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 Her...

Author: Hui Sun | Gabriel H. Travis

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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

For other titles published in this series, go to www.springer.com/series/7651

TM

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

vi

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

vii

viii

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

ix

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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).

4

Kane and Napoli

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)


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 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


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

22

Kane and Napoli

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).


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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Kane and Napoli RA 4,4-dimethyl-RA

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).


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).


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


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.


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.


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


6

7

0 0

5

10

15

20


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.


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.


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.


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.


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)


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


0.0010

atROL

0.0004

0.03

none

0.0010

0.0006

0.04


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


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)


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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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


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).


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


49

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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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

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5. 6.

7.

8.

9.

10.

11.

12.

13.

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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.

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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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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.

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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 (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

Measurement of the Mobility of All-Trans-Retinol

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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 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 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

HPLC / MSN Analysis of Retinoids

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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

30

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

30

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

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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


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.

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15.

16.

17.

18.

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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,

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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 ( 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 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

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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.

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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-

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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.

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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.

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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-

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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).


299

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


305

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 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

Specific Cell-Surface Receptor-Mediated Cellular Vitamin A Uptake

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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.

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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 Müller 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

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