-trans-RAL). After a brief period of activation, the pigment decays to yield free all-trans-RAL and apo-opsin. Apo-opsin regains light sensitivity by recombining with a new molecule of 11-cis-RAL to form another rhodopsin or cone-opsin pigment. The released all-trans-RAL is converted back to 11-cis-RAL by a multi-step enzyme pathway called the visual cycle (Fig. 19.1). The first catalytic step of the visual cycle, reduction of all-trans-RAL to all-trans-retinol (all-trans-ROL), takes place within the photoreceptor OS. The

Fig. 19.1. Visual cycle in RPE cells. 11-cis-RAL in rhodopsin is isomerized by absorption of a photon (hv). The resulting all-trans-RAL is reduced to all-trans-ROL by all-transRDH in the rod outer segment. The all-trans-ROL is taken up by an RPE cell where it is esterified to a fatty acid by LRAT. The resulting all-trans-RE is isomerized and hydrolyzed by Rpe65 to 11-cis-ROL. The 11-cis-ROL is oxidized to 11-cis-RAL by 11-cis-RDH. Both 11-cis-ROL and 11-cis-RAL are bound to CRALBP within the RPE cell. During transit through the extracellular space, all-trans-ROL and 11-cis-RAL are bound to IRBP (not shown).

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remaining steps take place within cells of the retinal pigment epithelium (RPE). The all-trans-ROL is released by photoreceptors into the extracellular space where it binds to interphotoreceptor retinoid-binding protein (IRBP). RPE cells take up all-trans-ROL where it is esterified to a fatty acid derived from phosphatidylcholine through the action of lecithin:retinol acyltransferase (LRAT) to yield an all-trans-retinyl ester (all-transRE) such as all-trans-retinyl palmitate (all-trans-RP). These alltrans-REs are substrates for Rpe65, which catalyzes the critical all-trans to 11-cis isomerization step. The resulting 11-cis-retinol (11-cis-ROL) product of Rpe65 binds to cellular retinaldehydebinding protein (CRALBP) before being oxidized by one of several 11-cis-retinol dehydrogenases (11-cis-RDHs) in RPE cells to yield 11-cis-RAL chromophore. 11-cis-RAL also binds to CRALBP. Finally, the 11-cis-RAL is released by RPE cells into the extracellular space, where it too binds IRBP, is taken up by the photoreceptors, and re-combines with apo-opsin to form a new light-sensitive pigment. Rpe65 is strongly associated with internal membranes although it contains no membrane spanning segments. Similarly, the substrates for Rpe65, all-trans-REs, are insoluble in water and present within the lipid bilayer. The remaining activities of the visual cycle in RPE cells, LRAT, and 11-cis-RDHs are integral membrane proteins of the endoplasmic reticulum (ER). Rpe65 activity can be assayed in total homogenates or microsomal fractions of RPE cells. Microsomal fractions contain higher isomerase-specific activity but lower total activity. The microsomal fraction also contains significantly lower endogenous retinoids. With mice it is convenient to assay isomerase activity in homogenates of eyecups (the posterior half of the eyeball containing RPE, choroid, and sclera after removing the cornea, lens, vitreous, and retina). The isomerase activity of Rpe65 in RPE homogenates can be assayed in vitro by two approaches. The first involves the use of all-trans-ROL substrate, which is much more water-soluble than all-trans-RP. This approach depends on LRAT to synthesize all-trans-REs and insert them into Rpe65containing membranes. Rpe65 utilizes these all-trans-REs as substrate to synthesize 11-cis-ROL. The second approach is to provide directly all-trans-RP as substrate. Here, the reaction must be carried out in the presence of detergent to solubilize the substrate. Müller glial cells of the retina express several proteins involved in the processing of visual retinoids. These include (i) CRALBP (1, 2), (ii) RPE-retinal G protein-coupled receptor (RGR-opsin), a non-photoreceptor opsin that effects light-dependent regulation of the visual cycle in RPE cells (3), (iii) CRBP1 (4), and (iv) retinol dehydrogenases types 10 and 11. However, Müller cells do not express Rpe65 (5) nor LRAT (6), which are both present in RPE cells. Therefore, Müller cells do not simply duplicate the

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function of RPE cells in the regeneration of visual chromophore. Several lines of evidence suggest that an alternate source of chromophore precursor is available to cones but not to rods. When frog retinas were separated from the RPE, cone opsins, but not rhodopsin, regenerated spontaneously after a photobleach (7–10). After photobleaching, isolated salamander cones recovered sensitivity with addition of either 11-cis-ROL or 11-cis-RAL, while isolated rods only recovered sensitivity with addition of 11cis-RAL (11). Müller cells in primary culture were shown to take up all-trans-ROL and synthesize 11-cis-ROL, which they secreted into the medium (12). Salamander cones were shown to dark adapt and regenerate visual chromophore in isolated retinas separate from the RPE (13). The intrinsic capacity of cones to recover sensitivity and regenerate visual chromophore in salamander retinas was lost after exposure to a selective Müller cell toxin (αaminoadipic acid), suggesting that Müller cells play a role in these processes (13). A retinoid isomerase activity, distinct from the activity catalyzed by Rpe65, is detectable in homogenates of cone-dominant ground squirrel and chicken retinas separated from the RPE. This isomerase-2 activity catalyzes the conversion of all-trans-ROL into 11-cis-ROL and 11-cis-retinyl esters (11-cis-REs) such as 11-cis-retinyl palmitate (11-cis-RP) (6, 14). The protein responsible for isomerase-2 activity has not yet been identified. Formation of 11-cis-ROL is favored when the reaction is carried out in the presence of CRALBP. Addition of palmitoyl coenzyme A (palm CoA) to the reaction mixture favors formation of 11-cis-RP. These observations suggest that isomerase-2 catalyzes the direct interconversion of all-trans-ROL and 11-cisROL. At equilibrium, all-trans-ROL is favored over 11-cis-ROL by ∼1000:1 (G  + 4.1 kcal/mol). Thus, removal of 11-cis-ROL from the equilibrium mixture is required to drive all-trans to 11cis isomerization of the retinoid. This appears to be accomplished by subsequent esterification of 11-cis-ROL by a palm CoAdependent retinyl ester synthase and acyl-CoA:retinol acyltransferase (ARAT). High-affinity binding of 11-cis-ROL by CRALBP also drives all-trans to 11-cis isomerization. The proteins responsible for the isomerase-2 and ARAT activities have not yet been identified.

2. Materials 2.1. Buffers and Solutions for Rpe65-Isomerase Assays

1. HEPES buffer: 20 mM HEPES (pH 7.4), 150 mM NaCl. One tablet of Roche Complete Protease Inhibitor (EDTA free) is dissolved in 50 ml of HEPES buffer. 2. 15% (w/w) bovine serum albumin (BSA) in HEPES buffer: 1.5 g of BSA dissolved in 10 ml of HEPES buffer.

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3. 0.5 M sodium cholate in HEPES buffer: 215 mg of sodium cholate is dissolved in 1.0 ml of HEPES buffer. 4. 400 μM all-trans-ROL in ethanol. 5. 400 μM all-trans-RP in hexane (HPLC grade). 6. 1.0% SDS solution: 1.0 g of sodium dodecyl sulfate is dissolved in 100 ml of double distilled water. 2.2. Buffers and Materials for Isomerase-2 Assay

1. 20% BSA. 2. 0.1 M NaOAc, pH 6.0. 3. 5.0 mM palm CoA in 0.1 M NaOAc, pH 6.0. 4. 1.0 mM all-trans-ROL in dimethylsulfoxide (DMSO). 5. Homogenization buffer: 10 mM HEPES (pH 7.5), 100 mM NaCl, 1.0 mM MgCl2 , 1.0 mM CaCl2 , protease inhibitor cocktail. 6. Assay buffer: 10 mM Tris-HCl (pH 8.0), 2.0 mM MgCl2 , 2.0 mM CaCl2 , 1.0 mM dithiothreitol (DTT). 7. Glass-to-glass homogenizer. 8. Pierce Micro BCA Protein Concentration Kit.

2.3. HPLC Analysis of Retinoids

1. Liquid chromatogram equipped with photodiode array detector, such as Agilent 1100. 2. Silica column for normal-phase HPLC such as Supelcosil LSSI 5 μm, 4.6 mm × 250 mm ID.

3. Methods 3.1. Assays for Rpe65-Isomerase in RPE 3.1.1. Preparation of Bovine RPE Homogenate

1. Fifty freshly slaughtered bovine eyeballs are dark adapted on ice for 1 h. 2. On ice, section the anterior one-third of the eyeball and discard the lens, vitreous, and retina. 3. Invert the eyecup and gently brush the RPE into HEPES buffer at 4◦ C. 4. Collect the pooled RPE cells in HEPES buffer by centrifugation at 500×g for 10 min. 5. Discard the supernatant and resuspend the pellet in 5.0 ml of HEPES buffer. 6. Disrupt the cells in a nitrogen cavitation bomb at 500 psi. 7. Remove cell nuclei and intact cells by centrifugation at 1,000×g for 10 min.

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8. Flash-freeze 50 μl aliquots of the supernatant (homogenate) in liquid nitrogen and store at −80◦ C for use in isomerase assays. 9. At time of use, determine the total protein concentration with a Micro BCA Assay Kit. 3.1.2. Rpe65-Isomerase Assay Using All-Trans-ROL as Substrate

1. Expose bovine RPE homogenate to handheld 365-nm UV light source for 5 min on ice to bleach endogenous retinoids (see Note 2). 2. Add 160 μl of 15% BSA in HEPES buffer to 50 μl of RPE homogenate (50–100 μg total protein) on ice. 3. Add 190 μl of HEPES buffer. Transfer samples to darkness (see Note 1). 4. Add 10 μl of 400 μM all-trans-ROL in ethanol and mix. 5. Incubate samples at 37◦ C in darkness for 1 h.

3.1.3. Extraction and Analysis of Retinoids by Liquid Chromatography

1. Add 100 μl 1% SDS, vortex briefly, incubate at room temperature 10 min to denature proteins. 2. Add 1 ml ice-cold methanol, vortex briefly. 3. To each sample add 2 ml hexane and vortex. Centrifuge at 3000×g for 5 min to separate phases. Transfer the hexane layer (top) to a clean 12-mm×75-mm test tube. Repeat extraction of aqueous phase and combine hexane extracts (4 ml total). These extracts may be stored at −20◦ C in the dark or analyzed immediately. 4. Filter extracts through a polypropylene 200-μm disposable filter column (e.g., Fisher Scientific #11-387-50). 5. Dry the extract in the dark under a stream of filtered nitrogen or argon gas. Dissolve the dried retinoids in 100 μl of hexane. 6. Separate retinoids by chromatography on a silica column using gradient elution (0.2–10% dioxane in hexane) at 2.0 ml/min in an Agilent 1100 liquid chromatograph equipped with a UV photodiode array detector. The wavelength of maximal absorbance (λmax ) for 11-cis-ROL is 318 nm; therefore this wavelength should be monitored during chromatography. The λmax for all-trans-ROL and alltrans-RP is 325 nm. Confirm identification of UV-absorbing peaks by acquisition of UV spectra and co-elution with authentic retinoid standards. 7. Quantitate each retinoid by comparing the sample peak area to a calibration curve established with an authentic retinoid standard. Figure 19.2 shows a representative chromatogram of retinoids following an assay for isomerase activity in RPE.

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Fig. 19.2. Representative retinoid chromatograms. (a) Chromatogram of retinoid standards showing UV absorption at 318 nm. Identified 11-cis-RP, all-trans-RP, 11-cis-RAL, all-trans-RAL, 11-cis-ROL, and all-trans-ROL peaks are labeled. (b) Chromatogram of chicken retina homogenate in the isomerase-2 assay mixture before incubation (t 0 ) showing 20–25 min elution times. This chromatogram shows endogenous 11-cis-ROL and 13-cis-ROL, plus all-trans-ROL substrate (off scale) added to the assay mixture. (c) Chromatogram of chicken retina homogenate in isomerase-2 assay mixture following 15 min incubation at 37◦ C (t15–min ). Note the higher 11-cis-ROL peak. The difference in 11-cis-ROL before and after incubation reflects the isomerase-2 activity. 13-cis-ROL increases during the incubation due to thermal isomerization of 11-cis-ROL product and all-trans-ROL substrate. (d) UV spectrum of 11-cis-ROL. The wavelength of maximum absorption (λmax ) for 11-cis-ROL is 318 nm. (e) UV spectrum of 13-cis-ROL (λmax , 328 nm). (f) UV spectrum of all-trans-ROL (λmax, 325 nm).

8. Determine the isomerase-specific activity by subtracting endogenous 11-cis-ROL (from the assay mixture at t0 ) from 11-cis-ROL in the post-incubation assay mixture (t15–min ) and dividing by the incubation time and total protein in the assay mixture. 3.1.4. Rpe65-Isomerase Assay Using all-trans-RP as Substrate

1. Expose RPE homogenate to UV light (see Note 2). 2. Add 160 μl 15% BSA in HEPES buffer to 50 μl RPE homogenate (50–100 μg total protein) on ice. 3. In the dark, dry 10 μl of 400 μM all-trans-RP in hexane under a stream of filtered nitrogen gas in a separate tube (see Note 1). 4. Add 5 μl of 500 mM sodium cholate solution plus 185 μl of HEPES buffer to the dried all-trans-RP substrate. Vortex for 2 min. 5. Add substrate solution to RPE homogenate (final assay volume 400 μl). Incubate in the dark for 1 h at 37◦ C. 6. Add 100 μl of 1.0% SDS to each sample to quench the reaction. Incubate at 25◦ C for 10 min. 7. Add 1.0 ml methanol to precipitate proteins.

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8. Pellet the precipitated proteins by centrifugation at 1000×g in a benchtop centrifuge. 9. Extract the supernatant twice with 2.0 ml hexane. Pool extracts and filter through a polypropylene 200-μm disposable filter column. 10. Dry the extract in the dark under a stream of filtered nitrogen gas. 11. Dissolve the dried retinoids in 100 μl of hexane. Analyze by HPLC as described in Section 2.4.6. 3.2. Assay for Retinol Isomerase-2 in Chicken Retinas 3.2.1. Preparation of Chicken Retina Homogenates

1. Obtain heads from freshly slaughtered chickens (see Notes 3–5). 2. Remove the eyes from the chicken head using a scalpel, surgical scissors, large scissors, and two forceps, one with and without grasping points on the end. 3. On ice, remove the anterior ∼1/3 of the eye with a scalpel. Place the opened eyeballs into small Petri dishes containing phosphate-buffered saline (PBS) on ice and place in the dark for 1 h to dark adapt the retina. This facilitates removal of the retina from the RPE. 4. Transfer to a clean Petri filled with fresh PBS in light. Using a Pasteur pipet, gently squirt PBS underneath the yellowish retina to separate it from the black RPE and choroid. After separating, free the retina from the optic nerve head by snipping with microsurgery scissors. Remove any adhering black RPE from the retina with forceps. 5. Transfer the retina to a fresh, tared 1.5 ml tube, determine the tissue wet weight, store on ice for immediate use. 6. Homogenize approximately two retinas (∼25 mg) in 1 ml of ice-cold homogenization buffer using the glass-to-glass homogenizer. 7. Assay the total protein content with the Micro BCA kit. Dilute the concentration to 2 mg/ml using Assay buffer. Store on ice.

3.2.2. Isomerase-2 Assay

1. Set up a labeled 12-mm × 75-mm glass test tube for each assay which will have a total volume of 500 μl. 2. Combine 250 μl of protein homogenate or microsomes, 200 μl of assay buffer, and 25 μl of 20% BSA to each tube, mix, and preincubate at 37◦ C for 5 min. 3. Add 15 μl of palm CoA, mix, and preincubate for 2 min.

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4. Add 10 μl of all-trans-ROL substrate to begin the reaction, mix, and incubate at 37◦ C for 10–20 min (see Note 1). 5. After incubating, transfer the assay mixtures to 12-mm× 75-mm extraction tubes. Follow the retinoid extraction protocol described in Section 2.4. 6. Analyze retinoids by HPLC as described in Section 2.4. 3.2.3. Saponification

1. After drying under a stream of nitrogen gas, dissolve the samples in 980 μl of ethanol (see Note 6). 2. Add 20 μl of 6 M KOH and vortex briefly (120 mM ethanolic KOH final). 3. Incubate at 55◦ C for 10 min. 4. Place the tubes on ice and add 1 ml of cold dH2 O. 5. Extract retinoids into hexane as described in Section 2.4. 6. Analyze samples by normal-phase HPLC as described in Section 2.4.

4. Notes 1. All manipulations of retinoids should be done in the darkroom under dim red light. 2. Bleaching of endogenous retinoids in the homogenate with UV light can be eliminated to yield slightly higher isomerasespecific activity. Eliminating this step will result in a higher 11-cis-ROL background. 3. Commercially slaughtered chickens are commonly scalded after killing to facilitate removal of feathers. This treatment will inactivate enzymes in the eye. Ensure that the chicken heads have not been exposed to hot water or steam. 4. Isomerase-2 assay conditions also detect Rpe65 activity, while the Rpe65 assay conditions with all-trans-RP substrate do not detect isomerase-2 activity. 5. Rpe65 and isomerase-2 activities can be distinguished by assaying homogenates of chicken retina and RPE under conditions described in Sections 2.5 and 3.3. Chicken retina homogenates contain virtually no all-trans-RP-dependent isomerase activity, while this activity is abundantly present in RPE homogenates (Fig. 19.3a). In contrast, chicken retina homogenates show much greater synthesis of 11-cis-ROL than RPE during incubation with all-trans-ROL and palm CoA (Fig. 19.3b). 6. The products of the isomerase-2 reaction are a mixture of 11-cis-ROL, 11-cis-REs, and all-trans-REs. Different

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Fig. 19.3. Rpe65 and isomerase-2 activities in chicken retina and RPE. (a) Isomerasespecific activities (pmol 11-cis-ROL per min per mg protein) in chicken retina and chicken RPE homogenates using the Rpe65 assay with all-trans-RP substrate. Rpe65 activity is almost undetectable in retina homogenates. (b) Isomerase-specific activities in chicken retina and chicken RPE homogenates using the isomerase-2 assay with alltrans-ROL and palm CoA substrates. Retinoids were saponified after assay incubation. Note the much higher activity in chicken retina versus RPE homogenates. Also note the difference in scale of specific activities between figure panels (a) and (b). During dissection, all visible traces of pigmented RPE material were removed from the retina samples. However, contaminating retina could not be removed from the RPE samples since retina is non-pigmented. Thus, RPE samples in this experiment probably contained contaminating retina material, while the retina samples were relatively free of RPE contamination.

fatty-acyl forms of retinol isomers co-elute during normalphase HPLC, which makes quantitation difficult. More accurate quantitation of isomerase-2 activity can be obtained by saponification (hydrolyzing in base) to yield the free retinols, which are readily separated by HPLC. References 1. Bunt-Milam, A.H., Saari, J.C. (1983) Immunocytochemical localization of two retinoid-binding proteins in vertebrate retina. J. Cell Biol. 97(3), 703–712. 2. Saari, J.C., Bredberg, D.L. (1987) Photochemistry and stereoselectivity of cellular retinaldehyde-binding protein from bovine retina. J. Biol. Chem. 262(16), 7618–7622. 3. Radu, R.A., et al. (2008) Retinal pigment epithelium-retinal G protein receptor-opsin mediates light-dependent translocation of all-

trans-retinyl esters for synthesis of visual chromophore in retinal pigment epithelial cells. J. Biol. Chem. 283(28), 19730–19738. 4. Eisenfeld, A.J., Bunt-Milam, A.H., Saari, J.C. (1985) Localization of retinoid-binding proteins in developing rat retina. Exp. Eye. Res. 41(3), 299–304. 5. Znoiko, S.L., et al. (2002) Identification of the RPE65 protein in mammalian cone photoreceptors. Invest. Ophthal. Vis. Sci. 43, 1604–1609.

Analysis of the Retinoid Isomerase Activities 6. Mata, N.L., et al. (2005) Chicken retinas contain a retinoid isomerase activity that catalyzes the direct conversion of all-transretinol to 1-cis-retinol. Biochemistry 44(35), 11715–11721. 7. Goldstein, E.B. (1967) Early receptor potential of the isolated frog (Rana pipiens) retina. Vision Res. 7, 837–845. 8. Goldstein, E.B., Wolf, B.M. (1973) Regeneration of the green-rod pigment in the isolated frog retina. Vision Res. 13, 527–534. 9. Hood, D.C., Hock, P.A. (1973) Recovery of cone receptor activity in the frog’s isolated retina. Vision Res. 13, 1943–1951. 10. Hood, D.C., Hock, P.A., Grover, B.G. (1973) Dark adaptation of the frog’s rods. Vision Res. 13, 1953–1963.

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11. Jones, G.J., et al. (1989) Retinoid requirements for recovery of sensitivity after visualpigment bleaching in isolated photoreceptors. Proc. Natl. Acad. Sci. USA 86(23), 9606–9610. 12. Das, S.R., et al. (1992) Muller cells of chicken retina synthesize 11-cis-retinol. Biochem. J. 285(Pt 3), 907–913. 13. Wang, J.S., et al. (2009) Intra-retinal visual cycle required for rapid and complete cone dark adaptation. Nat. Neurosci. 12(3), 295–302. 14. Mata, N.L., et al. (2002) Isomerization and oxidation of vitamin a in cone-dominant retinas. A novel pathway for visual-pigment regeneration in daylight. Neuron 36(1), 69–80.

Chapter 20 Techniques to Study Specific Cell-Surface Receptor-Mediated Cellular Vitamin A Uptake Riki Kawaguchi and Hui Sun Abstract STRA6 is a multitransmembrane domain protein that was recently identified as the cell-surface receptor for plasma retinol-binding protein (RBP), the vitamin A carrier protein in the blood. STRA6 binds to RBP with high affinity and mediates cellular uptake of vitamin A from RBP. It is not homologous to any known receptors, transporters, and channels, and it represents a new class of membrane transport protein. Consistent with the diverse physiological functions of vitamin A, STRA6 is widely expressed in diverse adult organs and throughout embryonic development. Mutations in human STRA6 that abolish its vitamin A uptake activity cause severe pathological phenotypes in many human organs including the eye, brain, lung, and heart. This chapter describes functional assays for STRA6 in live cells and on cellular membranes. These assays can be employed to study the mechanism of this new membrane transport mechanism and its roles in the physiology and pathology of many organs. Key words: STRA6, retinol-binding protein receptor, retinol, retinyl ester, vitamin A, retinoid, vitamin A uptake, HPLC.

1. Introduction Vitamin A has diverse biological functions (1–3). It exists in diverse forms, including alcohol, aldehyde, acid, and ester. Except for the ester, which is the storage form, all other forms of vitamin A are known to have biological activities. The acid form of vitamin A (retinoic acid) has the most diverse functions. Nuclear retinoic acid receptors regulate the transcription of a large number of genes (4, 5). In addition to its essential roles in embryonic development (6, 7), retinoic acid is also important in the function of many adult organs such as the nervous system (8), H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, DOI 10.1007/978-1-60327-325-1_20, © Springer Science+Business Media, LLC 2010

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the immune system (9, 10), the male and female reproductive systems (11, 12), the respiratory system (12, 13), and the skin (14). Retinoic acid was recently discovered to regulate protein translation in neurons (15, 16). The aldehyde form of vitamin A functions as the chromophore for visual pigments in the eye (17). It was also recently discovered to inhibit adipogenesis (18). The alcohol form of vitamin A serves as the substrate for retinolbinding protein (RBP) for delivery in the blood. In addition, the alcohol derivatives of vitamin A have distinct biological activities. For example, they control the growth of B lymphocytes (19) and function as survival factors in serum for fibroblasts (20). Retinol, but not retinoic acid, regulates BMP4 expression in male germ line cells (21) and maintains the pluripotency of embryonic stem cells (22). Plasma retinol-binding protein (RBP) is the principal carrier of vitamin A in the blood and is essential in mobilizing the hepatic vitamin A store (23–26). It was first proposed in the 1970s that there exists a cell-surface receptor that mediates vitamin A uptake from RBP on the retinal pigment epithelium and small intestinal cells (27–32). During the past three decades, there has been mounting evidence for the existence of RBP receptors on diverse types of tissues including the placenta (33–35), the choroid plexus (34, 36), Sertoli cells and peritubular cells of the testis (34, 37–40), macrophages (41), and skin (34, 42). There are also indirect pieces of evidence for the existence of an RBP receptor. For example, in an unbiased search for a serum factor that stimulates the growth of B cells, it was found that vitamin A/RBP complex (holo-RBP) is this factor (43). Using an unbiased strategy combining ligand-specific photo-crosslinking, highaffinity purification, and mass spectrometry, the RBP receptor was identified as STRA6, a multitransmembrane protein of previously unknown function (44) (Fig. 20.1). STRA6 binds to RBP with high affinity and specificity and mediates the uptake of vitamin A into the cell. STRA6 was originally identified as a retinoic acidstimulated gene in cancer cell lines (45, 46). STRA6 is widely expressed in embryonic development and adult organ systems (44, 45). Consistent with the diverse functions of vitamin A in human, STRA6 mutations cause severe pathological phenotypes including the absence of eyes (anophthalmia), mental retardation, congenital heart defects, and lung hyperplasia (47, 48). The point mutations identified in the human patients have been shown to abolish vitamin A uptake activity of STRA6 (49). In this chapter, we described techniques to study RBP receptor-mediated vitamin A uptake: a radioactive retinoid uptake assay, an HPLC-based retinol uptake assay, a cell-free retinoid uptake assay, and a method to quantitate of RBP binding to live cells. The advantage of the radioactive retinoid uptake assay is its high sensitivity. It does not distinguish between retinoids, but

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Fig. 20.1. Transmembrane topology of STRA6, the high-affinity cell-surface receptor for RBP (50). Transmembrane domains are indicated with Roman numerals. Missense mutations in human STRA6 associated with severe birth defects are indicated (47). Crystal structure of holo-RBP is based on structure 1HBP in Protein Data Bank.

measures total retinoid uptake. Although the HPLC-based assay is less sensitive, it can distinguish between retinoids. In addition, the HPLC-based assay can use serum as the source of holo-RBP and most closely mimics the in vivo uptake process due to the presence of transthyretin/RBP complex in the serum. The cellfree assay is based on the radioactive assay but is performed using cellular membranes. This assay makes it possible to acutely change the composition of the reaction. Quantitation of RBP binding to live cells is achieved using alkaline phosphatase-tagged RBP (APRBP). These assays have been successfully used for structural and functional analysis of STRA6’s role as the RBP receptor in mediating vitamin A uptake (44, 49, 50).

2. Materials 2.1. RBP Production

1. Human retinol-binding protein with a 6XHis tag on the N-terminus (His-RBP) cloned into the pET3a vector (New England Biolabs) 2. Competent BL-21 bacteria 3. Luria-Bertani (LB) media with 0.1 mg/ml carbenicillin

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4. 0.1 M IPTG (sterilized by filtration) 5. Complete protease inhibitors (Roche) 6. Phosphate buffered saline (PBS) 7. 20% Triton X-100 8. Sonicator 2.2. RBP Refolding

1. 7.5 M guanidine hydrochloride. 2. Sonicator. 3. 25 mM Tris–HCl, pH 9.0. 4. 30 mM cystine (prepared fresh): Dissolve 10.8 mg cystine in 50 μl of 1N NaOH. Heating at 37◦ C helps solubilization. Once cystine is completely dissolved, add water to 1.5 ml. 5. 300 mM Cysteine (prepared fresh). 6. Refolding buffer (prepared fresh): 25 mM Tris, pH 9.0, 0.3 mM cystine, 3.0 mM cysteine, 1 mM EDTA. 7. Amicon Ultra 15 concentrator, MWCO 10 K (Millipore). 8. 10 mM All-trans-retinol (ACROS): Dissolve 5.7 mg alltrans-retinol in 2 ml of 10 mM butylated hydroxytoluene (BHT) in ethanol. Keep the solution in dark. Store at −80◦ C. 9. 2 M dithiothreitol (DTT) (stored at −20◦ C in aliquots).

2.3. RBP Purification

1. Ni-NTA resin (Qiagen). 2. 10 mM Imidazole in PBS. 3. 100 mM Imidazole in PBS. 4. Dialysis tubing with MWCO of 10 kDa. 5. 25 mM Tris, pH 8.4, and 120 mM NaCl. 6. HPLC system with a photo-diode array detector (Agilent Technologies). 7. Weak anion exchange column AX-300 (Eprogen). 8. 25 mM Tris, pH 8.4. 9. 25 mM Tris, pH 8.4, and 2 N NaCl. 10. Nanodrop spectrophotometer.

2.4. Apo-RBP Production and Loading of 3 H-Retinol into Apo-RBP

1. Heptane. 2. Syringe with 22-gauge needle (Hamilton). 3. Nanodrop spectrophotometer. 4. [11,12-3 H]all-trans-retinol −20◦ C). 5. Ni–NTA resin (Qiagen). 6. 100 mM imidazole in PBS.

(PerkinElmer)

(stored

at

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1. Transfected COS cells (24 h after transfection). 2. 3 H-retinol/RBP. 3. Hank’s buffered salt solution (HBSS). 4. Serum-free medium (SFM). 5. 1% Triton-X100 in PBS. 6. Scintillation fluid (PerkinElmer). 7. Scintillation counter.

2.6. Cell-Free Retinoid Uptake Assay

1. PBS with protease inhibitors. 2. 3 H-retinol/RBP. 3. 2× reaction buffer: 3 H-retinol/RBP (40,000–50,000 CPM/reaction), 2.5 mg/ml BSA, 10 mM DTT, 500 mM sucrose, 1X PBS, and complete protease inhibitors. 4. Mechanical grinder (KONTES). 5. Syringe with 22-gauge needle (Hamilton). 6. Flask that connects to a vacuum source for radioactive waste. 7. Multiscreen vacuum manifold (Millipore). 8. MultiScreenHTS low protein binding 96-well filter plates (Millipore). 9. Scintillation fluid (PerkinElmer). 10. Scintillation counter.

2.7. HPLC-Based Retinoid Uptake Assays to Detect Retinyl Esters and Retinol

1. HPLC system with a photo-diode array detector (Agilent Technologies). 2. ZORBAX Eclipse XDB-C18 column, 5 μm, 4.6 mm × 150 mm (Agilent Technologies). 3. Purified holo-RBP. 4. Pooled human sera (Innovative Research). 5. Serum-free medium (SFM). 6. Hank’s buffered salt solution (HBSS). 7. PBS with 5 mM EDTA. 8. Mechanical grinder (KONTES). 9. 10 mM Butylated hydroxytoluene (BHT) in ethanol. 10. Hexane. 11. Nitrogen tank. 12. Ethylacetate (HPLC grade). 13. Methanol (HPLC grade). 14. Dark room with red lights.

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2.8. AP-RBP Production and Binding

1. AP-RBP fusion protein in SFM. 2. Hank’s buffered salt solution (HBSS). 3. Serum-free medium (SFM). 4. 2% Formaldehyde in PBS (freshly made). 5. 65◦ C incubator. 6. AP buffer: 0.1 M Tris, pH 9.5, 0.1 M NaCl, and 50 mM MgCl2 7. BCIP (Roche). 8. NBT (Roche). 9. pNPP (Sigma). 10. 3 N NaOH. 11. Microplate spectrophotometer.

3. Methods 3.1. Production of His-Tagged RBP (His-RBP) in Bacteria

1. Tag human cDNA for RBP on the N-terminus with 6XHis tag and clone it into the pET3a vector (see Note 1). Transform the construct into competent BL-21 bacteria. 2. Grow transformed BL-21 in 50 ml LB medium containing 0.1 mg/ml carbenicillin at 37◦ C with vigorous shaking. 3. Add 0.5 ml of 0.1 M IPTG to the culture medium when the O.D. at 660 nm reaches 0.1. Grow further for 5 h (see Note 2). 4. Pellet the cells by centrifugation at 10,000×g for 20 min. 5. Wash the cells with 50 ml PBS, pellet again, and resuspend in cold 20 ml PBS with complete protease inhibitors. 6. Sonicate the cells on ice for 1 min followed by 1 min cooling. This process is repeated four times. 7. Add Triton X-100 to a final concentration of 0.1%. 8. Freeze and thaw the cell lysate twice. 9. Pellet down the inclusion bodies at 20,000×g for 30 min at 4◦ C. RBP expressed in Escherichia coli is mainly found in inclusion bodies (see Note 3).

3.2. Solubilization and Refolding of RBP

1. Solubilize the inclusion body pellet in 20 ml 7.5 M guanidine hydrochloride. Brief sonication helps in solubilization (see Note 3). 2. After adding half volume (10 ml) of 25 mM Tris buffer, pH 9.0, add DTT to a final concentration of 10 mM (see Note 4). Incubate overnight at room temperature to fully reduce and denature proteins.

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3. Centrifuge the protein solution to remove guanidineinsoluble material at 16,000×g for 30 min. Split the solution into two batches of 15 ml. Freeze one batch at −20◦ C (see Note 5). 4. Bring all-trans-retinol stock to room temperature, keeping it in the dark (see Note 6). Prepare four volumes (60 ml) of refolding buffer (25 mM Tris, pH 9.0, 0.3 mM cystine, 3.0 mM cysteine, 1 mM EDTA) (see Note 7). 5. Degas the refolding buffer for 15 min and chill on ice. At the same time, cool both the protein solution and the refolding buffer on ice since high temperature can lead to protein aggregation. 6. Add 600 μl of 10 mM all-trans-retinol to the refolding buffer (to a final concentration of 1 mM) right before the refolding reaction. 7. Add refolding buffer to the protein solution dropwise with vigorous stirring of the protein solution (see Note 8). Carry out the refolding in the dark on ice or at 4◦ C for 5 h. 8. Spin down the refolding reaction at 24,000×g for 30 min at 4◦ C to remove protein aggregates. Misfolded RBP tends to aggregate together with correctly folded RBP. Therefore, it is essential to remove protein aggregates as soon as refolding is done. 9. Concentrate the supernatant solution using Amicon Ultra 15 concentrator (MWCO 10 K) to about 10 ml. Do not concentrate the refolded solution more than 10-fold. Too much concentrating causes protein aggregation and lowers the final yield of correctly folded His-RBP. 10. Dilute the concentrated sample with PBS to 100 ml to reduce the DTT and EDTA concentration (see Note 9). 11. Spin the solution one more time at 24,000×g for 20 min at 4◦ C, and transfer the supernatant to two 50-ml tubes. Add 1 ml of Ni-NTA resin to each tube and rotate the solution with the resin for 2 h (see Note 10). Longer incubation may increase yield slightly. Adding protease inhibitors may help; however, no major degradation has been observed without them. 12. Wash the resin with at least 20 column volumes of 10 mM imidazole in PBS before eluting His-RBP with 100 mM imidazole in PBS (see Note 11). 3.3. HPLC Purification of Refolded Holo-RBP

1. Dialyze His-RBP purified from the Ni-NTA resin against 25 mM Tris, pH 8.4, and 120 mM NaCl. 2. Set up HPLC with weak anion exchange column AX-300 (see Note 12).

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Fig. 20.2. Monitoring of the full absorption spectrum of the entire HPLC run during the holo-RBP purification. The peaks for holo-RBP and apo-RBP are indicated.

3. Clear samples for protein purification on HPLC by centrifugation at 16,000×g for 10 min at 4◦ C before each run. 4. Separate proteins by a NaCl step gradient (220 mM 10 min, 360 mM 15 min, and 1,000 mM 15 min) using 25 mM Tris (pH 8.4) as mobile phase at 1 ml/min. As NaCl concentration rises, holo-His-RBP is released whereas apo-His-RBP and misfolded RBP stay bound to the column. Monitor the full absorption spectrum of entire HPLC run (Fig. 20.2) (see Note 13). 5. Recover holo-RBP from the fractions at 360 mM NaCl. At 1,000 mM NaCl, all His-RBP bound to the column is released. 6. Fractions containing holo-RBP are pooled, concentrated, and dialyzed against PBS. 7. Purified holo-RBP can be quantified by Nanodrop spectrophotometer (see Note 14). 8. Store HPLC-purified RBP at 4◦ C (see Note 15). 3.4. Production of Apo-RBP and Loading of 3 H-Retinol onto Apo-RBP

Apo-RBP is prepared by removing retinol from holo-RBP using organic solvents such as hexane or heptane. 1. Add an equal volume of heptane to purified holo-RBP and mix gently by rotating overnight at 4◦ C. Harsh mixing will result in loss of the protein due to aggregation. 2. Centrifuge at 16,000×g for 10 min at 4◦ C to separate the aqueous phase and organic phase. Use a 22-gauge needle to carefully transfer the aqueous phase (bottom) to a new tube. A protein precipitate may form at the interphase. Be careful to avoid the precipitate. 3. Repeat this process three more times with a 3-h heptane incubation for each.

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4. Centrifuge the aqueous phase at 16,000×g for 10 min at 4◦ C to remove any insoluble materials. Check retinol depletion from His-RBP using a Nanodrop spectrophotometer. Apo-RBP should not show an absorption peak at 330 nm. If the protein solution still has significant absorption at 330 nm, repeat the heptane extraction until the absorption at 330 nm is the same as the baseline. 5. Add 40 μl of 15–30 μM3 H-all-trans-retinol stock to 10–20 μg of apo-His-RBP in 960 μl of PBS (see Note 16). 6. Rotate overnight at 4◦ C and purify His-RBP with 15 μl of Ni-NTA resin. 7. Wash the resin with 1 ml PBS four times and elute the 3 Hretinol/RBP in 200–1,000 μl of 100 mM imidazole in PBS. Measure the radioactivity of the eluted materials by scintillation. 3.5. Retinol Uptake Assay Based on 3 H-Retinol/RBP

The following protocol is designed for COS cells grown on 24well dishes (see Note 17). An example of this experiment is shown in Fig. 20.3. The volume of each buffer can be adjusted proportionally for cells grown on wells or dishes of larger areas.

Fig. 20.3. Vitamin A uptake activity from holo-RBP for human STRA6 mutants associated with severe birth defects (47, 49). Locations of these mutations in the transmembrane topology model of STRA6 are shown in Fig. 20.1. The activity of wild-type STRA6 is defined as 100%.

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1. At 24 h after transfection, media from each well is removed by vacuum suction. The cells in each well are washed with 500 μl HBSS. 2. Remove HBSS and add 250 μl of SFM containing 3 Hretinol/RBP (see Note 18). 3. After incubating for desired length of time at 37◦ C (e.g., 1 h), remove the supernatant and wash the cells in each well with 500 μl HBSS (see Note 19). 4. Lyse the cells by adding 500 μl 1% Triton X-100 in PBS to each well. 5. Transfer the cell lysate to a scintillation vial. 6. Repeat steps 4 and 5 for each well before counting with a scintillation counter (see Note 20). 3.6. A Cell-Free Assay for 3 H-Retinol Uptake

1. To prepare crude membranes, break up cell pellets on ice with a mechanical grinder in PBS with protease inhibitors (see Note 21). 2. Dilute the cell lysates in PBS with protease inhibitors. After centrifugation at 1,000×g for 3 min at 4◦ C to remove cell nuclei, centrifuge the lysate at 16,000×g for 30 min at 4◦ C to pellet the cell membranes. 3. Resuspend the membrane pellets in PBS (>50 μl) and break up the pellets by repeated passage through a 22gauge needle (see Note 22). 4. Prepare 2× reaction buffer that contains 3 H-retinol-RBP solution and add an equal volume to the membrane suspension. A typical final reaction volume is 100 μl. 5. Incubate for a defined amount of time (e.g., 1 h) at 37◦ C. 6. During the incubation, assemble the 96-well filtration plate and multiscreen vacuum manifold according to the manufacturer’s protocol. Open valve and test vacuum suction (see Note 23). 7. Wet each well with 10 μl PBS. Seal unused wells. 8. Mix samples by pipetting and add them to wells. Once the liquid is removed by suction, add 200 μl PBS. Repeat this wash to each well once more. Use of a multichannel pipettor is recommended when dealing with many samples. 9. Let the membrane dry by keeping suction force for 10 min. Then remove the back support from the MultiscreenHTS 96-well plate. Place the multiscreen plate on a few layers of Kimwipes to remove liquid on the backside. Let the plate dry for another 15 min. 10. Punch the membrane screen out of the plate and place it in a scintillation vial. Count radioactivity by scintillation.

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The following protocol is designed for transfected COS cells (see Note 24): 1. At 6 h after transfection, remove medium and quickly wash cells once with HBSS before culturing the cells in SFM at 37◦ C (see Note 25). 2. At 18 h after culturing in SFM, remove SFM and add holoRBP freshly diluted in SFM. The source of holo-RBP can be either purified holo-RBP or normal human serum, which contains RBP in the form of holo-RBP/transthyretin complex (see Note 26). 3. After incubation at 37◦ C for a defined period of time (e.g., 1 h), remove the medium and wash the cells briefly with 10 ml PBS. 4. Add 1.5 ml PBS containing 5 mM EDTA. Incubate for 2 min. 5. Detach cells by pipetting and collect cells in microcentrifuge tubes. 6. Pellet cells at 1,000×g for 3 min. Repeat steps 4 and 5 and combine cells. 7. Store the cell pellets at −80◦ C before retinoid extraction.

3.8. HPLC Analysis of Retinyl Ester and Retinol Contents in the Cells

All steps in the following procedure are done under red light: 1. 1. Resuspend cell pellet in 50 μl PBS in the presence of protease inhibitors. The following protocol describes experiments at this scale. 2. Add 50 μl of 10 mM BHT in ethanol to the cell lysate. Vortex for 30 s. 3. Add 500 μl of hexane. Vortex for 5 min. 4. Pellet down cellular debris at 5,000×g for 5 min. 5. Carefully remove and save the upper phase (hexane). 6. Repeat steps 3–5 twice and combine all hexane extracts. 7. Dry the hexane extractable material under nitrogen in the dark. 8. Solubilize the dried material in 450 μl of methanol by vortexing for 1 min. 9. For retinol analysis, add 50 μl water to the sample and filter the sample before HPLC (see Note 27). Separate samples using ZORBAX Eclipse XDB-C18 column (5 μm, 4.6 mm × 150 mm) at 1 ml/min using 90% methanol as the mobile phase. The retinol peak at 325 nm appears at around 7 min.

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Fig. 20.4. HPLC assay for retinyl ester uptake using transfected or untransfected COS-1 cells that were incubated with 20% normal human serum for 4 h. Retinyl ester was extracted and analyzed by HPLC. Mobile phase was changed linearly from 100% methanol to 100% ethylacetate. Retinyl palmitate peaks are indicated.

10. For retinyl ester analysis, separate filtered samples in 100% methanol using a linear gradient (100% methanol at time zero and 100% ethylacetate at 10 min). Flow rate is fixed at 1 ml/min. The same column for retinol analysis is used. The retinyl ester peak at 325 nm appears at around 8 min (Fig. 20.4) (see Note 28). 11. Peak areas for retinoid HPLC analysis are usually normalized by the protein peak by following the 280 nm absorbance. Using the conditions of this protocol, the protein peak appears at 2.3 min for retinyl ester or at 4.5 min for retinol analysis (see Note 29). 3.9. Visualization and Quantitation of the Binding of AP-RBP to Live Cells

The following protocol is designed to visualize AP-RBP binding to transfected or untransfected COS cells (see Note 30): 1. Wash COS cells grown on gelatin-coated coverslips once with PBS. 2. Incubate with AP-RBP diluted in SFM for 1 h at room temperature. 3. After washing the cells twice with HBSS, fix in fresh 2% formaldehyde in PBS at room temperature for 10 min. 4. After three more PBS washes, heat the cells in PBS for 1 h at 65◦ C. 5. After washing the cells once with AP buffer, perform the AP color reactions in the AP buffer containing 165 μg/ml of BCIP and 330 μg/ml of NBT for 1 h at room temperature.

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Fig. 20.5. Quantitation of RBP binding activities of human STRA6 mutants associated with severe birth defects (47, 49). Locations of these mutations in the transmembrane topology model of STRA6 are shown in Fig. 20.1. The activity of wild-type STRA6 is defined as 100%.

The following protocol is the liquid AP assay designed to quantitate AP-RBP binding to transfected or untransfected COS cells. An example of this experiment is shown in Fig. 20.5. 1. Wash COS cells grown on 12-well cell culture plates once with HBSS and incubated with AP-RBP diluted in SFM at 37◦◦ C for 1 h. 2. Stop the reactions by quickly washing cells three times with HBSS. 3. Lyse the cells in 150 μl of cold PBS containing 1% Triton X-100 and protease inhibitors per well. 4. Centrifuge the cell lysates at 3,000×g at 4◦ C for 5 min. 5. Remove the supernatants and heat at 65◦ C for 1 h. 6. Mix 50 μl of the heated lysate with 200 μl of pNPP and incubate at 37◦ C for 1 h for the AP color reaction. Use the lysate from untransfected cells as the negative control. 7. Stop the reactions by adding 50 μl 3 M NaOH and transfer to a 96-well plate for reading in a microplate reader at 405 nm.

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4. Notes 1. Tagging RBP on the N-terminus does not interfere with its interaction with STRA6, but a tag on the C-terminus may interfere with this interaction (44). Therefore, the 6XHis tag is engineered into the N-terminus of RBP. 2. IPTG may or may not be necessary in RBP induction. Basal RBP expression is already high (Fig. 20.2). LB is better than rich media for RBP production. 3. As an extracellular protein, native RBP has three pairs of disulfide bonds. Bacteria express RBP intracellularly and cannot properly form the correct disulfide bonds. For this reason, RBP expressed in E. coli is mainly found in inclusion body as misfolded proteins that are mostly inactive and do not bind retinol. This misfolded RBP needs to be denatured and refolded. Good refolding is critical for producing high-quality holo-RBP. Without good refolding, even HPLC cannot purify holo-RBP 100% loaded with all-transretinol. 4. The concentration of DTT in the solubilizing solution is critical. Excess DTT can inhibit the oxidation reaction during the refolding reaction and makes refolded – RBP unstable without disulfide bonds. 5. Increasing the scale of the refolding reaction can decrease the yield of correctly folded RBP. 6. The refolding solution includes all-trans-retinol, which helps proper refolding of RBP. Good quality retinol is essential during the refolding reaction. Without all-transretinol, refolding efficiency is very low (<10%). Once solubilized in ethanol, add BHT to 10 mM to prevent oxidation, seal the tube tightly, and store the unused retinol at −80◦ C. 7. Cystine and cysteine solutions need to be made fresh. Cystine is not easily solubilized without high pH and heat. Do not add water directly to solubilize cystine. To make the 30 mM cystine stock solution, dissolve 10.8 mg cystine in 50 μl of 1N NaOH. Heating at 37◦ C helps solubilization. Once cystine is completely dissolved add water to 1.5 ml. 8. Refolding buffer containing cystine and cysteine has to be added slowly at the start of the refolding reaction when the solution is being stirred. The slow addition of refolding buffer prevents the dramatic reduction of local guanidine concentration. As guanidine concentration is gradually

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diluted, RBP refolding starts. Refolding buffer contains cystine and cysteine at a ratio of 1:10. Cystine and cysteine facilitate the shuffling reaction by reducing and oxidizing the disulfide bonds in RBP. 9. High DTT and EDTA concentrations can reduce the efficiency of subsequent His-RBP purification using Ni-NTA resin. Dialysis to remove DTT and EDTA is not recommended because long buffer exchange tends to cause serious protein aggregation. 10. Ni-NTA resin binding of refolded RBP should be done on the same day as the refolding reaction. Refolded RBP solution without dilution tends to aggregate over an extended time period. 11. The typical yield at this step is >20 mg/100 ml of bacteria culture. In a typical preparation with successful refolding, 75–80% of His-RBP should be bound with retinol based on the 280 and 330 nm readings. 12. Since His-RBP has a pI of 6.47, a basic buffer is used to keep the protein in the negatively charged state. At physiological ionic strength, His-RBP binds to weak anion exchange column. It is important to calibrate the pH meter to prepare the mobile phase because the binding of RBP to the column is very sensitive to pH. High pH (pH> 8.4) can reduce the protein-binding capacity of the column and low pH (between pH 6.47 and 8.4) can make proteins bind too strongly to elute. 13. Holo-RBP can be monitored during purification either by its characteristic absorption at 330 nm or by the fluorescence intensity of the bound retinol. 14. Accurate quantitation of holo-RBP is not possible by spectrophotometer if free retinol is present in the solution. Even for purified holo-RBP, retinol in holo-RBP consistently causes a problem in accurate quantitation because the presence of retinol in RBP lowers the absorbance baseline significantly. Therefore, the baseline needs to be adjusted to reflect the presence of retinol. On a NanoDrop spectrophotometer, use PBS as baseline if holo-RBP is in PBS. Then read the absorbance of holo-RBP solution at 280, 330, and 500 nm. Note that the 500 nm value is reduced due to the presence of retinol in the solution. Add the reduced absorbance value at 550 nm to the values 280 and 330 nm to get the correct absorbance values. Alternatively, purified holo-RBP can be quantified using retinol fluorescence using holo-RBP of known concentrations. Although fluorescence is more sensitive, it cannot reveal how well retinol is loaded into RBP.

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15. High-quality holo-RBP is very stable and can be stored at 4◦ C in PBS for years without any loss or degradation of retinol. Freezing HPLC-purified RBP is not recommended, as it lowers the 330/280 ratio upon thawing. If RBP is frozen, centrifuge at 16,000×g for 10 min to remove any protein aggregates. 16. Apo-His-RBP tends to denature in the presence of a high concentration of ethanol. Therefore, the ethanol concentration should not exceed 4% in the loading reaction. The [11,12-3 H]retinol purchased from PerkinElmer has a specific activity of 30–60 Ci/mmol and a concentration of 1 mCi/ml or about 15–30 μM (in ethanol). At this concentration, the maximum concentration of 3 H-retinol is 0.6–1.2 μM in the loading reaction if 4% of the 3 H-retinol stock in ethanol is added. To achieve higher 3 H-retinol incorporation, the stock 3 H-retinol solution can be concentrated before use. Alternatively, purify His-RBP and repeat the loading reaction with fresh 3 H-retinol. 17. Since retinol uptake assays on live cells involve multiple washes, COS cells are advantageous because they do not detach as easily as HEK293 cells. HEK293 cells are more easily transfected and can be used for membrane-based assays. 18. For 24-well assays, stock 3 H-retinol/RBP is diluted in SFM so that each well contains 250 μl of SFM and 40,000– 50,000 CPM of 3 H-retinol-RBP. The sensitivity of the radioactive assay allows the use of 3 H-retinol-RBP at a much lower concentration (e.g., 3 nM) than the Kd of the RBP/STRA6 interaction. 19. The presence of divalent ions in HBSS makes it a better solution for live cell washes than PBS because it can minimize cell loss due to the wash. It helps to visually monitor cell attachment during the wash process. If the experiment is designed to account for 3 H-retinol/RBP bound to cell surface, it is important to perform the wash quickly since the RBP/STRA6 interaction is transient (e.g., wash two wells at a time). 20. Compared with HPLC-based assays, radioactive assays use much lower concentrations of holo-RBP (3 Hretinol/RBP) and have overall lower concentrations of proteins. Therefore, non-specific sticking of retinol/RBP to plastic dishes is a more significant source of background “uptake” signal in radioactive retinol-based assays, while it is negligible for HPLC-based assays. For a typical 3 Hretinol/RBP-based assay, this background signal is less than 10% of the real uptake signal. This background signal can

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be easily measured by incubating 3 H-retinol/RBP with empty wells that have been incubated with culture media but without cells. The equivalent amount of background in wells with cells can be calculated by taking into account of the confluence of the cells on the bottom of this dish and the area of the side wall in contact with the medium. 21. The cell-free assay for retinol uptake has the advantage of using more concentrated STRA6 in a small volume and allowing for the addition of soluble factors during the reaction. However, cellular membrane normally needs to be spun down to wash off non-specific-bound 3 H-retinolRBP. Repeated centrifugation can be time consuming. A filtering device can greatly reduce the washing time for cellular membrane-based retinol uptake. 22. Resuspend the membrane in 50–100 μl of PBS per reaction. Avoid bubbles during needle passage. Since a 96-well filtration device is used in this assay during the wash step, it is important not to use membranes prepared from more than one-quarter of a confluent 100-mm dish of cells per well (per reaction). Too much membrane can clog the filter and make it impossible to wash the membranes. 23. To provide constant vacuum pressure for the multiscreen system, cap the top of each well that will not be used. Alternatively, once PBS is added to wash each well, place a cover over the multiscreen plate and seal the side briefly. 24. Radioactive assays and HPLC assays for retinoid uptake require different scales of experiments due to the difference in sensitivity of detection. Radioactive assays can be performed on 24-well plates due to their high sensitivity. An HPLC-based assay to detect retinol uptake needs to be done at a much larger scale than an HPCL-based assay to detect retinyl esters. HPLC to detect retinyl esters can be performed on six-well plates. In contrast, a 100-mm dish is necessary for HPLC-based assay to detect retinol accumulation. 25. It is ideal to perform the retinoid uptake assay 24 h after transfection, since this is the time at which the transfected proteins just reach the peak of expression. However, fetal bovine serum used in cell culture does contain retinol-binding protein. Prolonged incubation after transfection will result in significant background retinoid uptake. The background level of uptake before the assay can be easily detected in HPLC by measuring retinoid levels without adding exogenous retinol-binding protein or human serum. Changing the culture media to SFM 6 h after transfection can reduce background retinoid uptake to non-detectable levels.

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26. Normal human serum contains high concentrations of holo-RBP (∼2 μM), which is much higher than the Kd of STRA6/RBP binding (∼50 nM). Therefore, retinol uptake from blood samples usually uses serum diluted in SFM (e.g., 25% serum). 27. Samples for retinoid analysis are filtered. Use of a guard column is preferred for good long-term performance of the column. 28. Flush the entire HPLC system with water if you do not plan to run HPLC for more than a few days. Salt precipitation will damage the pump. 29. An alternative way to normalize is to spike a fixed amount of retinyl acetate into the starting materials as an external control, and use the peak areas for retinyl acetate to normalize retinoid extraction efficiency across the samples. For absolute quantitation, a standard curve is made with stock retinyl palmitate or retinol solution. 30. AP fusion is an established method to label secreted proteins and study their interactions with cell-surface receptors (51). In this system, the GPI anchor of human placental AP was removed to make it a secreted protein. AP tagged at the N-terminus of RBP (AP-RBP) does not interfere with its interaction with STRA6 (44). To produce APRBP, COS cells are transfected with AP-RBP cDNA cloned into a mammalian expression vector. COS cell is preferred due to its strong attachment to the culture dish. At 12–24 h after transfection, the media is changed to SFM. AP-RBP fusion protein can be harvested from the supernatant of transfected cells in SFM 48 h later. If purified AP-RBP is desired, a 6XHis tag can be inserted between AP and RBP. This tag allows convenient purification of AP-RBP from the SFM. Quantitation of AP-RBP concentration can be performed by comparing the AP signal in AP-RBP with AP proteins with known concentrations. Detection of AP signal after AP-RBP binding is an effective method to quantitative study of the interaction between RBP and STRA6. Since this AP is heat resistant, heating is an effective way to eliminate endogenous AP activity. References 1. Blomhoff, R. (1994) Overview of vitamin A metabolism and function. In: Blomhoff, R. (ed.), Vitamin A in Health and Disease, Marcel Dekker, Inc., New York, Basel, Hong Kong, pp. 1–35. 2. Ross, A.C., Gardner, E.M. (1994) The function of vitamin A in cellular growth and dif-

ferentiation, and its roles during pregnancy and lactation. Adv. Exp. Med. Biol. 352, 187–200. 3. Napoli, J.L. (1996) Biochemical pathways of retinoid transport, metabolism, and signal transduction. Clin. Immunol. Immunopathol. 80, S52–S62.

Specific Cell-Surface Receptor-Mediated Cellular Vitamin A Uptake 4. Evans, R.M. (1994) The molecular basis of signaling by vitamin A and its metabolites. Harvey Lect. 90, 105–117. 5. Chambon, P. (1996) A decade of molecular biology of retinoic acid receptors. FASEB J. 10, 940–954. 6. Maden, M. (1994) Role of retinoids in embryonic development. In: Blomhoff, R. (ed.), Vitamin A in Health and Disease, Marcel Dekker, Inc., New York, Basel, Hong Kong. 7. Duester, G. (2008) Retinoic acid synthesis and signaling during early organogenesis. Cell 134, 921–931. 8. Drager, U.C. (2006) Retinoic acid signaling in the functioning brain. Sci STKE 2006, pe10. 9. Blomhoff, H.K., Smeland, E.B. (1994) Role of retinoids in normal hematopoiesis and the immune system. In: Blomhoff, R. (ed.), Vitamin A in Health and Disease, Marcel Dekker, Inc., pp. 451–484 10. Stephensen, C.B. (2001) Vitamin A, infection, and immune function. Annu. Rev. Nutr. 21, 167–192. 11. Chung, S.S., Wolgemuth, D.J. (2004) Role of retinoid signaling in the regulation of spermatogenesis. Cytogenet. Genome Res. 105, 189–202. 12. Wolbach, S.R., Howe, P.R. (1925) Tissue change following deprivation of fat-soluble A vitamin. J. Exp. Med. 42, 753–777. 13. Biesalski, H.K. (2003) The significance of vitamin A for the development and function of the lung. Forum Nutr. 56, 37–40. 14. Vahlquist, A. (1994) Role of retinoids in normal and diseased skin. In: Blomhoff, R. (ed.), Vitamin A in Health and Disease, Marcel Dekker, Inc., New York, Basel, Hong Kong, pp 365–424. 15. Chen, N., Onisko, B., Napoli, J.L. (2008) The nuclear transcription factor RARalpha associates with neuronal RNA granules and suppresses translation. J. Biol. Chem. 283, 20841–20847. 16. Aoto, J., Nam, C.I., Poon, M.M., Ting, P., Chen, L. (2008) Synaptic signaling by alltrans retinoic acid in homeostatic synaptic plasticity. Neuron 60, 308–320. 17. Travis, G.H., Golczak, M., Moise, A.R., Palczewski, K. (2007) Diseases caused by defects in the visual cycle: Retinoids as potential therapeutic agents. Annu. Rev. Pharmacol. Toxicol. 47, 469–512. 18. Ziouzenkova, O., Orasanu, G., Sharlach, M., Akiyama, T.E., Berger, J.P., Viereck, J., Hamilton, J.A., Tang, G., Dolnikowski, G.G., Vogel, S., Duester, G., Plutzky, J. (2007) Retinaldehyde represses adipogene-

19.

20.

21.

22.

23. 24.

25.

26.

27.

28.

29.

30.

359

sis and diet-induced obesity. Nat. Med. 13, 695–702. Buck, J., Myc, A., Garbe, A., Cathomas, G. (1991) Differences in the action and metabolism between retinol and retinoic acid in B lymphocytes. J. Cell Biol. 115, 851–859. Chen, Y., Derguini, F., Buck, J. (1997) Vitamin A in serum is a survival factor for fibroblasts. Proc. Natl. Acad. Sci. USA 94, 10205–10208. Baleato, R.M., Aitken, R.J., Roman, S.D. (2005) Vitamin A regulation of BMP4 expression in the male germ line. Dev. Biol. 286, 78–90. Chen, L., Khillan, J.S. (2008) Promotion of feeder-independent self-renewal of embryonic stem cells by retinol (vitamin A). Stem Cells 26, 1858–1864. Blomhoff, R., Green, M.H., Berg, T., Norum, K.R. (1990) Transport and storage of vitamin A. Science 250, 399–404. Newcomer, M.E., Ong, D.E. (2000) Plasma retinol binding protein: Structure and function of the prototypic lipocalin. Biochim. Biophys. Acta 1482, 57–64. Zanotti, G., Berni, R. (2004) Plasma retinolbinding protein: Structure and interactions with retinol, retinoids, and transthyretin. Vitam. Horm. 69, 271–295. Quadro, L., Hamberger, L., Gottesman, M.E., Wang, F., Colantuoni, V., Blaner, W.S., Mendelsohn, C.L. (2005) Pathways of vitamin A delivery to the embryo: Insights from a new tunable model of embryonic vitamin A deficiency. Endocrinology 146, 4479–4490. Bok, D., Heller, J. (1976) Transport of retinol from the blood to the retina: An autoradiographic study of the pigment epithelial cell surface receptor for plasma retinol-binding protein. Exp. Eye Res. 22, 395–402. Heller, J. (1975) Interactions of plasma retinol-binding protein with its receptor. Specific binding of bovine and human retinol-binding protein to pigment epithelium cells from bovine eyes. J. Biol. Chem. 250, 3613–3619. Heller, J., Bok, D. (1976) Transport of retinol from the blood to the retina: Involvement of high molecular weight lipoproteins as intracellular carriers. Exp. Eye Res. 22, 403–410. Rask, L., Peterson, P.A. (1976) In vitro uptake of vitamin A from the retinol-binding plasma protein to mucosal epithelial cells from the monkey s small intestine. J. Biol. Chem. 251, 6360–6366.

360

Kawaguchi and Sun

31. Maraini, G., Gozzoli, F. (1975) Binding of retinol to isolated retinal pigment epithelium in the presence and absence of retinolbinding protein. Invest. Ophthalmol. 14, 785–787. 32. Chen, C.C., Heller, J. (1977) Uptake of retinol and retinoic acid from serum retinolbinding protein by retinal pigment epithelial cells. J. Biol. Chem. 252, 5216–5221. 33. Sivaprasadarao, A., Findlay, J.B. (1988) The interaction of retinol-binding protein with its plasma-membrane receptor. Biochem. J. 255, 561–569. 34. Smeland, S., Bjerknes, T., Malaba, L., Eskild, W., Norum, K.R., Blomhoff, R. (1995) Tissue distribution of the receptor for plasma retinol-binding protein. Biochem. J. 305(Pt 2), 419–424. 35. Sivaprasadarao, A., Boudjelal, M., Findlay, J.B. (1994) Solubilization and purification of the retinol-binding protein receptor from human placental membranes. Biochem. J. 302(Pt 1), 245–251. 36. MacDonald, P.N., Bok, D., Ong, D.E. (1990) Localization of cellular retinolbinding protein and retinol-binding protein in cells comprising the blood-brain barrier of rat and human. Proc. Natl. Acad. Sci. USA 87, 4265–4269. 37. Shingleton, J.L., Skinner, M.K., Ong, D.E. (1989) Characteristics of retinol accumulation from serum retinol-binding protein by cultured Sertoli cells, Biochemistry 28, 9641–9647. 38. Bishop, P.D., Griswold, M.D. (1987) Uptake and metabolism of retinol in cultured Sertoli cells: Evidence for a kinetic model. Biochemistry 26, 7511–7518. 39. Bhat, M.K., Cama, H.R. (1979) Gonadal cell surface receptor for plasma retinol-binding protein. A method for its radioassay and studies on its level during spermatogenesis. Biochim. Biophys. Acta 587, 273–281. 40. Davis, J.T., Ong, D.E. (1995) Retinol processing by the peritubular cell from rat testis. Biol. Reprod. 52, 356–364. 41. Hagen, E., Myhre, A.M., Smeland, S., Halvorsen, B., Norum, K.R., Blomhoff, R. (1999) Uptake of vitamin A in macrophages from physiologic transport proteins: Role of retinol-binding protein and chylomicron remnants. J. Nutr. Biochem. 10, 345–352. 42. Torma, H., Vahlquist, A. (1984) Vitamin A uptake by human skin in vitro. Arch. Dermatol. Res. 276, 390–395. 43. Buck, J., Ritter, G., Dannecker, L., Katta, V., Cohen, S.L., Chait, B.T., Hammerling, U. (1990) Retinol is essential for growth of

44.

45.

46.

47.

48.

49.

activated human B cells. J. Exp. Med. 171, 1613–1624. Kawaguchi, R., Yu, J., Honda, J., Hu, J., Whitelegge, J., Ping, P., Wiita, P., Bok, D., Sun, H. (2007) A membrane receptor for retinol binding protein mediates cellular uptake of vitamin A. Science 315, 820–825. Bouillet, P., Sapin, V., Chazaud, C., Messaddeq, N., Decimo, D., Dolle, P., Chambon, P. (1997) Developmental expression pattern of Stra6, a retinoic acidresponsive gene encoding a new type of membrane protein. Mech. Dev. 63, 173–186. Szeto, W., Jiang, W., Tice, D.A., Rubinfeld, B., Hollingshead, P.G., Fong, S.E., Dugger, D.L., Pham, T., Yansura, D.G., Wong, T.A., Grimaldi, J.C., Corpuz, R.T., Singh, J.S., Frantz, G.D., Devaux, B., Crowley, C.W., Schwall, R.H., Eberhard, D.A., Rastelli, L., Polakis, P., Pennica, D. (2001) Overexpression of the retinoic acid-responsive gene Stra6 in human cancers and its synergistic induction by Wnt1 and retinoic acid. Cancer Res. 61, 4197–4205. Pasutto, F., Sticht, H., Hammersen, G., Gillessen-Kaesbach, G., Fitzpatrick, D.R., Nurnberg, G., Brasch, F., SchirmerZimmermann, H., Tolmie, J.L., Chitayat, D., Houge, G., Fernandez-Martinez, L., Keating, S., Mortier, G., Hennekam, R.C., von der Wense, A., Slavotinek, A., Meinecke, P., Bitoun, P., Becker, C., Nurnberg, P., Reis, A., Rauch, A. (2007) Mutations in STRA6 cause a broad spectrum of malformations including anophthalmia, congenital heart defects, diaphragmatic hernia, alveolar capillary dysplasia, lung hypoplasia, and mental retardation. Am. J. Hum. Genet. 80, 550–560. Golzio, C., Martinovic-Bouriel, J., Thomas, S., Mougou-Zrelli, S., GrattaglianoBessieres, B., Bonniere, M., Delahaye, S., Munnich, A., Encha-Razavi, F., Lyonnet, S., Vekemans, M., Attie-Bitach, T., Etchevers, H.C. (2007) Matthew-Wood syndrome is caused by truncating mutations in the retinol-binding protein receptor gene STRA6. Am. J. Hum. Genet. 80, 1179–1187. Kawaguchi, R., Yu, J., Wiita, P., Honda, J., Sun, H. (2008) An essential ligandbinding domain in the membrane receptor for retinol-binding protein revealed by large-scale mutagenesis and a human polymorphism. J. Biol. Chem. 283, 15160–15168.

Specific Cell-Surface Receptor-Mediated Cellular Vitamin A Uptake 50. Kawaguchi, R., Yu, J., Wiita, P., TerStepanian, M., Sun, H. (2008) Mapping the membrane topology and extracellular ligand binding domains of the retinol binding protein receptor. Biochemistry 47, 5387–5395.

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51. Flanagan, J.G., Cheng, H.J. (2000) Alkaline phosphatase fusion proteins for molecular characterization and cloning of receptors and their ligands. Methods Enzymol. 327, 198–210.

INDEX

A

E

ABC transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163–175 ABCA4 . . . . . . . . . . . . . . . . . . . . 130, 163–175, 233, 317, 319 A2E . . . . . . . . . . . 59, 164, 231, 235–237, 240–244, 315–326 Age-related macular degeneration . . . . . . . 56, 59, 164, 209, 236, 309, 315 all-trans-retinal . . . . . . . . 2, 11, 58, 62, 86–87, 96, 115, 130, 164–175, 217, 230–231, 240, 248, 316–320, 322, 324, 330 all-trans-retinal dimer . . . . . . . . . . . . . . . . . 316–320, 322, 324 APCI-MSN . . . . . . . . . . . . . . . . . . . . . . . . . 154, 156, 158–159 A2PE . . . . . . . . . . . . . . . . . . . . . . . . 58–59, 164, 316, 318–325

Electrophysiology. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .95–112 Embryonic stem (ES) cell . . . . . . . . . . . . . . . . . . . . 75–83, 342 Epifluorescence measurements . . . . . . . . . . . . . . . . . . 135, 137

F Feeder independent ES cell culture . . . . . . . . . . . . . . . . 75–83 Fluorescence recovery after photobleaching (FRAP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115–127 Fluorescence resonance energy transfer (FRET) . . . . . . . . . . . . . . . . . . . . . . . 210, 212–219 Fluorescence titration . . . 184–186, 210–211, 213, 221, 224

B

H

Bisretinoid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 315–326 Brain . . . . . . . . . . . . . . . . . . . . 14, 42, 120, 129, 150, 152–153, 278–279, 282–285, 288, 311 5-Bromomethyl fluorescein . . . . . . . . . . . . . . . . . . . . . . . . . 179

High performance liquid chromatography (HPLC) . . . . . . . . . . 4–5, 7–8, 17, 20, 22–28, 33, 47, 62, 67, 71, 151–152, 155–158, 165–167, 169–172, 174, 178, 232–237, 239–243, 248, 263–273, 278, 307, 319–320, 322–325, 333, 336–338, 342–345, 347–348, 351–352, 354, 356–358

C CCD camera . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135, 138, 141 Cellular retinaldehyde binding protein (CRALBP) . . . . . 57, 62–63, 100, 233, 330–332 Cellular retinoic acid binding protein (CRABP) . . . . . . . 78, 151, 178–187 Cellular retinol binding protein (CRBP) . . . . . . . . . . . 57, 76, 116, 231, 331 Chicken . . . . . . . . . . . . . . . . 92, 190–191, 297, 332, 335–338 Chromophore . . . . . . . . . . . . 57–58, 85–87, 92, 95–112, 116, 129–130, 132, 150, 229–231, 233, 235–236, 239, 241, 247–248, 315–326, 330–332, 342 11-cis-retinal . . . . . . . . . . . . 11, 57–60, 62–63, 68, 71, 86–87, 91, 95–104, 106–107, 129–130, 229–231, 233–234, 236–237, 240–241, 247–248, 318, 330 Cogan Plot . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211–213, 224 Cone . . . . . . . . . . . . 32, 56–59, 63, 85–93, 95–112, 130–133, 141–143, 145–146, 164–165, 230–231, 233, 248, 253, 330, 332 CREB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 284 CYP26 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 277–292 Cytochrome P450 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2, 278

I Immunoaffinity chromatography . . . . . . . . . . . . . . . 165–166, 168, 172, 174 Infrared light source . . . . . . . . . . . . . . . . . . . . . . . . . . . 118, 136 Inner filter effect . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211, 225 Interphotoreceptor retinoid-binding protein (IRBP) . . . . 57, 62–63, 66, 68–70, 116, 330–331 Intracellular lipid-binding proteins . . . . . . . . . . . . . . . . . . . 177 Isomerase . . . . . . . . . . . . . . . . . . . . 57, 101, 107, 248, 329–338 Isomerase-2 . . . . . . . . . . . . . . . . . . . . . . . . . . 332–333, 335–338

L LC/MS/MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4–5, 22, 30, 42 Leber congenital amaurosis . . . . . . . . . . . . . . . . . . . . . . . 56, 59 Lecithin retinol acyltransferase (LRAT) . . . . . . . . . . . . . 1–2, 57–58, 115, 231–233, 236, 238–239, 251, 330–331 Lipid droplet . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231, 251 Lipofuscin . . . . . . . . . . . . . . . . . . . . . . . . . . 164, 209, 231, 233, 235–236, 241, 315–326

D

M

Dark adaptation . . . . . . . . . 98, 102, 108, 125–126, 145, 233 Diet . . . . . . . . . . . . . . . . . . . . . 6, 12, 41–42, 46, 263, 295–311 Dietary assessment . . . . . . . . . . . . . . . . . . . . . . . . . . . . 299–300 Diffusion . . . . . . . . . . . . . . . . . . . . . 47, 58, 116–118, 122–126, 204–205, 217, 251, 278

Macular degeneration . . . . . . . . . . . . . . . . . . 56, 59, 163–175, 209, 236, 253, 309, 315 Mass spectrometry . . . . . . . . . . . . . . . 43, 152–154, 178, 234, 236, 243, 318, 324, 342

H. Sun, G.H. Travis (eds.), Retinoids, Methods in Molecular Biology 652, c Springer Science+Business Media, LLC 2010 DOI 10.1007/978-1-60327-325-1, 

363

RETINOIDS

364 Index

Microfluorometric measurement . . . . . . . . . . . . . . . . 129–146 Morphogenetic gradients . . . . . . . . . . . . . . . . . . . . . . . . . . . 278 Mouse . . . . . . . . . . . . . . . . . . . . . . 4, 14, 19, 30, 38–40, 42, 66, 68, 75–83, 97–98, 100–112, 131–133, 137, 141, 143–144, 236, 240–243, 248–249, 251–252, 256–257, 259, 263–273, 279, 282, 284, 289, 291, 302, 317 M¨uller cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130, 329, 331–332

N Nanog regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76, 79–80 N-(4-hydroxyphenyl)retinamide . . . . . . . . . . . . . . . . 195, 213 Non-canonical retinoic acid actions . . . . . . . . . . . . . . . . . . 277 N-retinylidene-phosphatidylethanolamine . . . . . . . . . . . 164, 318–319 Nuclear receptors . . . . . . . . . . . . . . . . . . . . . . 3, 150, 177, 264

O Opsin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85–93, 95–112, 130, 165, 231, 330–331 4-Oxo retinoic acid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2, 11

P Palmitoyl coenzyme A (palm CoA) . . . . . . . . . . . . . 332–333, 336–338 Pattern formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 278 Phospholipid vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . 107, 109 Photoreceptor cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56, 85, 118, 120–121, 126, 132–134, 139–140, 144, 164, 230, 248, 330 Phototransduction . . . . . . . . . . . . . . . . . . . . . . . . . . 58, 98, 101, 103, 110, 229 P450-linked oxidases . . . . . . . . . . . . . . . . . . . . . . . . . . 280–281 Polarized uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55–71

R RALDH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2, 78, 150–151, 278, 281–284 RAR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2–4, 78, 177–178, 279, 284–285 RARE-LacZ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279–289, 291 RA response element (RARE) . . . . . . . . . . 78, 279–289, 291 Recommended dietary allowance (RDA) . . . . . . . . . . . . . 300 Retina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56, 58–59, 67, 69, 85, 96–98, 110–111, 121, 125–126, 131–133, 140, 144–145, 174, 233, 236, 249, 251, 253, 256, 258–259, 279–283, 291, 309, 329–338 Retinal . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1–2, 6–8, 10–12, 14, 16–17, 19–21, 25–26, 29, 33, 40, 42–43, 45, 47–48, 55–71, 86–88, 90–92, 95–104, 106–107, 110, 115–116, 121, 125, 129–130, 140, 145, 163–175, 185, 217, 229–231, 233–237, 240–244, 247–248, 264, 315–326, 329–338, 342 Retinal degenerative diseases . . . . . . . . . . . . . . . . . . . . . . . . 309 Retinaldehyde . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57–59, 62–63, 78, 149–151, 155–159, 180–182, 233, 278, 280–281, 330–331 Retinaldehyde dehydrogenase (RALDH) . . . . . . . . . . . 2, 78, 150–151, 278, 281–284

Retinal pigment epithelial (RPE) . . . . . . . . . . . . . . . . . 56–64, 66–68, 70–71, 96, 98, 101–102, 130, 106–108, 116–117, 130, 164, 204, 230–231, 233, 235, 248–253, 255–259, 315–318, 320–321, 325, 330–338 Retinoic acid (RA) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2–3, 7, 11, 23–24, 58, 76, 78–79, 149–151, 156, 159, 177–187, 217–218, 235, 264, 277–291, 341–342 binding protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78, 151 biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 catabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 277–292 degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186 degrading enzymes . . . . . . . . . . . . . . . . 278, 282, 285, 288 Retinoid . . . . . . . . . . . . . . . . . . . . . . . . 1–49, 55–71, 85–93, 96, 98, 102, 108–111, 115–116, 149–161, 164–175, 177–178, 180, 186, 189, 192–193, 200–202, 210, 217–218, 221, 225, 229–244, 247–260, 263–273, 279–280, 296, 301, 311, 315, 329–338, 342–343, 345, 351–352, 357–358 extraction . . . . . . . . . . . . 67, 71, 172, 237, 265, 268–269, 273, 337, 351, 358 isomerase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57, 329–338 isomerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 239 quantitation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .33, 37 storage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 247–260 trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 248 Retinol . . . . . . . . . . . . . . . . 1–2, 4, 7–8, 10–14, 17–18, 20–21, 25–30, 33, 40, 42–43, 46–49, 56–58, 60, 62, 66, 68–71, 75–83, 86–87, 96, 115–118, 122–126, 129–146, 149–159, 166, 174, 185, 189–206, 209–226, 233, 235, 239–240, 248, 263–273, 296–301, 304–306, 309–311, 338, 342–345, 348–352, 355–358 Retinol binding protein (RBP) . . . . . . . . . . . . . 56–57, 62, 66, 69–70, 76, 116, 189–206, 209–226, 231, 264, 342–343, 357 Retinol binding protein receptor (RBPR) . . . . . . . . . . . . . . 58 Retinosome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231, 250, 253 Retinyl acetate . . . . . . . . . . . . . . . . . . . . . . . . . . . 11, 27–28, 48, 265–271, 273, 358 Retinyl ester . . . . . . . . . . . . . . . . . . . . . 1, 8, 18, 29, 57–58, 76, 101, 115, 150–151, 231–233, 235, 239–240, 248–253, 256–257, 259, 263–273, 298, 306, 331–332, 345, 351–352, 357 Retinyl ester storage structure (retinosome) . . . . . . 249–250, 256–257, 259 Retinyl palmitate . . . . . . . . . . . . . . . . . . . . . . . 2, 11, 27, 47, 62, 153, 265–268, 270–271, 273, 298, 304–305, 310, 331–332, 352, 358 Reverse-phase high performance liquid chromatography . . . . . . . . . . . . . . . . . . . . . 263–273 Rhodopsin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57, 86, 89, 91–93, 99, 102–107, 115–116, 125, 129, 132–133, 145, 165, 229–231, 233, 248, 251–252, 330, 332 Rod . . . . . . . . . . . . . . . . . . . . . . . 56–59, 63, 85–86, 89, 91–92, 95–112, 115–117, 122–126, 130–133, 136, 141, 143, 145, 164–165, 169–171, 174, 230–231, 233, 253, 330 Rod outer segment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91–92, 115, 117, 122–126, 132–133, 145, 165, 169–171, 174, 233, 330 RPE65 . . . . . . . . . . . . . . . . . . . . . 58, 101–102, 106–108, 130, 230–233, 236, 238–239, 242, 248, 330–335, 337–338

RETINOIDS 365 Index S

V

Spectrophotometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167 Stargardt macular degeneration . . . . . . . . . . . . . . . . . 163–175 STRA6 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58, 61, 76, 264, 342–343, 349, 353–354, 356–358

Visual cycle/retinoid cycle . . . . 55–60, 62–64, 96, 164, 248, 330–331 Visual pigment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58, 63, 86, 90, 95–112, 129–133, 143, 248, 257, 330, 342 Vitamin A deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150, 306–307, 310–311 supplementation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79, 300 uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341–358

T Transthyretin (TTR) . . . . . . . . . . . . . . . . . . 56, 66, 69–70, 76, 189–206, 209–226, 343, 351 Two-photon excitation . . . . . . . . . . . . . . . . 116, 249, 255, 258 Two-photon microscopy . . . . . . . . . . . . . . . . . . . . . . . 247–260