Laboratory Techniques in Biochemistry and Molecular Biology Vol 21: Isolated Hepatocytes Preparation, Properties and Applications

Isolated hepatocytes Preparation, properties and applications LABORATORY TECHNIQUES IN BIOCHEMISTRY AND MOLECULAR BIO...

Author: Michael N. Berry | Anthony M. Edwards | Gregory J. Barritt

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Isolated hepatocytes Preparation, properties and applications

LABORATORY TECHNIQUES IN BIOCHEMISTRY AND MOLECULAR BIOLOGY

Volume 21 Edited by R.H. BURDON - Department of Bioscience and Biotechnology, University of Strathclyde, Glasgow P.H. van KNIPPENBERG - Department of Biochemistry, University of Leiden, Leiden

ELSEVIER AMSTERDAM * NEW YORK * OXFORD

ISOLATED HEPATOCYTES PREPARATION, PROPERTIES AND APPLICATIONS Michael N. Berry Anthony M. Edwards Gregory J. Barritt with contributions by

Marlene B. Grivell Heather J. Halls Bren J. Gannon*

Department of Medical Biochemistry *Department of Anatomy & Histology. School 01Medicine The Flinders University of South Australia G.P.O. Box 2100 Adelaide, South Australia, 5001 A ustralia

and

Daniel S. Friend

Depurmienr of Pathology University of California San Francisco. 94/43. U.S.A .

1991

ELSEVI ER AMSTERDAM. NEW YORK . OXFORD

01991. Elsevier Science Publishers B. V. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical. photocopying, recording or otherwhe, without the prior permission of the puhlhher. E1,sevier Science Publishers B. V., P.O. Box 1527, 1000 EM Amsterdum, The Netherlands.

No responsibility is assumed by the Publisher for any injury an&or damage to persons or property as a matter of products liability. negligenee or otherwise. or from any use or operarion of any methorls. products, instructions or ideas contained in the material herein. Because of the rapid advances in the medical sciences, the puhlisher recommendasthat independent verification of diagnoses and drug dosages should he mudc. Special regulations for readers in the U.S.A.: This publication has been registerc.J with the Copyright Clearance Center, Inc. (CCC), Salem. Massachusetts. lnjormation cun be obtainedfrom the CCC ahout conditions under which the photoropying of purrs qft1ii.s publication may be made in the U S A . All other copyright questions, including photocopying outside the U.S.A. should be referred to the puhlisher.

ISBN 0-444-81299-7 (pocket edition) ISBN 0-444-81302-0 (library edition) ISBN 0-7204-4200-1 (series)

Published by:

ELSEVIER SCIENCE PUBLISHERS B.V. P.O.BOX 21 I lo00 AE AMSTERDAM THE NETHERLANDS

Preface

Since its introduction in 1969 the technique of high-yield preparation of isolated hepatocytes, by collagenase perfusion of the liver, has gained widespread acceptance. The adoption of the preparation in 1973 as the first step in monolayer culture of hepatocytes has further expanded its utility. Literature review suggests that several thousand papers based on use of the technique are published each year. In view of this high degree of utilization, exemplified not only by the numerous publications but also by the holding of several major symposia on the topic, it may reasonably be asked why a manual on isolated liver cell technology is necessary. Certainly there is enough information in the biochemical literature to allow any novice to make a satisfactory attempt at cell preparation. But a survey of the field quickly reveals an extraordinary degree of variability in the approaches employed. Many workers have introduced their own modifications, sometimes without an explanation of why such changes have been made. Moreover, when many workers report that they have prepared cells by the method of Berry and Friend (1969) or by that of Seglen ( 1 976), for example, on reading the Method Section it quickly becomes apparent that quite different conditions have been employed. It might be argued that isolated liver cell suspensions are so resilient that the method of preparation is hardly relevant, provided that a collagenase perfusion technique is used. Certainly their hardiness has very much contributed to the acceptance of the technique. Nevertheless, it is evident from published work that many cell-users are employing procedures that must inevitably result in inferior yields, with a high percentage of damaged cells. This is disadvantageous not only because poor yields are likely to limit the number of studies that can be performed, but

Vi

ISOLATED HEPATOCYTES

also because experience has shown that the quality of the cell preparation is frequently an important determinant of metabolic behaviour. There are a number of other cogent reasons why we consider a book of this kind is timely. A very large body of information about conditions for studying the metabolic properties of isolated liver cells, including their response to hormones, has now accumulated, but much of this is not easy to locate. We have endeavoured to include or cite here what we consider to be the most salient material, with emphasis on methods rather than results. We also review in depth measures of cellular integrity and responses to experimental damage. A substantial amount of work has been carried out with cells from species other than the rat, and we discuss the preparation of isolated liver cells from those species, including the human. Over thepast 20 years many researchers have developed innovative and valuable applications based on the isolated liver cell preparation. These are scattered through the literature, and have often been modified by other workers, again not infrequently without full explanation. We consider that it is appropriate to collect and describe within this Volume some of these developments, particularly those that we judge can be successfully utilized without resort to highly sophisticated and expensive equipment. We have also given special attention, on the basis of the high frequency of publications in the area, to the use of isolated hepatocytes for studies of drug metabolism and toxicity, and for exploring the intricacies of the interactions of calcium ions with the liver cell. Efforts to extend the use of isolated hepatocytes to investigations requiring the preservation of cells for many hours or days began with the availability of cells in high yield. The last decade in particular has seen rapid expansion both in applications of hepatocyte primary culture to a variety of problems and in innovative approaches to maintaining hepatocytes under defined conditions in vitro. The variety of approaches to hepatocyte primary culture, discussed in Chapters 10 and 11, has led not only to preservation of hepatocytes with functional characteristics increasingly like those of hepatocytes in vivo, but also to a better understanding of the factors which influence hepatocyte function in the intact liver.

PREFACE

vii

It was my original intention to write this book as a monograph. Very quickly I became aware of my considerable ignorance in regard to a number of key areas, and I was very grateful to be able to recruit Tony Edwards to write the chapters on cell culture, and Greg Barrittt to provide the chapters on calcium ions and hormone interactions. There are four additional contributors. It is a great pleasure to include a chapter on the morphology of isolated hepatocytes by Dan Friend, my original collaborator (transmission electron microscopy) and Bren Gannon (light and scanning microscopy). Significant contributions, spanning many of the chapters, have been made by Marlene Grivell and Heather Halls, both of whom have also acted with the utmost diligence and patience as sub-editors for the whole Volume. The book is intended to serve as a laboratory manual, and accordingly emphasis is placed on technique. Nevertheless, we believe that a firm understanding of the theory behind the practice is essential if sustained success is to be achieved. We have therefore taken pains to explain the theoretical basis for all approved procedures and to argue why certain approaches cannot be recommended. Collectively, we have had first-hand experience with all of the Protocols and most of the procedures described in the text. Not infrequently, in cases where a method was not familiar to any of us, or we were unsure about the validity of certain conclusions, we have taken steps to rectify this by establishing the procedure in our own laboratory. In instances where our comments are based solely on published information, we endeavour to make this clear in the text. It is our hope that this book will not only provide a useful benchmanual for those entering the area of isolated hepatocyte preparation for the first time, but also will serve as a source of valuable information for the experienced researcher in this field. It is gratifying that the technique of high-yield preparation of isolated liver cells has contributed to the advances in Biochemistry and Cell Biology over the last 20 years. We hope that this book will help make the preparation an even more useful tool. Adelaide, December 1990

MICHAELN. BERRY

This Page Intentionally Left Blank

Acknowledgements

Many people in addition to those listed on the title page have contributed to this book. Foremost amongst them is Ms. Julie-Anne Burton who typed the manuscript and prepared it for electronic processing, wit! extraordinary skill and dedication. Drs. Anthony R. Grivell, John W. Phillips, Roland B. Gregory provided detailed descriptions of some of the procedures described in the text. The excellent line drawings represent the skilled handiwork of Mrs. Susan A. Ruehlemann. A number of colleagues kindly read draft Chapters, including Drs. Peter F. Blackmore, John H. Exton, Robert A. Harris, Sten Orrenius, Chris I. Pogson, and Per 0. Seglen, while Dr. Fred J. Meijer read the whole of the draft manuscript. I am .grateful for their helpful suggestions. Mrs. Marlene Grivell and Mrs. Heather Halls are already listed as contributors for material that they wrote that has been included in the book, but special mention must be made of the time and effort they expended in editing the whole of the manuscript. I also would like to acknowledge the contributions of the many workers in the field, who have been responsible for developing the wide variety of techniques that are described in this book. Particular credit is due to Dr. Roger B. Howard who was the first to demonstrate the feasibility of preparing intact isolated hepatocytes.


Contents

Pwfuce . ..................................................................................

V

AcknowleclRement...................................................................

ix

Abbreviations..................................................................,.......

xxi

Chapter 1. Buckground.. .........................................................

i

The development development of of the the isolated isolated hepatocyte hepatocyte preparation preparation................... .......... The Factors involved involved inin cellular cellular organization organization and and adhesion adhesion........... ..................... Factors 1.2.1. Histology of the liver .............................. I .2.2. The nature of junctional complexes invol ..........,..,........ ... ston .....,,,... .............................................. ..._,_........... 1.2.3. The role of ........................................ 1.3. Mechanical methods for separation of parenchymal liver cells .......... 1.3. 1.3.1. Techniques Techniques................................. .......................................... ................. 1.3.1. ................. Properties ofof isolated isolated hepatocyt hepatocytes prepared by mechanical I .3.2 Properties 1.3.2 .............................. ........ techniques ............................. ................................................. 1.1. 1.1. .2. I I.2.

11

22 22 3

3 44

44

555

1.4.

................................................................ matic procedures procedures..................... Evolution of enzymatic nzymesused usedfor forpreparing preparin suspensi Properties of enzymes ........................,......... .... .... hepatocytes................. I .5. Principles ............ ............................. 1.5. cellsepa separation .......................... Principlesofof cell 1.5. ................................. ......................... I .4. I . I .4.2.

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

666

666 888 999

xii

1.6. I .7.

ISOLATED HEPATOCYTES 1.5.1. Role of Ca2+......................................................................... 1.5.2. Role of collagenase............................................................... 1.5.3. Role of mechanical disruption .............................................. Separation of cells from solid organs other than liver....................... Advantages of the isolated hepatocyte preparation ............................

Chapter 2. High-yield preparation of isolated hepatocytes from rat liver............................................................................................ 2.1. 2.2.

2.3.

2.4.

2.5. 2.6.

2.7. 2.8. 2.9.

Introduction........................................................................................ 2.2.1. Animals .................... 2.2.2. Apparatus...................................... 2.2.3. Reagents .... Two-step procedure............................................................................ 2.3.1. Modifications r step approach 2.3.2. Relative merit Evaluation of common modifications to the recommended method .. 2.4. I . Variation in enzyme content and concentration................... 2.4.2. Heparin as an anti-coagulant.. ........................................ 2.4.3. Inclusion of a chelating agent ............................................... 2.4.4. Inclusion of albumin ............................................................. 2.4.5. Inclusion of metabolites ................... 2.4.6. Exclusion of Mgz+..................................... 2.4.7. Control of oxygenation. pH and the choic 2.4.8. Surgical technique ................................................................. 2.4.9. Mechanics of perfusion ......................................................... 2.4.10. Temperature and oxygenation of washing media .................. Initial determination of cell quality .................................................... 2.5.1. Staining of ce Assessment of cell yiel 2.6.1. Estimation of cell number..................................................... 2.6.2. Estimation of hepatocyte mass and dilution of cells to a desired concentration......................................................................... 2.6.3. Cell yields.............................................................................. Reasons for unsatisfa 2.7.1. Causes of poor quality preparations ..................................... Removal of damaged Storage of hepatocytes 2.9. I . Short-term storage 2.9.2. Long-term storage

10

II 12 12 14

15

15 16 16 17 23 32 32 33 36 36 39 39 40 40 41 41 43 43

44 45 46 48 48 50 52 53 54 55 57 57 58

CONTENTS CONTENTS CONTENTS

xiii xiii xiii

Chapter Chapter 33.. Other Other methods methods for for hepatocyte hepatocyte isolation isolation..................... ..................... .....................

59 59 59

Introduction Introduction Introduction........................................................................................ ........................................................................................ ........................................................................................ Preparation Preparation Preparation of of of isolated isolated isolated hepatocytes hepatocytes hepatocytes from from from liver liver liverslices slices slices......................... ......................... ......................... Preparation Preparation Preparation of of of intact intact intact isolated isolated isolated hepatocytes hepatocytes hepatocytes without without without collagenase collagenase collagenase 3.3.1. 3.3.1. 3.3.1. Apparatus Apparatus Apparatus.............................................................................. .............................................................................. 3.3.2. 3.3.2. 3.3.2. Reagents Reagents Reagents............................................................ ............................................................ 3.4. 3.4. 3.4. Preparation Preparation Preparation of of of isolated isolated isolated hepatocytes hepatocytes hepatocytes from from from species species speciesother other other than than than rat rat rat....... ....... 3.4.1. 3.4.1. 3.4.1. Preparation Preparation Preparation of of of isolated isolated isolated hepatocytes hepatocytes hepatocytes from from from mouse mouse mouse.................. .................. .................. 3.4.2. Preparation 3.4.2. 3.4.2. Preparation Preparation of of of isolated isolated isolated hepatocytes hepatocytes hepatocytes from from from guinea guinea guinea pig pig pig Preparation Preparation of of of isolated isolated isolated hepatocytes hepatocytes hepatocytes from from from ruminants ruminants ruminants ........... ........... 3.4.3. Preparation 3.4.3. 3.4.3. Preparation Preparation of of of isolated isolated isolated hepatocytes hepatocytes hepatocytes from from fromavian avian avian species species species........ ........ ........ 3.4.4. 3.4.4. 3.4.4. Preparation Preparation Preparation of of of isolated isolated isolated hepatocytes hepatocytes hepatocytes from from from fish fish fish.......... .......... .......... 3.4.5. 3.4.5. 3.4.5. Preparation Preparation of Preparation of of human human human isolated isolated isolated hepatocytes hepatocytes hepatocytes.......................... .......................... 3.4.6. 3.4.6. 3.4.6. Preparation Preparation Preparation of of of isolated isolated isolated hepatocytes hepatocytes hepatocytes from from from other other other species species species........ ........ ........ 3.4.7. 3.4.7. 3.4.7. Preparation 3.5. 3.5. 3.5. Preparation Preparation Preparation of of of hepatocytes hepatocytes hepatocytes from from from foetal foetal foetal or or or neonatal neonatal neonatal animals animals animals............. ............. ............. 3.6. Preparation 3.6. 3.6. Preparation Preparation of of of hepatocytes hepatocytes hepatocytes from from from abnormal abnormal abnormal anima anima anima 3.7. Preparation 3.7. 3.7. Preparation of suspensions of damaged hepatocy Preparation of of suspensions suspensions of of damaged damaged hepatocy hepatocy 3.7.1. 3.7.1. 3.7.1. Reagents Reagents Reagents and and and apparatus apparatus apparatus............................... ............................... ............................... Preparation Preparation and and and purification purification purification of of of non-parenchymal non-parenchymal non-parenchymal cells cells cells .................... .................... 3.8. Preparation 3.8. 3.8.

59 59 59 59 59 59 61 61 61 63 63 63 63 63 63 64 64 64 64 64 64 67 67 67 67 67 67 68 68 68 69 69 69 70 70 70 72 72 72

3.1. 3.1. 3.1. 3.2. 3.2. 3.2. 3.3. 3.3. 3.3.

14 14 14

76 76 76 17 17 17 78 78 78 80 80 80

Chapter Chapter 44.. Assessment Assessment ooff integrity integrity of of isolated isolated hepatocytes hepatocytes...... ...... Introduction Introduction Introduction.................................................................. .................................................................. Measures Measures Measures of of of cellular cellular cellular integrity integrity integrity............................................................. ............................................................. ............................................................. 4.2. 4.2. 4.2.III.. Trypan Trypan Trypan blue blue blue staining staining staining ................................. ................................. ................................. 4.2.2. 4.2.2. 4.2.2. Succinate Succinate Succinate oxidation oxidation oxidation.................................. .................................. 4.2.3. 4.2.3. 4.2.3. Equipment Equipment Equipment and and and solutions solutions solutions for for for measuring measuring measuring succina succina succina 4.2.4. 4.2.4. 4.2.4. Enzyme Enzyme Enzymeleakage leakage leakage from from from isolated isolated isolated hepatocytes hepatocytes hepatocytes........ ........ ........ 4.2.5. 4.2.5. 4.2.5. Choice Choice Choice of of of aaa method method method for for for determining determining determining cell cell cell 4.3. 4.3. 4.3. Additional ...................... Additional methods methods for for determining determining cellular cellular integrity integrity ...................... Additional methods for determining cellular integrity 4.3. 4.3. 4.3.III.. Functional Functional Functional measures measures measures of of of cellular cellular cellular integrity integrity integrity.............................. .............................. 4.3.2. Other 4.3.2. 4.3.2. Other Other measures measures measures of of of cellular cellular cellular integrity integrity integrity ...................... ...................... ...................... 4.4. 4.4. 4.4. Assesment Assesment Assesment of of of cellular cellular cellular integrity integrity integrity following following followingcel cel cel

83 83 83 85 85 85 85 85 85 86 86 86

Chapter Chapter 55.. Microscopy Microscopy of of of isolated isolated hepatocyte hepatocyte............................... ............................... ...............................

99 99 99

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

99 99 99 99 99 99 I00 I00 I00

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

I03 I03 I03 I03 I03 I03

4.1. 4.1. 4.1. 4.2. 4.2. 4.2.

5.2. 5.2. 5.2.

5.1.2. 5.1.2. 5.1.2. Features Features Features of of of intact intact intact and and and damaged damaged damaged cel cel cel Morphology Morphology Morphology of of of isolated isolated isolated hepatocytes hepatocytes hepatocytes by by by tran tran tran 5.2.1. 5.2.1. 5.2.1.

lntro lntro lntro

88 88 88

90 90 90 93 93 93 94 94 94 95 95 95 96 96 96 91 91 91


xiv 5.2.2.

5.3.

5.4. 5.5. 5.6.

Features common to both hepdtocytes in situ and isolated

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

103

isolated cell .............................. 5.2.4. Signs of injury................................. Techniques ......................................................................... 5.3. I . Fixation ..................................................... ......... 5.3.2. Current procedure for thin-sectioned material ......................

105

5.3.4. Freeze-fracture. .......................... Morphological studies y Summary of studies by transmission electron microscopy ................ Scanning electron microscopy........................................................ ...............................,................. 5.6.1. Technique

I I4 I I5 I I7 1 I7

Chuprer 6. Biochemicul propertie...................................................... 6.1.

6.2.

6.3.

Basis for expressing cell activity......................................................... ............................................ 6.1.1. Hepatocyte dry weight 6.1.2. Relationship between is II dry weight. cell wet weight and whole liver wet weight ............................................ 6.1.3. Protein and DNA content of hepdtocylcs.............. 6.1.4. Cell number ........................................................................... 6.1.5. Choice of measure for expressing cellular activity ............... Measurement of cellular composition ................. 6.2. I . General considerations.................... 6.2.2. Separation of stable intracellular and by microcentrifugation of hcpatocytcs .................................. 6.2.3. Separation of labile intraccllular and cxtracellular components 6.2.4. Measurement of the intracellular constituents o f centrifuged hepdtocyte ...... 6.2.5. Use of inulin and other agcnts for mcasurcment ofcxtraccllular water ........... .......................................................................... Cellular composition ....... 6.3. I . Water and ion c 6.3.2. Metabolite content ................................................................ 6.3.3. Lipid. protein and enzyme 6.3.4. Adenine nucleotide ............................ ........................ 6.3.5. Glycogen

6.4. 6.4.1. 6.4.2. 6.4.3.

Incubation time and temperature .......................................... Cell mass. incubation volume and oxygenation .................... Incubation media..................................................................

I ox I I2 112 112 113

119

121 121

122 123 125 125 126 126 126 130 131 132

133 140 140

143 144 144 147

148 149 151 I55

CONTENTS

xv

6.4.4. Addition of albumin to incubation media 6.4.5. Addition of gelatin to incubation media ... 6.4.6. Osmolality of incubation media .................................. 6.4.7. Use of inhibitors................................................................... 6.4.8. Solubility of experimental substances .................................... 6.4.9. Availability of substrates to isolated hepdtocytes ................ Measurement of cellular 02-uptake ...................... 6.5. I . Pohrogrdphic measurement of c e h la r 0,- uptake ..... 6.5.2. Manometric measurement of cellular 0,-uptake ................. 6.5.3. AppardtuS..... ...................................................... 6.5.4. Preparation a tion of CO? buffer ......................... 6.5.5. Incubation p .................... 6.5.6. Measurement of rd formation 6.5.7. Measuring 02-uptake with an oxystat system ..................... Metabolic activities............................................................................. 6.6.1. Gluconeogenesis ................................... 6.6.2. Glycolysis .......... 6.6.3. Glycogen synthesis.................. 6.6.4. Ureogenesis ............................................................... 6.6.5. Lipogenesis............................................................................ 6.6.6. Lipid oxidation and ketogenesis........................................... 6.6.7. Protein synthesis............................. 6.6.8. Proteolysis ........................................... Transport activities............................................................................. 6.7.1. Transport across the sinusoidal membrane........................... 6.7.2. Secretion of biliary compone .................... Analysis of radiolabelled compounds .................................

I55 I56 157

Chapter 7. Utilization of hepator-vtes .for drug studies ...............

179

Introduction ......................................... ........ Drug uptake and metabolism by isola ........ 7.2.1. Measurement of drug uptake ................................................ 7.2.2. Study of drug metabolism ..................................................... Isolated hepatocytes and drug toxicity............................................... 7.3.1. Introduction ... ................................. 7.3.2. Indications of cell damage .................. 7.3.3. Investigation of solvent damage ............................... 7.3.4. Investigation of the metabolism of drugs associated with freeradical formation .................................. 7.3.5. Morphological in uced damage ............. 7.3.6 Glutathione depletion .......................................... 7.3.7. Lipid peroxidation ................................... 7.3.8. Covalent binding ........................................................... ............................................... Metabolism and effects of ethanol

179

6.5.

6.6.

6.7.

6.8.

7.1. 7.2.

7.3.

7.4.

15X 158 159 i60

161 162 164 164

I66 I66

167 i68 I6X 169 169 170 171

171 173 174 i76 176 I77 i78

I82 182 184 18a I8a 189 191

I92 194

195 196 198 I99

9

Chapter 8. Study of the effects of hormones .......

20 1

8.1. 8.2.

Hormonal responses exhibited by isolated hepatocytes ...................... Optimal conditions for observing the effects of hormones

8.3. 8.4. 8.5. 8.6.

.. . ........,............................... 8.2.2. Incubation m 8.2.3. Incubation ve .......................... .... Special conditions for ........................ Actions of insulin.. Action of steroid hormones ......................................... Hormone effects on mitochondria isolated from hepatocytes ............

20 I 202 202 205 206 208 209 210 21 1

Chapter 9. Ca2+ ion transport and compartmentation................

215

Measurement of intracellular Ca” ..................................................... 9.1 . I . Determination of total hepatocyte Ca*+............................... 9.1.2. Determination of Ca*+ content of the mitochondria and endoplasmic reticulum... Changes in Ca’+ movement m 9.2.1. Available methods ................................................................. Measurement of intracellular free CaZ+using fluorescent indicators. 9.3. I. Available methods .......................... ........... 9.3.2. Principle of the use of quin2 for measurement of intracellular free Ca*+............................................................................... 9.3.3. Technic rations ................................. ................................................................ 9.3.4. Use of a*+ inflow using glycogen phosphorylase or fluorescent Ca2+ indicators. .................. 9.4. I . Available methods ....

215 216

9.1.

9.2. 9.3.

9.4.

9.5.

....................................... 9.4.3. Major reagen ..,.................. 9.4.4. Comments on ..................... 9.4.5. Measurement ........,................................... 9.4.6. Comments on Assessment of available techniques for the measurement of Ca’ in .................................... ..................................... hepatocytes

218 219 219 224 224 224 221 228 229 229 230 230 233 233 235 235

Chapter 10. Hepatocyte isolation for primary culture and methods for non-adherent culture.......................................................

231

10.1. Introduction ........................................................................................

231

xvii

CONTENTS 10.2. Alternatives to perfusion with collagenase in isolating cells for

......................... 10.3. 10.3.2. Isolation media . 10.3.4. Elimination of non-hepatocytes ............................................ 10.3.5. Elimination of damaged hepatocytes .................................... 10.4. Preserving sterility of cell preparati 10.4.1. Handling of non-sterile ani 10.4.2. Sterilization of equipment and media ................................... 10.5. A procedure for hepatocyte isolation .................................................

.................................................................... .................. 10.6. Suspension culture of hepatocytes 10.6.1. Comparison of suspension and monolayer cultur 10.6.2. Maintenance of oxygenated suspensions of hepatocytes .......................................................... 10.6.3. Media for suspension culture ................................................ 10.6.4. Methods for suspension culture of hepatocytes .................... 10.7. Other non-adherent culture methods .................................................. 10.7.1. Multicellular spheroids ................ 10.7.2. Microencapsulation of hepatocytes ..........................

239 24 I 24 I 242 242 243 245 246 241 247 248 249 252 252 254 256 257 260 260 262

Chapter 11 . Monolayer culture of hepatocytes..............................

265

I I . I . Characteristics of adult primary monolayer cultures: an overview.... I I .1.1. Reviews of the literature ....................................................... I I .1.2. Limitations of simple culture systems................................... 1 I . 1.3. Cell density and cell function in cultur 11.1.4. Contribution of complex media to main function ................................................................................. 1 1 .1.5. Significance of extracellular matrix ....................................... 1 I . 1.6. Heterologous cell interactions in co-culture .......................... 11.1.7. Growth of hepatocytes in culture ................. I 1 . 1.8. Summary ....................................................... 11.2. Attachment of hepatocytes to substrata for monolayer culture 11.2.1. Significance of cell attachment to substrata .......................... 11.2.2. Characteristics of attachment to various substrata .............. 11.2.3. Conditions for hepatocyte attachment . .................. 11.3. Preparation. use and functional effects of speci ata .............. 11.3.1. Plastic ..................................................... 11.3.2. Collagen films or gels............................................................ 1 I .3.3. Matrix glycoproteins: fibronectin laminin ............................ 11.3.4. 'Matrigel' and other extracellular matrix preparations .........

265 265 266 268

.

270 212 274 275 278 278 278 279 280 282 282 282 287 288

xviii


I 1.4. Choice of culture media..................................................................... 11.4.1. Useful media available commercially 11.4.2. Amino acid requirements...................................................... Salts and trace metal ions..................................................... Other components ............. Antibiotics Oxygenati Hormonal functions .........,..................................................,......,.........,.. 1 I .4.9. Hormones and growth factors with growth-stimulatory effects I I .4.10. Effects of proteoglycans or glycosaminoglycans ............ I I .4. I 1. Selective media Non-physiological supp 11.5.1. DMSO ................................................................................... 11.5.2. Phenobarbital ........................................................................ 11.5.3. Nicotinamide 11.5.4. Butyrate ........... ...............,....... .................... Homologous cell inte 11.6.I. Effects of hepatocyte density on preservation of differentiated function ............... ............... 1 I .6.2. Effects of hepdtocyte density on growth-related parameters. Heterologous cell interactions ............................................................. I I .7. I . Rationale and characteristics of co-culture systems ............. 11.7.2. Choice of cell type for co-culture with hepatocytes .............. 11.7.3. Plating and culture procedure for co-culture ..... Selecting appropriate methods for monolayer culture ....................... Variations on monolayer culture ..... 11.9. I . Monolayer culture on microcarriers ...................................... 11.9.2. Culture on filters or memb 11.9.3. Perifused monolayer culture 11.9.4. Culture of periportal or perivenous hepatocytes ................. Culture of hepatocytes from foetal, neonatal and suckling rats 11.10.1. Foetal liver....... ................ 11.10.2. Neonatal hepatocytes ............................................................ 1 I . 10.3. Hepatocytes from suckling rats ............................................. Culture of hepatocytes from other species I1.Il.l. Adult hepatocytes from humans or other primates ............. I I . 11.2. Human foetal hepatocytes ..................................................... I I. 11.3. Hepatocytes from mice. rabbits and other species ............... Some commonly used experimental methods in studies with hepatocyte monolayers .......................... 11.12.I. Assessment of hepatocyte integ 11.12.2. Release of hepatocytes from culture substrata 11.4.4. 11.4.5. 11.4.6. I I .4.7. I 1.4.8.

11.5.

11.6.

11.7.

11.8. 11.9.

11.10.

1 I. 1 1.

11.12.

292 293 30 I 302 303 304 305 305 306 309 313 314 315 315 317 317 318 319 319 320 32 1 32 I 323 324 326 333 333 334 335 336 337 337 340 342 342 342 344 346 347 347 348

xix

CONTENTS 11.12.3. Preparation of whole cell homogenates ................................ I I. 12.4. Preparation of subcellular fractions from hepatocyte monolayers. ................ ..... 11.12.5. Expression of relative bi culture ............................... I I. 12.6. Fixation of cell monolayers ...................................................

.

350 352 353 353

Chapter 12. Selected specialized techniques...................................

355

12.1. 12.1. Introduction Introduction_.... .... 12.2. Sub-cellular fra 12.3. Methods of subcellular fractionation .............................. 12.3.1. Rationale of method of digitonin fractionation ....................

355 355 355 355 357 357 357 357 358 358 359 359 360 360 362 362 362 362

12.3.3. 12.3.3. 12.3.4. 12.3.5. 12.3.5. 12.3.6. 12.3.6. 12.3.7. 12.3.7.

Preparation Preparation of oftubes tubes........ ......... Validation of digitonin fr Variatio Variatio ................................. Alternat Alternat Use Use ofof the the metabolite metabolite indicator indicator method method for for determining determining metabolite metabolite distribution distribution.......................................................... ..........................................................

12.5. 12.5. Preparation Preparation of of cell cell fragments fragments ('cytospheres'). ('cytospheres'). 12.5.1. 12.5.1. Preparation Preparation of of solutions solutions and and cell cellsu suspension ........................ 12.6. 12.6. Preparation Preparation of of plasma membrane fractions ........................................ .......................... 12.7. 12.7. Permeabilized Permedbilized cells 12.7.3. 12.7.3. Other Other chemical chemical methods methods for for hepatocyte hepatocyte permeabilization permeabilization.... .... 12.7.4. 12.7.4. Electro-permeabilization Electro-permeabilization........................................................ ........................................................ 12.8. 12.8. Perifusion Perifusion ofof hepatocyte hepatocytes ... .,..................................................... 12.8.1. 12.8.1. Approaches Approaches to to ... ................ 12.8.2. 12.8.2. Description Description of of 12.8.3. 12.8.3. Use Use of of aa small small 12.9. Measurement of cellula 12.9.1. Measurement of plasma membrane potential. 12.9.2. Measurement of inner mitochondria1 membrane potential ... 12.9.3. .......................................................... 12.9.3. Procedure Procedure... ... .......................................................... ...................... 12.9.4. Calculation of A$ ,,,.......... ...................... ............................................ 12.9.5. Other methods of measuri 12.9.6. Significance of measureme 12.10. .................... 12.10. Measure Measurement of intracellular pH pH................. ..................................... ...................... ......................

I . .

363 363 364 364 365 365 365 365 369 369 312 312 372 372 372 372 373 373 375 375 376 376 378 378 379 379 38 38I I 383 383 384 384 384 384 386 386 386 386 387 387 388 388 389 389 390 390 391 391

xx


12.1 .1 . Transplantation procedures........... 12. I .2. Measurement of hepatocyte functi 12.1 .3. Cryopreservation.......................................................... 12.12. Studies with hepatocyte 'doublets'................. 12.12.1. Electrophysiological studies.................................... 12.12.2. Chemical studies............................... 12.13. Separation of periportal and perivenous hepatocytes ....................... 12.13.1. Separation by collagenase-digitonin perfusion...................... 12.13.2. Separation by centrifugal elutriation ..................................... 12.14. Future developments ................................................................

392 393 395 395 396 396 397 397 399 399

Appendix 1 Composition of media ..................................................

401

Appendix 2 Addresses of suppliers and manufacturers...............

403

References..................................................................................................

409

Subject Index............................................................................................

441

Abbreviations

BSA DMSO EDTA EGTA GSH GSSG HEPES LDH PBS PCA TCA TES

Bovine serum albumin Dimethylsulphoxide Ethylenediaminetetraacetic acid Ethylene glycol-bis(0-aminoethyl ether) N,N , N ’.N ’-tetra-acetic acid Reduced glutathione Oxidized glutathione N-2-hydroxyethylpiperazine-N’-2-ethanesulfonic acid Lactate dehydrogenase Phosphate buffered saline Perchloric acid Trichloroacetic acid 2-(2-hydroxy- I , I -bis( hydroxymethy1)-ethyl1amino)ethane-sulfonic acid


CHAPTER 1

Background

1.1. The development of the isolated hepatocyte preparation The intact isolated hepatocyte suspension is a relatively new experimental preparation, first introduced less than 25 years ago (Howard et al., 1967). Since then its use has increased rapidly, facilitated by the development of a high-yield preparative technique (Berry and Friend, 1969) and by the demonstration of its value by a large number of workers in different fields (for reviews, see Tager et al., 1976; Harris and Cornell, 1983). The successful development of the technique occurred well after the establishment of methods for the preparation of isolated organelles. A major reason for this was the generally held view that preparation of isolated cells from solid organs such as the liver was not feasible. It was known that, in the liver, the hepatocytes are firmly connected to one another, and so the prospect of separating each cell from its neighbours did not seem encouraging. Certainly, until the contribution of Howard et al. (1967), all efforts had yielded highly damaged preparations (Berry and Simpson, 1962). The lack of initial success was due in large measure to an ignorance of the properties of the components responsible for cellular adhesion. In consequence, early attempts at preparing isolated hepatocytes were

2


largely empirical in nature. In time, a firm theoretical basis for cell separation evolved, based on studies involving biochemistry, cell biology, microbiology, histology and electron microscopy. The relevant experimental findings, which led eventually to practical success, are described below.

1.2. Factors involved in cellular organization and adhesion 1.2.1. Histology of the liver

In most textbooks the term ‘parenchymal cell’, used in relation to the liver, refers to the hepatocyte, while the remaining cells are classified as non-parenchymal. The hepatocytes represent 6&65% of the cells of the liver, but because they are larger than the other cells, they occupy about 80% of the volume of the organ (Weibel et al., 1969; Greengard et al., 1972; Blouin et al., 1977; Drochmans et al., 1978). Hepatocytes are arranged in a series of branching, anastomosing, perforated plates, to form a labyrinth between which run the sinusoids (Leeson and Leeson, 1970). The plates are usually one cell thick, although any single cell is bordered by several others. Thus, each cell will be firmly attached to a number of its neighbours by means of the junctional complexes described below. That part of the plasma membrane of each hepatocyte, not involved in forming the junctional complex, abuts on a perisinusoidal space which is separated from the sinusoid by a layer of sinusoidal cells, comprising endothelial, Kupffer, fat-storing (lipocytes) and pit cells. Sinusoidal cells account for about 40% of the total number of hepatic cells, but less than 10% of the liver volume. The remainder of the liver volume is made up of extracellular spaces, both sinusoidal and perisinusoidal. The entire organ is permeated by a fibroconnective tissue skeleton composed of collagenous fibres. Within each hepatic lobule there is a fine meshwork of reticular and collagenous fibres around the sinusoids and within the perisinusoidal spaces. These fibres are

Ch. I .

BACKGROUND

3

embedded in an extracellular ground substance consisting of a complex mixture of proteoglycans, the most common containing heparan sulphate or chondroitin sulphate. 1.2.2. The nature of junctional complexes involved in cellular adhesion Fawcett (1961) records that Bizzozero in 1864 first described junctions between two opposed cells of stratified squamous epithelium, assuming them to be adhesive plates to which both cells contributed. Light microscopists (Schaffer, 1927) identified areas of thickening and adhesiveness at cell boundaries and gave them the name terminal bars. However, it was not until the introduction of the electron microscope that the tine structure of cell junctions was elucidated (Fawcett, 1961). Even then, there was some confusion about both nomenclature and morphology that was clarified by the in depth studies of Farquhar and Palade (1963). These workers defined the junctional complex as composed of the tight junction (zonula occludens) where the cell membranes of two contiguous cells appear united but are not fused (Gumbiner, 1987), the zonula adherens or intermediate junction, which is characterized by a true intercellular space measuring approximately 20 nm across, and the desmosome (macula adherens). The desmosome was identified by the presence of laminar densities in the intercellular space and by high local concentrations of dense amorphous and fibrillar material in the subjacent cytoplasmic matrix. In 1967, Revel and Karnovsky described the gap (communicating) junction, which was subsequently included in the definition of the junctional complex (Friend and Gilula, 1972). 1.2.3. The role of Ca2+ Chambers (1940) draws attention to early studies by Ringer (1890), with tadpoles and a green alga (Laminaria),that demonstrated the importance of Ca2+for cell adhesion. Ringer showed that the action of distilled water, in loosening the cement substance that binds cells

4


together, was prevented by bicarbonate of lime. A role for Ca2+in cellular cohesiveness was subsequently confirmed by studies at the turn of the century by Herbst (1900) and Lillie (1906). These early observations prompted workers in the field of organ culture to omit Ca2+ from the incubation medium when cell separation was desired. It is now known that the function of cell adhesion molecules is dependent on their interaction with Ca2+(Geiger et al., 1987).

1.3. Mechanical methods for separation of parenchymal liver cells 1.3.1. Techniques

Initial attempts to prepare isolated hepatocyte suspensions were more concerned with cell yield than with cell integrity. One of the earliest relevant reports is a publication by Schneider and Potter (1943) in which they describe forcing liver tissue through cheesecloth and then through bolting silk. The isolated cells were separated from tissue debris by repeated low-speed centrifugation and resuspension of the pellet in hypotonic phosphate buffer. Bucher et al. (1951) obtained isolated cells by placing rat liver slices in an aqueous medium and shaking them with glass beads on a mechanical shaker. As much as lo'%) of the liver was converted to isolated (though not intact) cells. They observed that low temperatures and the inclusion of Ca2+in the suspending medium reduced cell yields. Anderson (1953) described the first method for preparation of (damaged) isolated cells from rat liver in high yield. A liver from an adult rat was distended by retrograde perfusion via the inferior vena cava and hepatic veins. The perfusing fluid was an approximately isotonic solution, free of Ca2+and containing a Ca2+binding agent such as pyrophosphate, ethylenediaminetetraacetic acid (EDTA), or citrate. The distended liver was cut into small pieces, transferred to a homogenizer, and gently dispersed with a loosely fitting pestle in

Ch. I .

BACKGROUND

5

about ten volumes of Locke's solution, without Ca2+.Shreds of connective tissue were removed by straining the suspension through bolting silk. The hepatic cells were centrifuged and resuspended several times in order to separate them from tissue debris and erythrocytes. All procedures were carried out at M 0 C .It was found that cell yields varied considerably even when identical preparative conditions were employed, but usually over 40% of the liver was converted into isolated cells. Anderson's method was modified by Branster and Morton ( 1957). who found that the use of a chelating agent was unnecessary provided that perfusion of the liver was performed via the portal vein. They observed that the ease of isolation of cells increased with step-wise rises in the temperature of the perfusion and dispersion solutions up to 50°C. Normally perfusion and dispersion were carried out at 37°C. Under these conditions 0.25-0.4 M sucrose or mannitol at pH 7.4 proved highly satisfactory for the preparation of mouse liver cells (Berry, 1962). Other mechanical separation techniques employed steel screens (Kaltenbach, 1954) or repeated passage of liver tissue through a narrow orifice (Longmuir and Ap Rees, 1956), but none of these techniques gave yields as high as Anderson's procedure (Laws and Stickland, 1956).

1.3.2. Properties of isolated hepatocyres prepared by mechanicul techniques When the metabolic properties of hepatocytes prepared by mechanical techniques were examined, it became immediately apparent that the cells would oxidize only one physiological substrate, succinate (Laws and Stickland, 1956; Kalant and Young, 1957; Lata and Reinertson, 1957; Leeson and Kalant, 1961). Endogenous respiration was absent, as was glycolysis. It was also observed that the cells lost most of their cytoplasmic enzymes during the preparative procedure, but retained glucose-6-phosphatase and glutamic dehydrogenase (Henley et al., 1959; Zimmerman et al., 1960). A morphological basis for this

6


behaviour was provided by the studies of Berry and Simpson (1962) who demonstrated that mechanically prepared cells underwent marked changes in fine structure. The plasma membranes of the cells were extensively disrupted and the mitochondria and endoplasmic reticulum became vesiculated. Berry (1962) found that these cells were capable of endogenous respiration and could oxidize Krebs cycle intermediates, but only when incubated in a medium that was suitable for sustaining the respiration of isolated mitochondrial suspensions. The inability of many workers to obtain respiratory activity, except with succinate as substrate, was due to their use of media containing Ca2+,which penetrated the cells and rapidly led to mitochondrial swelling and damage (Berry, 1962).

1.4. Development of enzymatic methods for isolated hepatocyte preparation 1.4.1. Evolution of enzymatic procedures

The use of trypsin to loosen cells from tissue has a long history dating back at least as far as Rous and Jones (1916). For many years trypsin remained exclusively the enzyme of choice for workers in the field of tissue culture (Moscona, 1952), until Mandl et al. (1953) isolated and characterized collagenase from Clostridium histolyticum. Lasfargues (1957) utilized this enzyme for cell separation during the cultivation of normal mammary epithelium, and in 1961 collagenase was prepared on a commercial scale by Worthington Biochemical Corporation. Subsequently, Rodbell (1964) used the enzyme for the isolation of intact fat cells from adipose tissue. Three years later, Howard et al. (1967) reported the first successful isolation of intact hepatocytes from rat liver. These workers injected a balanced-saline solution (Hanks’ medium), free of added Ca2+and containing 0.05% collagenase plus 0.1% hyaluronidase, into the major sinusoids of a rat liver. The injection was made under sufficient pressure to blanche the organ, which

Ch. 1.

BACKGROUND

7

was then cut into thin slices that were incubated in enzyme solution in vessels agitated in a shaking water bath. The incubation vessel contents were filtered after which the freed cells were separated by centrifugation and washed several times in enzyme-free medium. of the original liver were Quantities of intact cells representing 3-5% obtained. Although yields of intact cells obtained by the technique of Howard et al. (1967) were low, this work provided unequivocal evidence that the preparation of suspensions of intact parenchymal cells was possible, and stimulated attempts to find an enzymatic procedure that gave high yields of cells. It seemed desirable to try to obtain better exposure of the connective tissue of the liver to enzyme and better survival of the cells during digestion. This was achieved by perfusing the liver with an oxygenated, balanced-saline medium, lacking added Ca2+but containing collagenase and hyaluronidase (Berry and Friend, 1969). During perfusion the liver swelled considerably and its collagen matrix was digested and loosened. The liver was removed from the animal and the external capsule ruptured with a spatula. The released cells were shaken in large conical flasks in a water bath for 15 min at 37°C before the parenchymal cells were purified by centrifugation at 50 x g for 2 min followed by resuspension in enzyme-free medium. Yields of intact cells represented 30-50'%, of the original liver. Since the description of the collagenase perfusion technique, numerous minor changes have been introduced. However, perhaps the only important modification has been the now widely employed twostage procedure, originally introduced by Seglen (1972,1976), in which perfusion of the liver with a medium free of Ca2+and enzyme is followed by a perfusion with a Ca2+-enrichedcollagenase-containing medium. The various approaches to hepatocyte preparation are discussed in detail in Chapters 2 and 3. Three other enzymes should be mentioned in discussing procedures for preparing isolated liver cells. Lysozyme has been used for digestion of foetal liver ( H o m e s et al., 1971), and pronase for the preparation of non-parenchymal cells, since this enzyme selectively destroys

8


hepatocytes while leaving Kupffer cells intact (Berg and Boman, 1973). Dispase is a neutral protease sometimes used in combination with collagenase. 1.4.2. Properties of enzymes used for preparing suspensions of isolated hepatocytes

A variety of enzymes have been tested to determine their usefulness for the preparation of isolated undamaged hepatocytes. These include: (1) Collagenase EC 3.4.24.3 (previously EC 3.4.4.19) from CI. histolyticum, more accurately termed clostridiopeptidase A. A collagenase is defined as an enzyme capable of causing hydrolytic cleavage of molecules of collagen in their native conformation and at their helical region. These enzymes, which are very specific for collagen, are proteases with a specificity for the X-Gly bond in the sequence Pro-X-Gly-Pro, where X is usually a neutral amino acid. They require Ca2+ for activity and usually contain Zn2+. Collagenases can be isolated from many bacterial sources (e.g. Achromobacter iophagus, Mycobacterium tuberculosis, Pseudomonas aeruginosa and other microorganisms) and also can be obtained from animal tissue that is undergoing rapid reorganization such as tadpole, human and rat skin and bones (see Seifter and Harper, 1971). However, CI. histofyticum provides the main commercial source of collagenase. Crude commercial preparations of collagenase from this source can be separated into six different isoenzymes (Bond and Van Wart, 1984). These crude preparations also contain trypsin, clostripain and caseinase activities, and differences in the relative quantities of these enzymes may be responsible for some of the observed lot-to-lot variation in efficacy when these products are used for hepatocyte preparation. (2) Hyaluronidase EC 3.2.1.35, now called hyaluronoglucosaminidase, is an endoglycosidase with a specificity for endo-Nacetylhexosaminic bonds. It hydrolyses 1,4 linkages between 2-acetamido-2-deoxy-~-~-glucose and D-glucuronate residues in hyaluronic acid, a glycosaminoglycan found in the ground substance

Ch. 1.

BACKGROUND

9

of virtually all connective tissue, cell coats, the vitreous of the eye and the synovial fluid of joints. The commercial supply of this enzyme is from bovine and sheep testis, leeches and Streptomyces hyalurolyticus. Although originally recommended for dissociation of liver tissue (Howard et al., 1967), the enzyme is now considered unnecessary for this purpose. (3) Trypsin EC 3.4.21.4 (formerly EC 3.4.4.4). This enzyme is a pancreatic serine protease capable of hydrolysing peptides, preferentially at bonds involving the carboxyl group of the basic amino acids, L-arginine or L-lysine. The enzyme also shows some esterase and amidase activity. Purified trypsin is not suitable for use by itself in the dissociation of adult liver tissue, as it is ineffective against native collagen. However, it occurs as a contaminant in collagenase preparations from C1. histolyticum and may have a synergistic action under these circumstances, since it can hydrolyse the portions of collagen which are not tightly wound in the helical form. (4) Clostripain EC 3.4.22.8 (formerly EC 3.4.4.20). CI.histolyticum is the source of this enzyme which is highly specific for the carboxyl peptide bond of arginine. It is activated by Ca2+and is found in most commercial collagenase preparations. ( 5 ) Lysozyme EC 3.2.1.17. This enzyme is an endoglycosidase which catalyses the hydrolysis of 1,4-@-linkagesbetween N-acetylmuramic acid and N-acetylglucosamine residues in glycosaminoglycans. The main commercial sources are hen or turkey egg white and human milk. (6) Pronase EC 3.4.24.4. Pronase is a proprietary name for a nonspecific protease isolated from Streptomyces griseus. It hydrolyses practically all naturally occurring peptide bonds. (7) Dispase EC 3.4.24.4 is a protease that hydrolyses bonds involving leucine or phenylalanine.

1.5. Principles of cell separation It is apparent that the successful methods for hepatocyte isolation,

10


described above, involve treatment of the liver tissue in three different ways - exposure to a medium very low in Ca2+,digestion with collagenase and gentle mechanical disruption. 1.5.1. Role of Ca2+

The most critical step in the preparation of isolated hepatocytes is treatment of the liver with a medium very low in Ca2+,or containing a Ca2+-bindingagent. It is now well-established that the desmosome is the initial and critical site affected by perfusion with such a medium. Berry and Friend (1 969) recognized that during the perfusion procedure the regions of cell membranes that contain desmosomes invaginate (Section 5.2.3). The invaginations subsequently pinch off, forming endosomes that include hemi-desmosomes on their cytoplasmic surfaces. During invagination and vacuole formation, desmosomal fibres separate from their plaques. These changes, which are irreversible, closely resemble events occurring in embryonic tissue incubated in a Ca2+-freemedium containing trypsin (Overton, 1968). Ca2+ lack, rather than proteolytic enzymes, is responsible for desmosome cleavage. This is apparent from studies where cell separation has been achieved without the use of proteolytic enzymes, by incubating epidermal (Hennings and Holbrook, 1983) or hepatic tissue (Berry et al., 1983; Wang et al., 1985) in Ca2+-freemedium. Splitting of the desmosomes is the key element in liver cell separation procedures, since no success is obtained if Ca2+are present throughout the incubation period. Conversely, it is clear that Ca2+are essential for desmosomal integrity. Mattey et al. (1987), subjected various epithelial cell cultures to Ca2+concentrations lower than 0.1 mM and observed that the desmosomes cleaved. It can be inferred, therefore, that during the first stage of the twostage procedure introduced by Seglen (1972, 1976), pre-perfusion with a medium lacking Ca2+brings about cleavage of the desmosomes to a stage of irreversibility. In consequence, subsequent perfusion of the liver with a medium rich in Ca2+is without effect on cell adhesion.

Ch. 1.

BACKGROUND

11

However, desmosomal separation is not instantaneous, and a too rapid restoration of Ca2+,before irreversible changes have taken place, can lead to re-adhesion of the hemi-desmosomes and a consequent severely reduced yield of isolated cells, many of which are damaged (see Section 2.3.2). The role of Ca2+in cell separation is seen to be complex. These ions are known to be essential for collagenase activity, so that the inclusion of any Ca2+-bindingagent in the medium will prevent collagenase action. Yet removal of Ca2+ is essential for desmosome cleavage. It is fortuitous that sufficient Ca2+are present in the perfusate, either derived from the crude collagenase preparation (approx. 0.5-1 pmol Ca2+/50 mg collagenase) or released from the hepatocytes or interstitial fluid, to ensure adequate collagenase activity yet allow desmosomal cleavage. Collagenase binds Ca2+rather tightly (Gallop et al., 1957), and it seems possible that the enzyme could actually scavenge trace amounts of Ca2+from the perfusion medium. We have observed that in the absence of any pre-perfusion procedure, the addition of concentrations of CaZ+above 20 pM to the medium at the commencement of perfusion impairs desmosomal cleavage. Such treatment produces poor yields of cells with multiple blebs, low ATP content and low rates of gluconeogenesis from lactate. However, trypan blue staining of the cells may remain quite low (< 10%) at initial concentrations of Ca2+in the perfusate as high as 50 pM. Wittenberg et al. (1986) also observed that the addition of 20 pM CaZ+to the perfusion medium increases the proportion of damaged cells in myocyte preparations. 1.5.2. Role of collagenase

It might be thought that the role of collagenase in cell separation is self-evident. However, there is perhaps more uncertainty about this element than about any other feature of the procedure. While there is no doubt that crude collagenase is effective in assisting disruption of the liver, it seems equally clear that the purified enzyme is much

12


less efficacious. The question of whether collagenase acts synergistically with other proteases, and other properties of the enzyme, are discussed in Section 2.4.1. As indicated in Section 1.5.I , collagenase is not an essential component for the preparation of substantial quantities of intact isolated hepatocytes, although the collagenase-free procedure yields a large percentage of damaged cells. A collagenase-free method is described in Section 3.3 and the procedure necessary to remove damaged cells in Protocol 2.5. 1.5.3. Role of mechanical disruption

A number of the early methods for preparing isolated hepatocytes relied on mechanical disruption, carried out by perfusion of the liver under pressure with a medium low in free Ca2+,or containing a Ca2+-bindingagent. Such treatments invariably gave rise to damaged cells. Parenchymal cells appear very sensitive to shearing stresses, which readily lead to plasma membrane rupture. Nevertheless, gentle mechanical treatment is an essential feature of all effective procedures. From the morphological studies of Berry and Friend (1969) it is clear that neither short-term digestion of the liver with collagenase, nor removal of Ca2+,fully cleaves tight or gap junctions. Instead, the separation of two opposed cells is achieved by an actual tearing of the plasma membrane of one of the cells after desmosome breakage. In consequence, the gap and tight junctions are retained by one cell and the resultant defect in the plasma membrane of the other is probably repaired (see Section 5.2.3). Final yields of intact hepatocytes, prepared by collagenase perfusion procedures, are so large that it must be inferred that cells with plasma membranes torn during loss of portions of the junctional complex usually survive.

1.6. Separation of cells from solid organs other thun liver It is gratifying to find that the mechanisms for separating hepatocytes have proved relatively straightforward, despite the complex nature of

Ch. 1.

BACKGROUND

13

the processes involved in liver cell adhesion. A particularly fortuitous element of isolated hepatocyte preparation is that the cells are tolerant to Ca2+-depletion, at least for short periods (approx. 40 min). Myocardial cells are much more susceptible to Ca” removal (Farmer et al., 1983), and in consequence the high-yield preparation of intact isolated myocytes (except from neonatal tissue) has proven very difficult. However variations of the one- and two-step methods described in Chapter 2 have been reported (e.g. Montini et al., 1981: Wittenberg and Robinson, 1981), in which myocytes have been isolated that are tolerant to incubation with physiological concentrations of Ca2+.Recently, Haworth et al. (1989) reported that addition of 1 mM Ca2+to a Ca2+-freeKrebs-Henseleit medium, 15 min after collagenase had been added, yielded a preparation in which 77.5% of cells were Ca2+-tolerant, rod-shaped myocytes. Preparation of isolated cells from some solid organs such as the kidney is more difficult and in some cases may not be feasible, apparently because junctional complexes fail to separate. Methods using collagenase and hyaluronidase to digest the kidney by perfusion (Balaban et al., 1980) or incubation of minced tissue (Stokes et al., 1987) produce preparations of isolated renal tubules and collecting ducts, rather than individual cells. Stokes and co-workers were unable to separate isolated collecting ducts into single cells by using Caz+-freemedia or a number of proteolytic enzymes, presumably as a result of the failure of desmosomes to separate. However, Eveloff et al. (1980) were able to produce a population of single cells from the rabbit kidney medulla, but only by using repeated tryptic digestion of the collagenase-treated preparation. Collagenase has also been successfully used for the isolation of exocrine pancreatic cells (Amsterdam and Jamieson, 1974), pancreatic islets (Lacy and Kostianovsky, 1967)and adrenocortical cells (Kitabchi and Sharma, 1971). An extensive bibliography of enzymatic methods for the isolation of cells from many sources including liver, mammary gland, muscle, lung and brain tissue is provided in a valuable publication by Worthington Biochemical Corporation ( 1990).

14


1.7. Advantages of the isolated hepatocyte preparation For many years the surviving slice was the mainstay of liver cell research, particularly when it was considered that a relatively intact preparation was required. After the introduction of the isolated perfused liver preparation (Trowell, 1942; Miller et al., 1951) it rapidly became evident that the properties and biochemical activities of the surviving slice fell far short of those demonstrable with the perfused organ (Miller et al., 1951; Hems et al., 1966). Unfortunately, maintenance of the latter in good physiological condition is technically rather difficult, and severe limitations are placed on the experimental scope by the need to supply all substrates or other agents through a single perfusing medium. In essence, this tends to limit experimental studies to a single set of conditions for each perfusion. The isolated hepatocyte suspension is easier to prepare than surviving slices and shows a far superior range of activities. In most instances its metabolic behaviour closely emulates that of the perfused liver. The special attraction of the isolated cell preparation is that a suspension of, say, 50 ml can be obtained from one liver, divided up into aliquots, and used in a single study to explore the effects of up to 50 different experimental combinations. Once separated, the cells are relatively resistant to mechanical damage, so that they can be shaken in incubation vessels for several hours at a time. The ease of preparing aliquots of the suspension greatly facilitates the establishment of dose-response curves when examining the properties of drugs or hormones. It is therefore hardly surprising that the isolated hepatocyte preparation has become a widely used experimental technique.

CHAPTER 2

High-yield preparation of isolated hepatocytes from rat liver

2.1. Introduction A turning point in the development of methods for the preparation of intact isolated hepatocytes came with the demonstration by Howard et al. (1967) of the value of collagenase as a tool for liver cell separation. Two years later Berry and Friend (1969) introduced the technique of collagenase perfusion of the liver as a means of greatly increasing hepatocyte yield. Since then numerous modifications of the collagenase perfusion method have been described, but in almost all instances little or no explanation has been given for the changes made, and hence they are not easy to evaluate. The modifications include alteration to the composition of the perfusion medium or the duration of the perfusion, removal of the liver from the animal prior to perfusion, or changes to cell washing procedures. As discussed in Chapter 1, any method for the preparation of isolated hepatocytes that utilizes collagenase perfusion will have two conflicting requirements. The cells must be exposed to a very low concentration of Ca2+to allow the cleavage of hepatic desmosomes. On

16


the other hand, the activity of collagenase requires the presence of Ca2+.This conflict has been resolved in two ways, generally termed the ‘one-step’ and ‘two-step’ procedures. The one-step procedure, which essentially follows the method of Berry and Friend (1969), as modified by Berry (1974a), takes advantage of the fortuitous circumstance that the concentration of Ca2+required for adequate collagenase activity is substantially less than that required for desmosomal integrity (see Section 1.5.1). The contrasting approach of the two-step procedure is to allow the hepatic desmosomes to cleave irreversibly during a pre-perfusion of the liver with Caz+-freemedium for at least 10 min, and then to add collagenase and Ca2+in physiological or supraphysiological concentration to the perfusate (Seglen, 1972, 1976). In this Chapter both the one-step and the two-step procedures will be described in detail and their relative merits discussed. Each of these approaches has proved satisfactory in the hands of numerous workers in the field, and both are recommended. In fact, the two procedures as practised today differ very little. Not only are similar perfusion media employed, but an early re-exposure of hepatocytes to Ca2+is now also a recommended part of the one-step procedure, in order to provide optimal conditions for maintenance of cellular integrity. The usefulness of many of the modifications suggested by other workers are also evaluated in this Chapter.

2.2. One-step method For the sake of clarity the requirements for the one-step method are reported below. The minor modifications necessary for the two-step procedure are set out in Section 2.3. 2.2.1. Animals

The description given here applies to the preparation of hepatocytes from adult rats in either the fed or fasted state. In general, rats of

Ch. 2.

HIGH-YIELD PREPARATION

17

150-300 g body weight are the most convenient from a surgical point of view; the liver weights of such animals range between 5 and 10 g. For reliable results it is desirable to maintain consistency in regard to the strain, sex and weight of the rat. Of equal importance is maintenance of a regular feeding schedule, and a consistent light and dark cycle. Variations required to prepare isolated liver cells from other species and from young animals are described in Chapter 3. 2.2.2. Apparatus

Perfusion of the liver for 10-30 min with collagenase-containing perfusate is obligatory if high yields of intact hepatocytes are required. Thus, because of the expense of collagenase, a recirculating perfusion is highly desirable. However, there is no need to use an elaborate perfusion system of the type described by Miller et al. ( 1 95 1) or Mortimore (1961). The components of a simple perfusion system, together with possible sources, are listed in Table 2.1. Of course, similar and equally satisfactory components are available from other manufacturers. The chief elements are a 250-ml reservoir, a peristaltic pump and a means of warming the perfusate to 37°C before it enters the liver. Additional requirements are a bubble trap, a magnetizable stainless steel or plastic and steel platform, approximately 15 x 25 cm, portal vein and outflow cannulas and latex and tygon connecting tubing. An oxygenating system, whereby the water-saturated gas mixture is vigorously bubbled through the perfusate in the reservoir, is also recommended, or alternatively a multibulb glass oxygenator (Krebs et al., 1974) (Fig. 2.l), or the silastic tubing ‘lung’ (Fig. 2 . 2 ) devised by Hamilton et al. (1974)(but see Section 2.4.7).A coarse filter should be inserted in line to remove debris such as hair. Thermistors for sensing the temperature of the liver or perfusate are more convenient than a thermometer. Temperature maintenance can be achieved by placing all fittings and tubing in a thermostatted plexiglass cabinet, or by running the perfusate through a water-jacketed coil, immediately prior to its passage

TABLE 2.1 Components of a perfusion system and their sources Component

Source

Peristaltic pump and controller

Cole-Palmer Instrument Co., 7425N Oak Park Avenue, Chicago, ILL., 60648, U.S.A. Model No. 7546-14

Jacketed coil

I

x v Dimensions

r

& 4

6 E

Internal diameter - 2 mm External diameter - 4 mm 24 loops Coil diameter - 30 mm Coil length - 100 mm

Silastic tubing

Dow Corning Corporation, Medical Products, Midland, MI 48640, U.S.A. Cat. No. 601-405

Internal diameter 0.228 mm External diameter 0.303 mm

Tygon tubing

Cole-Palmer Instrument Co. 7425N Oak Park Avenue, Chicago, ILL., 60648, U.S.A. Formulation R3606; Cat. No. 6408-41

Internal diameter - 1.66 mm External diameter 4.94 mm

28

:i-2:

Natural latex tubing

Kent Latex Products Inc., Kent, Ohio, 44240, U.S.A. Cat. No. 402R

Internal diameter Wall - 0.79 mm

Inflow (portal vein) cannula

Teflon Jelco, Critikon Johnson & Johnson Medical k.k. 15-5 Ichiban-Cho, Chiyoda-ku, Tokyo, Japan Cat. No. 4054

18 G, 44 mm

Outflow cannula

Teflon Jelco, Critikon Johnson & Johnson Medical k.k 15-5 Ichiban-Cho, Chiyoda-ku, Tokyo, Japan Cat. No. 4058

14 G , 57 mm

Water circulator

Haake DI, Haake Mess-Technik, Dieselstrasse 14, D-7500Karlsruhe 41, Germany Model No. 000-3350

- 3.18

mm

0

P rJ

20


Fig. 2.1. Multibulb glass oxygenator.

through the liver. Although groups already established in the field of liver perfusion have tended to favour use of a cabinet, many new workers engaged in isolating hepatocytes use a water-jacketed coil (Seglen, 1976) or oxygenator (Krebs et al., 1974) (Fig. 2.1). If a coil

Ch. 2.


21

Fig. 2.2. Silastic tubing 'lung'.

is used, a thermostatted water circulator will also be required. The maintenance of a liver surface temperature close to 37"C, as measured by means of a thermistor, is desirable. A simple version of the bubble trap can be constructed from the inverted barrel of a 10 ml plastic syringe, fitted with a rubber stopper that has been pierced with two short lengths of 1-2 mm (internal diameter) stainless steel, glass or plastic tubing, to allow inflow and outflow of fluid, and an on-off tap, to enable the barrel to be filled with perfusion medium (Fig. 2.3). The drip chamber of a disposable saline perfusion kit can also be utilized. The various elements are assembled to form a perfusion circuit as

22


Fig. 2.3. Bubble trap.

illustrated in Fig. 2.4. Fine natural latex tubing is used to interconnect each item, but tygon tubing is used for the segment that passes through the peristaltic pump. The length of tubing running from the reservoir to the peristaltic pump includes an in-line, on-off tap and the coarse filter. The pump should be capable of delivering 60 ml/min and is connected to the jacketed coil (or oxygenator) which in turn is joined to the bubble trap. A further length of tubing passes from the bubble trap to the portal vein cannula. The perfusate exits the liver via the hepatic veins, which empty into the inferior vena cava. A length of tubing, connected to the outflow cannula, drains fluid from the vena cava back into the reservoir. Efflux from the liver is assisted by a siphoning effect, achieved by mounting the rat on a platform some 15 cm above the reservoir (Fig. 2.4). The apparatus should be kept clean and free of bacterial growth. After each perfusion it is rinsed thoroughly with deionized water, and

Ch. 2.


H p L BT RES GI GO 0 T F 4

heaters Pump lung bubbletrap resetvoir gasinlet gasoutlet on-off tap tray filter direction of flow

23

1. 2. 3. 4. 5.

6. 7.

30cm 40cm 35cm 2x2m 44cm 60cm 60cm

Fig. 2.4. Perfusion cabinet and circuit.

the tubing is flushed weekly with 70% ethanol. The tubing generally requires changing at twice-yearly intervals, but should be renewed more frequently if there are signs of contamination. 2.2.3. Reagents The perfusion medium routinely used in our laboratory contains 160 mM Na+, 5.4 mM K+, 0.8 mM Mg2+,139 mM C1-, 0.8 mM SO,'-, 1 m M phosphate and 25 mM HC0,-.The medium is initially

24


equilibrated to a pH of 7.4 at 37°C in a water bath, by bubbling carbogen, a mixture of 0, and CO, (95:5) through it. A useful check of equilibration is provided by including 0.001% phenol red and noting whether or not the indicator shows an appropriate orange-pink colour. If retention of the glycogen content of cells from fed rats is desired, the perfusion fluid should also contain at least 15 mM glucose. For consistent results it is desirable to use only the highest quality reagents and preferably to reserve containers of the necessary salts solely for perfusion purposes. The purity of the water used to make solutions is also critical. For all reagents we use deionized rainwater, passed through a Permutit Hi-Pure laboratory water system. Water is not collected from the water-purifier until the conductivity is below 0.07 pS/cm at 22°C. If a commercial water-purifying system is not available, deionized double-distilled mains water ( c 0.1 pS/cm) is generally suitable for cell preparation.

Protocol 2.1 Preparation of working solutions (i) Prepare a ten-fold concentrated stock balanced salt solution comprising 80 g NaCl, 4 g KCl and 2 g MgS04.7H,0 dissolved in 1 1 H,O. Freeze this stock solution in 200-ml quantities in plastic bottles. Also make up a 100-fold concentrated stock phosphate buffer, comprising 2.4 g Na,HPO, and 0.4 g KH,P04, dissolved in 200 ml H,O, and store frozen in 20-ml plastic bottles. (ii) To prepare bicarbonate-free phosphate saline, add each item listed, with mixing, to a 2-1 container in the following order: 1.5 1 H,O; 200 ml concentrated stock balanced salt solution; 20 ml concentrated stock phosphate buffer; 10 ml 0.2% phenol red solution.

Make up this mixture to 2 1 with H,O and store at 0-4"C. Do not keep the solution for more than a few days, If the number of cell preparations will be small, make up lesser quantities.

Ch. 2.


25

(iii) To prepare perfusion medium, add 7.5 ml of 1 M NaHCO, (stored at 0 4 O C ) to 293 ml of phosphate saline. Warm the mixture to 37"C, while gassing it with carbogen to bring the pH to 7.4. The perfusion medium, which should be prepared fresh daily, has a bicarbonate concentration similar to that of plasma (25 mM). (iv) Prepare cell washing medium daily by mixing 3 ml of 80 mM CaCI, with 237 ml of phosphate saline. Adjust the pH to 7.4 and store at 0 4 ° C . This medium does not contain NaHCO,. (v) Dissolve collagenase (50 mg) in 10 ml of perfusion medium as soon as it is certain that liver perfusion has been successfully established. When subsequently mixed with the circulating perfusion medium, the final concentration of collagenase in the perfusate will lie between 0.03 and 0.04% (w/v). Sources and quality of collagenase are discussed in detail in Section 2.4.1.

Protocol 2.2 Preparation of hepatocyte suspensions (i) Prime the apparatus with perfusion medium, and re-circulate the solution at about 40 ml/min. Approximately 200 ml of medium will be required in systems that use bubbling or a silastic tubing lung to oxygenate the perfusion buffer to an 0,-tension > 500 mmHg. (ii) Anaesthetize the rat with intraperitoneal pentobarbital (60 mg sodium pentobarbitone/kg rat body weight). Place the unconscious animal, abdomen upwards, on the platform, securing its legs to the platform with adhesive tape (Fig. 2.5). Moisten the abdomen with ethanol, grasp the abdominal wall with a pair of forceps and open the abdominal cavity from the tip of the sternum to the pubis, along the midline. Commencing at the mid-point of the vertical incision, make a horizontal cut on each flank from the centre line around towards the back. With cuts of appropriate depth it is possible to construct a well out of the abdominal wall. This cavity will be used to retain perfusate oozing from the liver (Fig. 2.6). Reflect the intestine to your right-hand side, thereby exposing the liver. Gently push the two main lobes of the liver proximally (towards the animal's head)

26


Fig. 2.5. Diagramatic representation of liver perfusion.

with a wet swab and uncover the portal vein (the major vessel supplying the liver) and, lying deeper, the inferior vena cava. With the aid of a small pair of curved forceps draw a nylon thread under the vena cava, above the branch to the right kidney, and tie a loose half-knot round the vein, ensuring that the vessel is in no way obstructed. A small bulldog clip may be attached to the ends of the ligature to facilitate its later identification. Now repeat this procedure for the por-

Ch.2.


27

Fig. 2.6. ‘Well’ for collection of leaked perfusate.

tal vein, although in this case it is not necessary to clip the ends of the tie. To make subsequent cannulation easier, do not place the portal vein ligature closer than 1 cm from the point of entry of the vein into the liver. The positions of cannulas and ligatures are illustrated in Figs. 2.5 and 2.7. (iii) Cannulation of the portal vein is most easily achieved by use of a trocar and cannula (Fig. 2.8). To cannulate the portal vein, fix the vessel by grasping a small portion of its uppermost wall with very fine forceps, and puncture it with the trocar about 1.5 cm from the point where the vein branches to enter the liver lobes. The trocar should be held almost parallel to the portal vein during the insertion, to avoid piercing the opposite wall. Manoeuvre the tip of the cannula to lie about 0.5 cm proximal to the branch-point (Fig. 2.7). Immediately withdraw the trocar and pull tight the ligature round the portal vein. Blood should flow back from the liver, thereby completely filling the cannula. This is important because an air bubble introduced into the liver can seriously impair the perfusion. At this point sever the inferior vena cava, with a pair of scissors, below the level of the kidney. (iv) Now slow the pump to deliver approximately 10 ml/min, and connect the inflow tubing to the portal cannula, taking great care to avoid air bubbles. The portal cannula line is held in a clamp, with a heavy base containing a permanent magnet, to prevent movement


28

ligature

7\

\\

cannula

Fig. 2.7. Placement of portal cannula.

and accidental withdrawal of the cannula, as well as to maintain correct position (Fig. 2.5). Perfusate will flow through the liver, blanching it, and then seep through the proximal cut end of the vena cava. Place a large cotton swab in position to absorb the fluid. (v) Open the thorax with a pair of heavy scissors, by snipping through the diaphragm and cutting around to the dorsum on both sides. Then pick up the sternum and rib-cage and pull them forward with a pair of forceps. Cut away the whole of the front of the chest and remove it, after freeing the sternum from ligaments attached to the mediastinum. Place a ligature around the inferior vena cava below the heart. Locate the right atrium and grasp it with a small pair of blunt forceps, Make a small hole in the heart with a pair of fine scissors. Gently push the outflow cannula vertically down through this hole into the inferior vena cava, so that its tip comes to lie just above where

Ch. 2.


29

Fig. 2.8. Trocar and cannula.

the hepatic veins empty into the vessel (Fig. 2.5), and tie in place. There is no need to use a trocar for this insertion. If the placement is satisfactory, perfusate will flow freely from the outflow cannula and will almost cease to flow from the cut section of the vena cava. At this point locate the clip attached to the ends of the loose ligature around the vena cava and pull the tie tight. (vi) Connect the vena cava cannula to the outflow tubing, which is held in a clamp to help maintain cannula position. The outflow tubing passing downwards from the platform towards the reservoir acts as a siphon that reduces the risk of hepatic swelling. Increase the flow rate gradually over a period of about 30 s to 3-4 ml/g tissue per min (i.e. 2-0 ml), depending on liver size. Allow the first 50 ml of perfusate to flow to waste. Re-position the outflow tubing so that the perfusate flows back into the reservoir. Since an additional loss of 10-20 ml medium occurs in the time (1-2 min) taken to place the outflow cannula, about 130 ml of perfusion medium are now recirculating in the system. At this stage the liver should appear normal in size and yellowish in colour, due to the washout of blood. A mottled appearance with residual red patches indicates that the perfusion

30


has not been satisfactory due, for example, to air-bubbles or clots. Under no circumstances should the liver be allowed to swell at this stage. Swelling indicates that the outflow cannula is not properly sited in relation to the hepatic veins. (If the liver begins to swell, stop the perfusion by turning off the pump, and attempt to make the necessary adjustments.) Once it is certain that the perfusion is technically satisfactory, pour the collagenase solution into the reservoir. At this stage swab the abdominal cavity clean of blood clots and perfusate. (vii) After about 10 min, fluid commences to ooze freely from the surface of the liver and accumulates in the thoracic and abdominal cavities. The liver itself takes on a somewhat turgid look but is not grossly distended, although the edges of the lobes adopt a rounded appearance. If blood clots have been removed, the fluid accumulating in the abdominal cavity will not be contaminated. Collect this fluid continually with a syringe and return it to the reservoir for the duration of the perfusion. Within 15-30 min, the consistency of the liver will have become so soft that it will rupture on pressure if not handled gently. We obtain higher yields of cells, with no apparent decrease in cell quality, if perfusion continues until fissures form spontaneously in the liver without any applied pressure or manipulation. The liver will also be moderately swollen and have a speckled appearance, with yellow dots representing islands of cells in a sea of pink perfusion medium. Because of variation in the quality of collagenase, it is not feasible to prescribe an exact perfusion time and some experience is required in making a judgement as to when perfusion should be terminated. If gentle pressure with a finger tip on the underneath of a liver lobe results in fracture of the liver surface, digestioncan be taken as complete. (viii) At the end of the digestion period, remove the softened and disintegrating liver. With care it can be virtually pulled away from its ligamentous attachments to the mesentery, though some use of scissors will be necessary, particularly to dissect the liver from its connections to the stomach and duodenum. Transfer the extirpated organ to a beaker and disrupt it gently, using pointed scissors to tear open the liver capsule, to cut connective tissue strands, and if necessary to

Ch. 2.


31

tease the tissue apart. An alternative method recommended by Seglen (1976) is to comb the liver with a stainless steel dog comb (3 mm between teeth). If the digestion has been satisfactory the tissue will now have the consistency of a very concentrated homogenate. (ix) Retrieve about 50 ml of medium from the perfusion reservoir, add 1 ml of 80 mM CaC1, and disperse the liver tissue in this solution. (The reason for the addition of Ca2+at this time is discussed in Section 2.3.2). Transfer the suspension in approximately equal portions to two 250-ml conical wide-necked flasks, which are shaken (80 cycles/min) at 37°C for 10 min in a reciprocating water bath after being gassed with an atmosphere of carbogen, delivered through a gassing manifold. This incubation serves to break up cell clumps and to digest isolated nuclei and damaged cells. Cells of borderline quality appear to be removed by this treatment, since experience has shown that if this incubation step is omitted, the final yield is greater but the suspension contains a higher percentage of overtly damaged cells. (x) Rinse the contents of the two flasks through nylon mesh (250-pm pore-size) into a clean 250-ml beaker with about 100 ml of washing medium at U ' C . Distribute the dilute suspension evenly among four ice-cold 50-ml centrifuge tubes and centrifuge at 40 x g for 2 min at 0 4 ° C in a refrigerated centrifuge. Aspirate the supernatants carefully and repeat the washing and centrifuging procedures twice. At each step carefully resuspend each cell pellet in about 5 ml of washing medium by using a wide-mouthed glass tube, e.g. the wide end of a pasteur pipette, and then gently mix the suspension with an additional 25 ml of washing medium. After the penultimate wash, combine the cell pellets by flushing them, with small portions of washing medium, through 100-pm nylon mesh into a single tared tube. Make the suspension volume up to 30 ml with washing medium and centrifuge at 50 x g for 4 min. Remove the supernatant as completely as possible without disturbing the cell pellet and weigh the pellet. Finally suspend the cells in the required volume of washing medium at U ' C (Protocol 2.4). Washing removes constituents that have leaked from intact or damaged cells, cell debris and a substantial proportion of the damaged cells, as well as almost all non-parenchymal

32


cells. Higher centrifugal forces result in slightly larger yields of hepatocytes but more contamination with damaged hepatocytes and non-parenchymal cells. Liver perfusion requires a certain amount of practice. Not infrequently the beginner encounters problems with placement of cannulae, so that swelling of the organ occurs, or alternatively, the portal vein is not successfully cannulated, or the cannula slips out of the vein during the perfusion. Nevertheless, the technique should be within the capability of virtually all research workers, and generally we find that novice research assistants can learn to make high quality cell preparations within a few days of commencing training.

2.3. Two-step procedure 2.3.1. Modifications required to the one-step procedure for the two-step approach The essential feature of the two-step procedure for isolated hepatocyte preparation is that the liver is first flushed with about 500 ml of a Ca2+-freemedium, a process that takes approximately 10 min. Then the perfusion medium is changed to one containing Ca2+and collagenase. The pre-perfusion medium can be identical to the Caz+-freeperfusion medium used in the one-step procedure, although some workers use a medium buffered with N-2-hydroxyethylpiperazineN’-2-ethanesulfonic acid (HEPES) (Seglen, 1976). The effluent is not returned to the reservoir but allowed to flow to waste. When the preperfusion medium is almost depleted, a recirculating perfusion is established by returning the effluent to the reservoir, to which is added 100 ml of perfusion medium containing both collagenase and Ca2+.All other aspects of the procedure are identical with the one-step method.

Ch. 2.


33

The two-step approach was introduced by Seglen (1976) as a consequence of his studies of the effects of collagenase on the isolated perfused liver (Seglen, 1972). It was observed that the liver swelled more rapidly on addition of collagenase and 5 mM Ca2+,than when collagenase alone was added, provided that there had been a preliminary perfusion with medium very low in Ca2+.Seglen argued that since collagenase activity is dependent on Ca2+,and since a virtual absence of Ca2+is a prerequisite for cell separation, it was good practice to first perfuse the liver with a medium free of Ca2+and subsequently perfuse with a Ca2+-rich medium containing collagenase. Seglen (1976) recommended a Ca2+concentration of 4.8 mM in the final perfusion medium to enhance collagenase activity. However scrutiny of his data (Seglen, 1972) suggests that, after a 10-min preperfusion with Ca2+-freemedium, 1 mM is an equally effective concentration. Since the rat liver is normally exposed to blood containing about 1.25 mM free Caz+,it would seem good practice not to raise the concentration in the perfusion medium much above this level. Accordingly, the majority of workers who have adopted the two-step procedure have substantially reduced the Ca2+concentration in the perfusion medium, and our own studies confirm that 1 mM Ca2+is sufficient for a maximal rate of digestion. 2.3.2. Relative merits of the one-step and two-step procedures

Recent surveys of the literature suggest that the two-step procedure is currently more popular, though no comparisons of the relative merits of the one- and two-step methods have been published. Because of the higher Ca2+concentration in the collagenase-containing perfusion medium in the two step procedure, the overall perfusion time is about 5 min less than in the one-step approach. Nevertheless, the two procedures, performed correctly, give virtually identical yields and proportions of intact cells. This is explicable on the basis that the slightly longer perfusion associated with the one-step procedure compensates

34


for the fact that the activity of the collagenase in the perfusion medium is less than that observed under the conditions employed by Seglen (1976). In view of the similarity of the two approaches in terms of performance, it may be asked why the two-step procedure has proved more popular. Soon after the original descriptions were published, findings from a number of laboratories indicated that isolated hepatocytes, incubated in the absence of added Ca2+,are unable to achieve and maintain normal intracellular concentrations of Na+ and K+, and that the cells can be irreversibly damaged by long-term deprivation of Ca2+(Barnabei et al., 1974; Baur et al., 1975; Kolb and Adam, 1976; Edmondson and Bang, 1981; Thomas and Reed, 1988a,b). These potential problems were not encountered by workers using the twostep procedure in which Ca2+are replaced early. Information about Ca2+-lossfrom the cells, or the deleterious effects of Ca2+-deprivation,was not available when the original onestep perfusion method for hepatocyte preparation was devised. We have found that in the one-step procedure the perfusate leaving the liver after the first 50 ml flush has a Ca2+concentration of 15-35 pM. At the end of the perfusion period the medium contains Ca2+at a concentration of 30-75 pM. These Ca2+are not merely a contaminant of the added collagenase, but rather represent loss from the liver cells or interstitial fluid. In order to counteract this loss, the one-step procedure has now been modified by replacement of Ca2+at the incubation step immediately following perfusion of the liver (Protocol 2.2), and this has proved as satisfactory as replacement of Ca2+during perfusion, in preventing damage due to Ca2+-deprivation. If unwashed cells are assayed immediately following the perfusion procedure, hepatocytes prepared by the two-step method have Na+ and K+ contents closer to normal than those isolated by the original one-step method. However, the 10-min incubation carried out in a Ca2+-containingmedium immediately post-perfusion, as part of the modified one-step method, corrects abnormal ion gradients (Section 6.3.1). Hence, as shown in Chapter 6, differences in integrity and ion content between cells prepared by the one-step or two-step methods are not detectable.

Ch. 2.


35

When compared to whole liver, cells isolated by collagenase perfusion techniques have been shown to have depleted levels of carnitine (Christiansen and Bremer, 1976) and amino acids, including glutamate and aspartate, which are important components of the malate/aspartate shuttle (Krebs et al., 1974, 1976; Cornell et al., 1974). Reduced levels of glutathione have also been demonstrated by several groups, the actual decrease being dependent on the length of perfusion (Hogberg and Kristoferson, 1977) or the presence or absence of ethylene glycol-bis(P-aminoethyl ether) tetra-acetic acid (EGTA) in the pre-perfusion medium of the two-step procedure (Vida et al., 1978). An important function of the liver is to maintain the steady-state level in the blood of many important metabolites such as amino-acids (Schimassek and Gerok, 1965). It follows that the liver might be more severely depleted of some or all of these substances by the large volume non-recirculating initial perfusion of the two-step procedure than by the continuously recirculating perfusion of the one-step method. However, there have been no controlled studies to examine this question. It should be noted that in the two-step procedure, a large volume initial perfusion is not absolutely essential. At the time of Seglen's early investigations (Seglen, 1972), it was not widely appreciated that irreversible desmosomal cleavage, as a consequence of exposure of the liver to a perfusion medium low in Ca2+,is a time-dependent function. In other words, the flushing of the liver with large volumes of solution, as recommended by Seglen (1976), is effective in cleaving the desmosomes, not only because extra Ca2+are eluted from the junctional complexes by the large quantity of medium (approx. 500 ml) used for the initial non-recirculating liver perfusion, but also because the wash-out procedure takes at least 10 min to accomplish. In accord with this, adequate isolated cell preparations have been obtained after pre-perfusion of livers with Caz+-freemedium with flows of only 150 ml over 12 min (Vida et al., 1978) or 200 ml over 10 min (van de Werve, 1980). Nevertheless, our own studies using the twostep method, indicate that Caz+-depletionof the desmosomes, and the greatest yield of healthy cells, is best achieved by perfusion with a much larger Caz+-freevolume as recommended by Seglen (1976). Thus, if

36


it is considered desirable to keep the pre-perfusion volume low, the most reliable way to do so is to employ the one-step procedure where only 50 ml of perfusate are flushed through the liver before recirculation of the medium is established. In summary, the theoretical advantages of adding Ca2+back to the medium during perfusion are not realized in practice in terms of better yields. CaZ+replacement during the perfusion (two-step procedure) or immediately post-perfusion (one-step procedure) are equally effective in enabling hepatocytes to correct abnormalities in ion distribution that occur during hepatocyte isolation. To ensure good cell yields, Ca2+replacement during two-step perfusion should not be effected until at least 10 min has elapsed from the time of commencement of the initial perfusion. If these precautions are followed, both the one-step and two-step procedures are satisfactory in producing good yields of intact hepatocytes. Each has been used successfully for a multiplicity of experimental studies over the past 15-20 years, and it does not seem reasonable to attempt to select either as superior in practice to the other. For the newcomer to the field the best advice would seem to be to try each technique and determine which appears to give the best results for the question under study, bearing in mind that both of the original techniques have been modified to the point that, as carried out today, there is actually minimal difference between them. Indeed, we have performed extensive comparisons of the two procedures and can detect no significant differences in cell yield or performance.

2.4. Evaluation of common modifications to the recommended method 2.4.1. Variation in enzyme content and concentrution

Early methods for the preparation of isolated hepatocytes (Howard et al., 1967; Berry and Friend, 1969; Krebs et al., 1974) employed O.l'%l

Ch. 2.

HIGH-Y IELD PREPARATION

37

hyaluronidase, as well as collagenase, as digestive enzymes. Although there is some suggestion that hyaluronidase improves cell yields, its effect is marginal, and hence it is best omitted, to avoid any possible deleterious effects of this enzyme on the cell surface. The key enzyme in hepatocyte isolation is collagenase, but there is surprisingly little definitive information about what constitutes a good enzyme preparation in terms of its efficacy in cell separation. The original perfusion procedure recommended a final collagenase concentration of 0.050/0, and the majority of workers have used a similar concentration. There are, however, a number of reports in the literature recommending enzyme concentrations of half this amount, or even less (Johnson et al., 1972; Wagle and Ingebretsen, 1975).In our hands, use of collagenase levels lower than 0.03'%1 yields considerably higher proportions of damaged cells. Seglen (1976) reports that 0.05% is the optimal collagenase concentration for the two-step procedure. The greatest uncertainty lies in relation to the quality of the collagenase. At this time Worthington, Boehringer Mannheim and Sigma are the major suppliers of the enzyme, and each company offers various grades of collagenase, containing different amounts of contaminating enzymes. On the basis of previous experience and practical tests, the companies endeavour to select batches of collagenase considered to be particularly suitable for hepatocyte preparation, but the assessments of company scientists do not always coincide with those of the researcher in the field. Unfortunately, after 20 years it is still not feasible to predict from assay of the activities of a crude collagenase sample and the contaminating proteases. whether the batch will be good for hepatocyte isolation or will allow a particular cellular property to be demonstrated. However, there appears to be a general consensus that if the behaviour of receptors is under study. it is preferable to avoid collagenase preparations rich in trypsin. Some groups have included soybean trypsin inhibitor (Sigma) in the perfusion medium to counteract potential trypsin action (Crane and Miller, 1977). but there is no evidence that this is beneficial and some indication that it may be harmful (Dickson and Pogson, 1977).

38


It is surprising that, more than 20 years after isolated hepatocytes were first prepared using collagenase, there is still uncertainty concerning what components of the crude mixture are essential for satisfactory cell separation. We have tested several collagenases from CI. histolyticum that were classified as being ‘crude’ preparations and contained clostripain, tryptic and caseinase activities in varying ratios. There appeared to be no relationship between the quality or yield of hepatocytes and the proportions of the different enzymic activities. Some products that had been recommended specifically for the preparation of cells other than hepatocytes (e.g. adipocytes) very rapidly digested the liver and produced high yields of good quality cells. In the one-step perfusion procedure, it is critical that the Ca2+content of the enzyme preparation be low. For example, we were unable to obtain satisfactory hepatocytes when we used collagenase isolated from A . iophugus both with and without dispase (a neutral protease). The Ca2+ concentration in our A . iophugus collagenase/dispase preparation was found to be sufficient to raise the perfusing medium Ca2+ concentration a further 50 pM. When we used this collagenaseldispase preparation in the two-step procedure, in which the Ca2+content of the collagenase is of no consequence, a moderate yield of cells was achieved. Our results, however, were not as good as those of Queral et al. (1984) who obtained their largest yields and lowest degree of cell damage using a collagenase/dispase preparation from A. iophugus. Seglen (1976) has obtained success with commercial purified preparations of collagenase, substantially free of protease, but possibly containing clostripain and tryptic activities. Others have not achieved similar results with single fractions purified from crude collagenase (Hatton et al., 1983). It may be necessary to use specific collagenases to preserve some hepatocyte enzymic activities. Quibel et al. (1986) and Marteau et al. (1988) found it necessary to use chromatographically purified collagenase to isolate hepatocytes with heparin-releasable neutral triester lipase EC 3.1.1.3 activity preserved at the cell surface. Whereas most commercial collagenases inactivated this lipolytic activity, it was

Ch. 2.


39

found associated with hepatocytes when either Sigma Type VII or Worthington type CISPA collagenase from CI. histolyticum, each of which contain only small amounts of contaminating proteases, were used. However, cell yields were only 3 x lo7 intact cells compared with the expected 25-50 x 107 cells for livers of comparable size digested with crude collagenase. Experience indicates that the only reasonably sure way to secure an effective batch of collagenase is to obtain three or four small samples from the supplier of choice and test them. Fortunately all major suppliers recognize the problem of choosing an appropriate enzyme batch, and will provide free samples for the researcher to test. When a suitable sample is identified, a larger quantity of that batch number can be ordered. In view of the high cost of collagenase, this approach is highly recommended. Companies such as Worthington, that have been preparing collagenase for many years, have built up a large body of knowledge concerning the product and can assist the researcher in batch selection. Even so, practical trial remains the only sure way to determine if the collagenase selected is suitable for the research worker’s needs. 2.4.2. Heparin as an anti-coagulant

It has been a common practice in many laboratories to use heparin as an intravenous anti-coagulant prior to commencing perfusion of the liver. This does not appear to be necessary as, in our experience, blood clears completely from the liver without its addition. Also, since heparin is known to effect some surface proteins (e.g. neutral triester lipase, Marteau et al., 1988) and can penetrate into hepatocytes and intracellular compartments (Khatsernova et al., 1989) it would seem preferable to avoid it during hepatocyte preparation. 2.4.3. Inclusion of a chelating agent

In order to achieve complete removal of Ca2+some workers have in-

40


cluded 0.1-1 mM EDTA or EGTA in the pre-perfusion medium used in the two-step procedure. The presence of a chelating agent is not necessary to obtain good yields of cells. However, Vifia et al. (1978) have reported that 0.1 mm EGTA (but not EDTA) prevents the loss of cellular reduced glutathione (GSH). It seems likely that the loss of GSH is related to the presence in the perfusate of heavy metals (Fe, Cu, Mn) that catalyse the oxidation of GSH by 0,. These metals are unlikely to be present if good quality water is used to make up reagents. 2.4.4. Inclusion of albumin

A number of researchers have included albumin in the perfusion medium (Johnson et al., 1972), in the hope of binding potentially noxious agents in the perfusate. While it is likely that crystalline albumin will have no ill-effects, some preparations of albumin that are economically feasible to use contain amounts of Ca2+sufficient to interfere seriously with cell separation. There is no evidence that greater yields or better quality cells are achieved in the presence of albumin; hence there seems little justification for its use. On the other hand, the inclusion of defatted albumin in the incubation medium is frequently desirable during metabolic studies (Section 6.3.3). 2.4.5. Inclusion of metabolites

Glycogen is rapidly depleted in hepatocytes from fed animals, unless perfusing and washing media contain at least 15 mM glucose. As mentioned previously (Section 2.3.2), other metabolites are also lost from the cells, and because of this a number of workers choose to add substrates such as pyruvate, Krebs cycle intermediates or amino-acids to the perfusing or washing media (Zahlten et al., 1973). Since some of these metabolites are rather expensive, perhaps a more economic approach is to add them, if desired, to the cell suspension at the time of incubation (Section 6.4).

Ch. 2.


41

2.4.6. Exclusion of Mg’+

Seglen ( 1976) found that 5 mM Mg2+inhibited collagenase-induced swelling of the liver and recommended the exclusion of Mg2+from perfusion media. He further pointed out (Seglen, 1979) that Mg2+ supports the attachment of hepatocytes to tissue culture plates (Seglen and Fossa, 1978) and suggested the possibility of Mg2+-dependent junctions between cells. However, we have found no difference in cell yield or the time required for liver digestion when 0.8 mM Mgz+is included in these media and we suggest on general principles that physiological concentrations of this ion should be included in perfusion solutions. 2.4.7. Control of oxygenation, p H and the choice of buffers

The early successes in the preparation of isolated hepatocytes (Howard et al., 1967; Berry and Friend, 1969) were achieved using Hanks’ medium (Appendix l), a salt solution low in buffering capacity and containing only about 4 mM bicarbonate ion (Hanks and Wallace, 1949). Experience showed that when livers were perfused with this medium, the pH tended to drop as low as 7.1, even when the perfusing medium was continually equilibrated with 100%)0,. Nevertheless, satisfactory yields of intact cells were consistently obtained. Subsequently Cornell et al. (1973) substituted the bicarbonate-saline of Krebs and Henseleit (1932) (Appendix 2), with Ca2+omitted, and employed carbogen as the gas phase. This perfusion medium was found to give yields similar to, but no better than, those obtained with Hanks’ medium. From first principles, it would seem desirable to maintain physiological conditions as closely as possible during the perfusion procedure, even though on technical grounds there seems no advantage in replacing Hanks’ medium with Krebs-Henseleit bicarbonate-saline. The latter solution provides a higher capacity buffering system that more closely resembles plasma and, like

42


plasma, requires a gas phase containing 5% CO,. For these reasons the use of Krebs-Henseleit bicarbonate-saline is recommended. Seglen (1976) has taken a different approach, arguing that the use of a concentrated synthetic buffer (e.g. 100 mM HEPES) is more convenient than the use of bicarbonate, which, though a better buffer, requires a gas phase of CO,, to allow the correct pH to be maintained. Moreover, Seglen (1973, 1976) has simplified the perfusion on the basis that, in his view, continuous oxygenation of the perfusing medium is not required. Under these circumstances, the employment of HEPES is obviously advantageous. Even so, the great majority of workers would consider oxygenation of the medium during the perfusion step to be desirable, since under hypoxic conditions the breakdown of ATP to AMP in the rat liver is frequently accompanied by degradation of AMP, primarily by deamination to IMP followed by dephosphorylation to inosine (van den Berghe et al., 1989). We have conducted a number of studies in which cells from fasted animals were prepared according to Seglen’s recommendations. On most occasions there was no evidence of cellular deterioration induced by lack of oxygenation of the perfusion medium. Moreover, in most instances the hepatocytes, on incubation, showed no fall in levels of ATP, and, as previously reported (Le Cam et al., 1976), exhibited undiminished rates of gluconeogenesis.We also observed that a medium containing only 10 mM HEPES appeared satisfactory for cell preparation, despite suggestions that a concentration of 100 mM was necessary for obtaining good yields of intact cells. It would appear, therefore, that hepatocytes in situ are rather more resilient than has been believed. Notwithstanding this, on occasions rather poor preparations, with a higher percentage of damaged cells with blebbing plasma membranes (Section 5.1.2) were obtained in the absence of an augmented 02-supply, leading to the conclusion that, at least for the relatively unskilled worker, a technique that employs adequate oxygenation of the liver is desirable. Krebs et al. (1974) points out that the bicarbonate-CO, system is not merely a buffer, since CO, takes part in many metabolic reac-

Ch. 2.


43

tions. Depriving the liver of CO, will prevent those reactions from occurring. Of course the critical issue is whether or not the metabolic changes, induced by various manipulations during the preparative procedures, are irreversible. It appears that the great majority of them are not, so that most of the insults to which the liver tissue is exposed during cell preparation do not lead to permanent damage. Despite this, it seems reasonable to argue that if a ‘physiologically balanced’ saline will bring about the desired results, that should be the medium of choice. Some authors have included erythrocytes in the perfusion medium to facilitate oxygenation of the liver, but there is no evidence that this is necessary. Ultimately, the test must be whether or not the phenomenon that the researcher wishes to study is optimally demonstrable under the chosen preparation and experimental conditions. 2.4.8. Surgical technique

A number of workers choose to remove the liver from the carcass of the animal during the perfusion period. This procedure is recommended by Seglen (1976) but, as he points out, the surgical removal of the liver is not easy without training. On the other hand removal of the liver facilitates the containment of perfusate oozing from its surface, and is aesthetically more satisfactory. The minor advantages of hepatectomy do not seem to compensate for the extra difficulties involved, and the procedure is not recommended. Some authors have suggested that removal of the liver is desirable if the cells will subsequently be cultured, on the basis that bacterial contamination is more easily avoided. However, numerous laboratories successfully prepare uncontaminated cell suspensions using perfusion in situ (see Section 10.4.1). 2.4.9. Mechanics of perfusion

As discussed earlier, the length of the perfusion is of some importance. Satisfactory yields of intact isolated cells cannot be expected if insuf-

44


ficient time is allowed for the desmosomes to cleave. In the two-step procedure an adequate total perfusion time is about 20 min, whereas about 5 min longer is generally required for the one-step procedure, but as previously mentioned in Protocol 2.2 (vii), these times are very dependent on collagenase quality. Some authors have claimed success with perfusion procedures lasting half this time, in some cases not even attempting to establish a re-circulating perfusion. This approach is not to be recommended, since it invariably leads to lower yields and increased proportions of damaged cells. The perfusion should be continued until the liver consistency is so soft that it fractures with gentle pressure. Another practice to be deplored is that of deliberately allowing the liver to swell, by partially obstructing the outflow, or increasing the perfusion pressure, which is normally about 12 cm of water. Again, from experience gained over many years, it can be predicted that a rise in pressure sufficient to swell the liver will generally increase the proportion of damaged cells. 2.4.10. Temperature and oxygenation of washing media An issue which has caused some debate is whether the cells should be cooled during the washing procedures, or handled at room temperature. It is certainly true that cells cooled below 20°C lose K+ and gain Na+, and that such ion movements are accompanied by a substantial degree of cellular swelling. Nevertheless, it appears that these changes are reversible, so that cells cooled during the washing procedure regain their normal cation balance during subsequent incubation (Section 6.3.1). We have observed that cellular concentrations of ATP are invariably lower in cells washed with media at room temperature, even if the solutions have been previously gassed with 0, or, where appropriate, carbogen. Moreover, both the synthetic ability and ultimate K+ content of such cells are less than those of cells washed in cooled medium, while enzyme leakage is greater, suggesting a degree of cellular damage (see Section 4.2.4). Our recommendation is to wash cells in ice-cold

Ch. 2.


45

media (i.e. held at OAOC), which do not need to be gassed. However, if for experimental purposes it is desirable to use uncooled media for washing purposes, it is important to pre-oxygenate them. Note that oxygenation should never be carried out by bubbling gas through the cell suspension, since this is a sure way to damage the cells. The oxygenating gas should be flowed over the cells as they are swirled mechanically or by hand in the flask. Our normal practice is to stand tubes containing cell suspensions on ice whenever we are not sampling from them, and to use a refrigerated centrifuge for cell-washing procedures. All aqueous media are kept refrigerated when not in use or otherwise on ice when feasible (i.e. ‘ice-cold’). We have not systematically monitored the temperature of cell suspensions or media under these conditions, but we aim at maintaining them at 0 4 ° C . We quote this temperature range in this book to indicate all circumstances where we are trying to keep aqueous media and cells in suspension as cold as possible without actually freezing.

2.5. Initial determination of cell quality After the isolated cell suspension has been prepared, it is necessary to determine both the quality and quantity of the isolated cells. This facilitates decisions concerning, for example, the number of incubations that can be run and, perhaps more important, allows an early decision to be made to abort the experiment should the yield or proportion of intact cells be unacceptably low. Unfortunately, neither the determination of cell quantity nor the identification of the proportion of damaged cells present in the preparation are entirely straightforward. Methods for assessing cell yields are presented in Section 2.6, and the issue of cell integrity is discussed in greater depth in Chapter 4.In this Section we present a description of what we consider to be the most rapid and practical way of determining the proportion of damaged hepatocytes in a suspension during the preparative procedures.

46


2.5.1. Staining of cells with trypan blue

The standard and long-established method for identifying damaged cells is by use of the dye, trypan blue (Evans and Schulemann, 1914; Pappenheimer, 1917; Phillips, 1973). In the method used in our laboratory 0.4 g of the dye is dissolved in 100 ml of 0.15 M NaCl and the pH adjusted to 7.4. The dye solution (0.4%) is filtered through a 0.22-pm membrane filter and stored frozen in portions of approximately 10 ml. The bottle in use is kept refrigerated. A quick, albeit approximate, assessment of the proportion of damaged cells can be obtained by placing a drop of cell suspension, mixed with an equal volume of trypan blue solution, on an ordinary microscope slide, covering the droplet gently with a coverslip and examining it under a microscope to determine what proportion of cells stain with trypan blue. While this is useful for a crude estimation of the percentage of cells staining with the dye, damaged cells will tend to distribute at the outer edges of the coverslip when this technique is used. The accurate measurement of the degree of trypan blue staining requires the use of a cell counting device. It is extremely important to develop a standard technique when using this, otherwise inconsistent results are inevitable. Cells settle rapidly in suspension and must be resuspended evenly just prior to sampling. Whenever possible, it is our practice to examine the cells 1 min after mixing the suspension with dye solution, and always to use the same final concentration of dye (0.2%). We use a gold-plated Improved Neubauer haemocytometer (chamber depth 0.1 mm), and a microscope with a 10 x eyepiece and 10 x objective. Other brands of haemocytometer (e.g. Burker) are available. The percentage of cells staining with trypan blue is also affected by pH (Baur et al., 1975), the number increasing as the pH is lowered from 8 to 4. This observation may, in part, explain why the number of cells staining increases with time. The effect is not due to irreversible plasma membrane damage as it can be abolished by returning the pH to 7.4.

Ch. 2.


47

Protocol 2.3 Trypan blue staining for detecting damaged cells (i) Lightly moisten the supporting ridges of a haemocytometer chamber and apply the polished ground-glass coverslip, pressing firmly so that light interference patterns appear. (ii) Pipette into a small vessel, such as a microcentrifuge tube, 50 p1 of trypan blue, 30 pl of washing medium and 20 p1 of well-mixed cell suspension, approximately 30 mg dry weightlml (see Protocol 2.4), and mix gently but thoroughly. (iii) Allow one drop of the well-mixed suspension to flow smoothly by capillary action under the cover-slip of the cytometer, so as to fill the measuring compartment completely, yet not overflow. (iv) The percentage of freshly prepared isolated cells staining with trypan blue remains constant in our experience for at least 30 min on a cooled slide. Thus, if the cells cannot be viewed immediately, store the stained sample on ice. Count at least 200 cells and calculate the percentage staining with trypan blue. If the cell count reveals that more than 10% of the cells are stained, prepare and examine a fresh sample. Should this sample also show more than 10% of cells staining, discard the preparation, or remove the damaged cells (Section 2.8). The working rule in our laboratory is that at least 90% of the cells must exclude trypan blue for the suspension to be considered suitable for experimental work, and we prefer to see a value closer to 95% prior to the incubation. Not infrequently, publications report the use of suspensions where the proportion of damaged cells may be as high as 20%. Values as large as this generally indicate an unsatisfactory preparative procedure, and where feasible it is recommended that such a preparation be discarded and a fresh start made. Under normal circumstances, the preparative procedures described in this Chapter will regularly yield preparations in which less than 10% of the hepatocytes takes up trypan blue prior to cell incubation.

48


2.6. Assessment of cell yield Because of the wide variety of experiments that can be conducted with the isolated hepatocyte preparation, it is not possible to make a single recommendation concerning the amount of cells likely to be required for experimental purposes. Important considerations concerning choice of the optimal quantity of cells for a particular set of incubation conditions are discussed in Chapter 6. Once a decision has been made as to the appropriate quantity of cells to be used in a given incubation, it is necessary to ensure that the correct quantity of cells is dispensed. In this Section we describe two ways to achieve this, based on two different methods for obtaining total cell yield. 2.6.1. Estimation of cell number

Perhaps the most obvious way to determine cell number is to count the hepatocytes. Unfortunately, this is not quite as easy as might be anticipated. Whereas an accurate measure of the proportions of intact and damaged cells can be obtained by counting in a haemocytometer chamber (Protocol 2.3), determination of the total number of cells present is subject to considerable inaccuracies. For instance, the droplet of cell suspension may not spread evenly under the cover-slip, some clumping may occur, or the suspension may not have been mixed or sampled quickly enough, as the cells settle rapidly. It is good practice, therefore, to perform the cell count in duplicate, counting at least 200 cells each time. The central portion of an improved Neubauer haemocytometer is divided into 25 medium-sized squares that are each further divided into 16 small squares. The dimensions of a medium square are 0.2 mm x 0.2 mm. When counting cells, all of those touching the top or right hand edges of each square are included. Any contacting the bottom or left hand sides are ignored. A four-fold dilution of a 30 mg dry weight suspension of hepatocytes (Protocol 2.4) will give 2.5-3.5 x lo6 cells/ml and approximately 10 cells/medium square. Count the number of cells in all 25 medium

Ch. 2.


49

squares in the central section. Since the depth of the chamber beneath the coverslip is 0.1 mm, each medium square represents a volume of 4 X cc. Thus, the number of cells per ml in the diluted sample is calculated by multiplying the average number of cells per medium square by 250,000. About 95-98% of the cells present in the isolated liver cell preparation are hepatocytes, so that it is unnecessary to make an allowance for the presence of other types of cell. In any case, hepatocytes are considerably larger than all other kinds of cell likely to be present. An alternative way of counting cells is by means of a Coulter Counter. This devise is frequently encountered in Clinical Haemotology departments, but its cost is probably too great for most basic science laboratories, particularly since the haemocytometer, a much cheaper alternative, is available. Sweeney et al. (1978) and Otto et al. (1983) used a Coulter counter, model B (Coulter Electronics), fitted with a 100-pm aperture, in conjunction with a Coulter size distribution analyser (Model P-64 or ClOOO) and an X-Y plotter to determine the number and size distribution of a suspension of hepatocytes (0.25 mg wet weight/ml). The cells were suspended in ISOTON (Coulter Electronics) solution, and polystyrene particles or latex particles (17 pm diameter) were used to calibrate the equipment. However, electronic cell counters cannot distinguish between living or damaged cells, suspended particles, or even between single cells and clumps. Hence, it is necessary also to examine the cell preparation microscopically. A large number of workers report their results on the basis of cell number, and this can be taken as a good indication that cell counting is a satisfactory means of determining the quantity of hepatocytes in the preparation. Nevertheless, in our experience the counting of cells is not quite as reliable as other means of determining yield, and for this reason cell counting is not used in our laboratory as a routine procedure for calculating the quantity of cells to add to each incubation vessel. Instead, we prefer to centrifuge the cell suspension, measure the weight of the pellet and use this information to determine how the cell suspension should be diluted and divided.

50


2.6.2. Estimation of hepatocyte mass and dilution of cells to a desired concentration

For metabolic experiments we have found that a convenient amount of tissue that provides sufficient cell mass for metabolite analysis, yet not so much activity that substrates are depleted during the incubation period, yields approximately 30 mg dry weight when 2 ml of suspension is precipitated with perchloric acid (PCA) (Protocol 6.2). To achieve an appropriate dilution of the cells it is necessary to know what volume of newly prepared suspension contains 30 mg dry weight of hepatocytes. Of course, at the time of initial cell isolation it is not possible to obtain instantaneously the dry weight of a measured portion of the suspension. Instead, it is necessary to determine its wet weight, and to assume a wet weighudry weight ratio, based on previous empirical observations. Factors influencing cell water (and hence the pellet wet weighudry weight ratio) are discussed in detail in Section 6.3.1. Our observations indicate that the cell water forms about 70% of the cell weight, and a somewhat lower percentage of the centrifuged pellet due to the presence of entrained water. There can be considerable differences between the ratios of cell pellet wet weight/dry weight under different metabolic conditions. When rats are starved for 48 h hepatocytes lose one-quarter to one-third of their weight, and this loss is mainly water. Hence there is an increase of approximately 3 5 4 5 % in the number of cells/g pellet wet weight in comparison with cells from fed animals (Harrison, 1953; Otto et al., 1983). Moreover, the intracellular volume of cells cooled to 0 4 ° C is substantially greater than that of cells immediately after incubation at 37°C. Also, the time of addition of Ca2+ during preparation of isolated hepatocytes affects intracellular volume. These differences are reflected in the centrifuged pellet wet weightldry weight ratios. The quantity of entrained water will also vary, depending on the speed of centrifugation. It follows that to achieve a consistent pellet wet weight/dry weight ratio from experiment to experiment, it is necessary to maintain a regular protocol when

Ch. 2.


51

manipulating the cell suspension. The procedure for achieving a consistent cell dilution commences following the final 4-min centrifugation step of the cell washing procedure (Protocol 2.2).

Protocol 2.4 Preparation of cell suspension containing a desired weight of cells (i) After weighing the cell pellet, add 4.5 ml washing medium for each gram of cells (pellet wet weight) to a 50-ml measuring cylinder. Pour about half this medium into the centrifuge tube and gently mix cells and medium to give a homogeneous suspension. Transfer the suspension to a 125-ml conical flask using the remaining washing medium to rinse the centrifuge tube. (ii) Take two 500-pl samples of the thoroughly mixed suspension and centrifuge them in tared microcentrifuge tubes (Eppendorf, Model 3810) at 12,000 x g for 2 min in a microcentrifuge (Eppendorf centrifuge, model 5415). Decant the supernatant and recentrifuge the tubes for a further 2 min. Aspirate the residual supernatant with a syringe and weigh the pellet. The sum of the pellet weights gives the wet weight of cells in mg (P) in 1 ml of suspension at U " C . If the pellet weights vary by more than lo%, repeat the determination with fresh samples. (iii) Calculate the required extra volume of medium (V,) necessary to add to the suspension (VJ, to give the desired cell concentration according to the equation:

v,

=

P x X

v, - vs

If a negative value for V, is obtained, the suspension is obviously too dilute to yield the desired concentration. The cell suspension should be allowed to stand on ice for a few minutes, during which time the cells tend to settle out. The necessary amount of medium can then be removed from the top of the flask. The factor, X, is determined experimentally and will vary according to the degree of swelling

52


associated with the preparative technique. In our hands, using a fasted rat, a value of 97.5 yields a suspension containing 30 mg dry weight of celldml.

=

2.6.3. Cell yields

The number and mass of parenchymal cells per cc of fresh rat liver tissue changes markedly during foetal and post-natal life (Greengard et al., 1972). Variation in the percentages of haematopoietic and Kupffer cells and the volume of parenchymal cells are responsible for the changes during foetal life, while the ploidy state and volumeincrease per parenchymal cell are the main contributing factors during the post-natal period (Epstein, 1967; Wheatley, 1972; James et al., 1979). As mentioned in Section 2.6.2, the nutritional state of the donor affects the number of parenchymal cells per cc, with fasting for 48 h increasing this value by 3 5 4 5 % due mainly to loss of glycogen stores and water (1 7-20%~ and 70'%,respectively, of total cell weight loss). The total number of hepatocytes per liver, in the individual animal, remains relatively constant after the first 6 weeks of post-natal life (Epstein, 1967; Greengard et al., 1972; James et al., 1979) and also after short-term starvation (Harrison, 1953). This value (approximately 1O9 hepatocytes per liver) is thus relatively constant for rats weighing from 150 to 300 g (about 7-20 weeks of age for hooded Wistar rats, which are commonly employed as donors). The yield of isolated hepatocytes in the final preparation can thus be expressed as the total number of cells per liver or as a percentage of this maximum theoretical yield. The majority of workers, however, express yield in terms of the number of hepatocytes isolated per g of liver. Values of between 30 and 60 x lo6 cells/g liver have generally been reported in preparations from either fed or fasted rats using standard techniques or minor modifications of them. At least two groups (Drochmans et al., 1975; Gustavsson and MBrland, 1980) have obtained values as high as 70 x loh cells/g liver. For comparative purposes the nutritional state of

Ch. 2.


53

the animal must be given. Rats fasted for 24 h, and weighing between 150-200 g, have approximately 130 x lo6 hepatocyteslg of fresh liver (Weibel et al., 1969)when allowance is made for a 20%)binuclearity of parenchymal cells (Wheatley, 1972; Seglen, 1973). Using heavier rats (approx. 290 g; nutritional conditions unstated) Greengard et al. (1972) have found approximately 95 x lobparenchymal cells/g of intact liver. Thus, a yield of 30-60 x lo6 hepatocytedg from a fasted rat represents about 25-50% of the maximum theoretical yield. Our usual cell preparations from fasted rats have average yields of 60 x lo6 hepatocytedg or 45-50%) of the maximum theoretical yield. Data concerning the number of hepatocytes per cc provided by Greengard et al. (1972), should be useful for determining cell yields during other stages of rat development. There may also be differences due to species or strain, as well as the pre-treatment of the rat prior to isolation of hepatocytes. The one-step or two-step procedures performed under optimal conditions appear to give similar yields. If we assume that 40% of a 10 g liver is converted to isolated cells, this is equivalent to P total cell yield of at least 4 x lo8. For the vast majority of experiments, the numbers of cells used per incubation vessel ranges from 106-107. Hence, a reasonable preparation will yield sufficient cells for 4-00 incubation vessels! Thus, cell yield should not be a serious limitation. In those instances where analytical requirements demand that the number of cells in each vessel be as high as lo7, the yields from two rat livers may be combined.

2.7. Reasons for unsatisfactory preparations The methods for preparation of isolated hepatocyte suspensions, described in this Chapter, have been in use in the author’s laboratory for 20 years, and have been used extensively by other workers for at least 15 years. The thousands of published papers, describing work using isolated hepatocytes, attest to the reliability of the technique. Nevertheless, it is recognized that on occasion a poor quality prepara-

54


tion may result. The yield may be low, or alternatively an unacceptably high proportion of cells may stain with trypan blue. Often in such circumstances,both yield and integrity are down. Moreover, many workers have infrequently encountered the alarming situation where the poor quality preparation becomes not the exception but the rule. 2.7.1. Causes of poor quality preparations

A frequent cause of an occasional poor quality preparation of cells is the use of a faulty medium. This may be due to poor quality water, a contaminated preparative reagent, a solution made up incorrectly or subsequently contaminated. A possible source of contamination is detergent not adequately removed after washing of glassware. We soak all glassware in 1% Extran 300 (British Drug Houses), and then rinse each piece three times with tap water and then three times with filtered water (see Section 2.2.3). Contamination problems can usually be eliminated by a complete change of preparative reagents and by paying careful attention to washing up procedures. Other causes of poor quality yields are incorrect perfusate temperature or other problems arising during the perfusion. These include obstruction to the outflow that causes the liver to swell, or puncture of the portal vein by the inflow cannula, which prematurely ends the perfusion. These mishaps are readily identified and with a little practice can be avoided. More difficult to eliminate are circumstances whereby a series of perfusions result in poor yields with an unacceptably high proportion of cells staining with trypan blue. Fortunately, such occurrences are rare, but when they do occur, elucidation of the cause may be difficult or perhaps impossible. The first step is to discard all preparative solutions and make up fresh ones. If this fails to remedy the problem, it is necessary to examine a wide range of potential causes, such as the health of the donor rats, the cleanliness of the perfusion apparatus or the quality of the collagenase. In one instance, for example, it was found that the time-clock governing the hours that the rats were exposed to light had failed. Nevertheless, despite these very occasional

Ch. 2.


55

and sometimes inexplicable set-backs, adequate preparations will regularly be achieved by the procedures described in this Chapter. There is no evidence that either the one-stage or two-stage procedures are more susceptible to failure, provided an adequate pre-perfusion time is allowed in the latter.

2.8. Removal of damaged cells from the preparation As indicated in Section 2.5, it is considered highly undesirable to use for experimental work hepatocyte preparations in which more than 10% of the cells stain with trypan blue. In general the best approach, when faced with a poor quality preparation, the reason for which is known, is to discard it and start again. However, on occasion the supply of hepatocytes is limited, because the donor rats have been specially treated and are few in number, so that it becomes imperative to use the available preparation. A preparation with a high proportion of damaged cells is hardly suitable for metabolic studies, and hence it is necessary to remove the damaged hepatocytes before commencing experimental work. Damaged hepatocytes are much more susceptible to digestion by trypsin than are intact cells, and this has led to the suggestion that damaged cells can be removed from the preparation by treatment of the suspension with trypsin (Bellemann et al., 1977). While such an approach is feasible, it has the disadvantage of exposing the hepatocyte plasma membranes to a high concentration of a proteolytic enzyme, known to be capable of causing damage to surfaceassociated components such as receptors. In an alternative and preferred approach, damaged cells are removed by Percoll treatment (Kreamer et al., 1986). The method takes advantage of the fact that damaged cells will float, and intact hepatocytes will pellet, in a 1.06 g/cm3 density ( p ) isotonic Percoll solution. Centrifugation of hepatocytes through 5-300/0 Metrizamide ( M r 789) gradients (Munthe-Kaas and Seglen, 1974) or onto a cushion of buffered 30% Metrizamide ( p = 1.16) (Seglen, 1976) has also been


56

used to separate intact and damaged cells. Under these conditions damaged cells sediment to the bottom of the tube, while intact cells either form a well-defined, homogeneous peak at p = 1.12 or remain at the interface. Metrizamide apparently penetrates the plasma membrane of damaged cells making them more dense than intact cells. It seems that Percoll, a colloidal suspension of silica particles coated with polyvinylpyrrolidine, is unable to enter either damaged or intact cells and in this situation, the swollen damaged cells, because of their larger water space and loss of cytoplasmic proteins, have a lower density than intact cells (Seglen, 1979).

Protocol 2.5 Removal of damaged hepatocytes by treatment with Percoll (i) Prepare 102 ml isotonic Percoll solution as follows: 90 ml Percoll (Pharmacia); 10 ml stock balanced salt solution (Protocol 2.1); 2 ml stock phosphate buffer (Protocol 2.1).

Adjust pH to 7.4 with 0.1 M HCl. The composition of stock and working solutions is given in Section 2.2.3. (ii) Dilute the final cell suspension (30 mg dry weight/ml) 1:1 with washing medium. This will give approximately 5 x lo6 cells/ml. (iii) Add equal volumes of cell suspension and Percoll solution to 50-ml centrifuge tubes. Mix gently and centrifuge at 0--4"C for 10 min at 50 x g . Cells capable of excluding trypan blue will pellet, whereas damaged cells, non-parenchymal cells, cell aggregates and debris will float in the upper region of each tube. (iv) To remove the Percoll, aspirate the supernatant and gently resuspend the pelleted cells in washing medium and centrifuge for 3 min at 40 x g (0-4OC). Repeat this washing step. Resuspend the pelleted cells in two-thirds of the volume of medium in which they were suspended before Percoll treatment. Carry out a determination of wet

Ch. 2.


57

weight according to steps (ii) and (iii) of Protocol 2.4. Store the cells on ice. This procedure is considerably simpler than those involving use of a Percoll gradient (Brown and Bidlack, 1988). Memon et al. (1989) demonstrated that high-speed centrifugation of hepatocytes through such gradients impaired the ability of insulin to stimulate both the oxidation of labelled succinate to CO, and the incorporation of carbon atoms from this compound into protein. Results obtained with succinate are of questionable significance since, as pointed out in Section 4.2.2, the ability of succinate to penetrate intact cells and hence be oxidized, is very low. Nevertheless, there is merit to the recommendation of these workers that results obtained with cells purified by methods that include high-speed centrifugation through Percoll be compared with those obtained from cells prepared without the inclusion of such a step. Note that the method in Protocol 2.5 employs only low-speed centrifugation.

2.9. Storage of hepatocytes 2.9.1. Short-term storage

For certain types of experiment it is necessary to store the isolated cell suspension for periods up to an hour or more. The question arises as to whether such cells can be maintained in a state suitable for experimentation. We have explored the effects of temperature and oxygenation of buffers on the stability of cells during storage. Cells stored on ice maintain, for at least 1 h, ATP levels and normal synthetic rates on subsequent incubation. There appears to be no requirement for oxygenation during storage. Cells stored at ambient temperature are also capable of maintaining ATP concentrations and synthetic rates, provided that oxygenation and continuous shaking of the cells is employed during the storage period. However, since this

58


procedure inevitably results in a significant degree of cellular metabolism, storage on ice is the recommended approach. 2.9.2. Long-term storage

Farrell and Lund (1983) have found that hepatocytes from rats, starved 48 h, could retain about 80% of their capacity to synthesize glucose from lactate, urea from ammonium chloride, and acetoacetate and 0hydroxybutyrate from oleate under appropriate storage conditions. The hepatocytes were stored in a shallo,w layer under an atmosphere of carbogen or N,:CO, (955) at 0-4"C for 24 h, in the presence of the substrates lactate (10 mM), pyruvate (1 mM), oleate (0.5 mM), NH,CI (10 mM) and ornithine (1 mM); the inclusion of 1.3-3.75% (w/v) albumin was important for cell stability. It was later recognized (Lund and Farrell, 1985) that the albumin (provided it contained very low levels of Cu2+)was protecting the cells from damage, by scavenging free-radicals produced from some of the substrates. Lund and Wiggins (1988) demonstrated that hepatocytes stored for 24 h with 10 mM xylitol, sorbitol or glycerol as substrates, under an atmosphere of carbogen or N,:CO, (955) at 0-4"C, lost substantial amounts of cellular ATP and total nucleotides but their rates of glucose synthesis increased. Hence, it seems that cellular ATP and total nucleotide concentrations cannot always be used as a precise measure of cell integrity. Gluconeogenic substrates such as xylitol appear to act as freeradical scavengers. This demonstration that isolated hepatocytes can tolerate substantial periods of storage at 0 4 ° C is encouraging. However, where possible, the experimental incubation of cells as soon after preparation as is feasible is recommended. Hepatocytes have also been successfully maintained for extended periods by means of cryopreservation (see Section 12.11.3).

CHAPTER 3

Other methods for hepatocyte isolation

3.1. Introduction The one- and two-step procedures described in Chapter 2 are the methods of choice for the isolation of intact hepatocytes in high yield. The key feature of these methods is perfusion of the liver with a medium containing collagenase. A number of methods, including the all important contribution of Howard et al. (1967) have not employed liver perfusion, but instead have relied on incubation of liver slices with collagenase. In other procedures collagenase has been omitted. In this chapter these variations will be evaluated. In addition, the modifications to the recommended method that are required for species other than the rat and for foetal or neonatal animals, will be discussed.

3.2. Preparation of isolated hepatocytes from liver slices In 1967, Howard et al. described the first successful method for preparing intact isolated hepatocytes. These workers injected a balanced saline solution (Hanks’ medium), free of added Ca2+and containing 0.05% collagenase, into the major sinusoids of a rat liver and then cut

60


the liver into thin slices that were incubated in collagenase solution. A considerable quantity of intact isolated parenchymal cells (up to 5% of total liver volume) was obtained. Since then, a number of other workers have used a slice-making technique in the belief that preparing slices is simpler than perfusing the liver. Also it has been argued that for large animals, such as sheep or goats, or for preparation of cells from human liver, perfusion is not feasible. Preparation of isolated hepatocytes by procedures which omit a perfusion step are not recommended and therefore, a detailed description of such methods is not provided. The yield of cells by these techniques is always very low and the proportion of damaged cells greater (Carlsen et al., 1981). In consequence, a very large amount of cell debris is present and it can be anticipated that enzyme release from the large number of damaged or broken cells is likely to have deleterious effects on the undamaged hepdtocytes. Moreover, it is very difficult to maintain adequate oxygenation in sliced material (Umbreit et al., 1964). It is not surprising, therefore, that a number of studies (Gustavsson and Msrland, 1980; Carlsen et al., 1981; Sturdee et al., 1983) have shown that cells prepared by non-perfusion techniques emulate the behaviour of hepatocytes in vivo to a much lesser degree than do cells prepared by collagenase perfusion. The liver perfusion procedure described in Chapter 2, in which the liver is not excised, is a relatively simple one. Indeed contrary to what might be thought, it is considerably easier to perfuse a liver than to cut slices of desirable thinness from it. Moreover, it has been shown that use of a perfusion technique is often feasible even in the case of cell preparation from large animals. Hepatocytes from cattle, sheep, or goats, for example, can be obtained by removing and perfusing the caudate lobe (Clark et al., 1976; Shull et al., 1986; Aiello and Armentano, 1987). Perfusion of hurnan cadaver liver or biopsy specimens is also feasible by cannulating veins exposed on sectioning the extirpated specimen (Houssin et al., 1983). However, it is recognised that a situation may arise where liver perfusion is anatomically not feasible, as in certain species of fish.

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If the use of sliced material is unavoidable, readers can use the methods described by Howard et al. (1973) or by Fry et al. (1976). The latter authors claimed that if liver slices are shaken with a medium containing a Ca2+-chelatingagent, prior to treatment with collagenase dissolved in a Caz+-containingmedium, yields of cells twice as great as those obtained by Howard et al. (1973) are obtained. We have not attempted to validate this. Both methods give an unacceptably high proportion of damaged cells, but most of these can be removed by centrifugation through Percoll-saline (Protocol 2.5). Nevertheless, where hepatic perfusion is possible, it should be employed (Green et al., 1983; Pogson et al., 1984), preferably through the portal vein rather than retrograde through the vena cava (Klaunig et al., 1981a).A survey of the current literature indicates that the vast majority of scientists who prepare isolated hepatocytes, choose to do so by techniques employing collagenase perfusion of the liver via the portal vein.

3.3. Preparation of intact isolated hepatocytes without collagenase Concern has frequently been expressed that treatment of liver cells with a crude collagenase preparation that also contains other proteolytic enzymes might lead to deleterious alteration of their surface properties (Quibel et al., 1986; Marteau et al., 1988; Meredith, 1988). In fact, experimental studies of these surface properties (Carlsen et al., 1981; see also Chapter 8) indicate that, in general, cells isolated by collagenase perfusion retain binding proteins, surface receptors and plasma membrane transport functions. Even so, it may be necessary in an experimental study to demonstrate that a particular property of the isolated hepatocyte preparation is not merely an artefact of exposure of the cells to collagenase. One approach to this is to prepare intact cells without the use of collagenase (Berry et al., 1983; Wang et al., 1985). It may not be widely appreciated that it is feasible to prepare intact

62


cells without the use of collagenase or other proteases. The method is based on the premise that removal of Ca2+is more crucial for cell separation than is digestion of connective tissue. However, for the method to be successful, it is necessary to loosen gap and tight junctions, as well as desmosomes, and to achieve this, Ca2+-removalmust be virtually complete. Hence a chelating agent is essential and in our hands, perfusion must be carried out for at least an hour if reasonable yields of cells are to be obtained. In consequence, the hepatocytes are exposed to a medium very low in Ca2+for much longer than occurs during collagenase perfusion and this may be detrimental to their subsequent performance. Nevertheless, hepatocytes isolated without collagenase have been successfully used for certain studies (Berry et al., 1983; Wang et al., 1985), and it can be assumed that they will prove valuable in a variety of circumstances, provided that their potential limitations are recognised. The method may be particularly advantageous for preparing cells for culture. Recently, Meredith (1 988) compared cells grown in monolayer culture after preparation either by the collagenase-free technique or by the standard collagenase method. Cells prepared without collagenase were able to maintain total cytochrome P-450 concentrations over 5 days in culture, whereas values decreased rapidly in cells that had been exposed to collagenase. Glutathione levels remained more stable and did not exhibit the marked decline shown in cells prepared with collagenase. Moreover, the marker of dedifferentiation, y-glutamyl transpeptidase did not increase over 5 days in cells prepared without collagenase, in contrast to the rapid rise in cells prepared by the enzymic procedure. Meredith (personal communication) now reports yields of intact cells similar to those obtained by Berry et al. (1983). He finds that the perfusion time can be shortened by doubling the normal flow rate of perfusate through the liver, a process that inevitably causes the organ to swell. This procedure is not recommended for preparing cells for immediate study. However, Meredith reports that cells prepared in this manner are excellent for purposes of cell culture.

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3.3.1. Apparatus The apparatus is a simplified version of that used for collagenase perfusion (Section 2.2.2). The constituents of the collagenase-free perfusion medium are sufficiently inexpensive to permit a single-pass petfusion. Nevertheless, because of the possibility that a nonrecirculating perfusion can seriously deplete the liver of those metabolites capable of crossing the plasma membrane, a recirculating system is preferred. 3.3.2. Reagents

The perfusion medium contains no collagenase and includes 2 mM EDTA, but otherwise is identical with that used for the normal onestep procedure (Section 2.2.3, Protocol 2.1). The washing medium is similar in composition to the perfusion medium, but 1.3 mM CaCl, is added and bicarbonate and EDTA omitted.

Protocol 3.1 Intact rut hepatocyte preparation without collagenase (i) Establish hepatic perfusion by following the steps set out in Protocol 2.2, allowing the first 50-100 ml to flow to waste. The flow rate for the recirculating medium (Section 3.3.2) should be about 4 ml/g liver. (ii) Continue the perfusion for 60 min. During this period the liver will become softer and ooze freely, though not to the degree seen during perfusions with media containing collagenase. (iii) Remove the liver, mince it lightly with scissors and break it up with a blunt spatula in about 20 ml of washing medium at 0 4 ° C . Add further medium to make the suspension up to 60 ml and filter it through coarse nylon mesh (250 pm pore size). (iv) Centrifuge the cells at 50 x g for 2 min and resuspend the pellets

64


in 60 ml of ice-cold washing medium. Repeat this process twice, filtering the suspension through 100-pm nylon mesh before the final centrifugation. (v) The damaged cells usually represent 40-60% of the total cell population of the suspension (as determined by trypan blue staining). Remove these by treatment with Percoll, according to Protocol 2.5. The yield of cells by this method is, as expected, somewhat lower than for the standard collagenase preparation. Although collagenasefree perfusion ultimately produces a high quality preparation of isolated hepatocytes, it will be appreciated that the generation of substantial numbers of damaged cells during the procedure is unavoidable - hence the requirement for the use of Protocol 2.5. It is feasible that enzymes leaking from such damaged cells could have deleterious effects on the surface of intact cells, but this possibility has not been adequately explored. In most circumstances, the collagenase perfusion technique for hepatocyte preparation is the method of choice.

3.4. Preparation of isolated hepatocytes from species other than rat A wide range of species, both homeothermic and poikilothermic, have been used for the preparation of isolated hepatocytes. Poikilothermic species include frog, fish and eel, while homeotherms that have proved suitable as liver cell donors embrace farm, domestic and laboratory animals. In most instances cell preparation can proceed along standard lines (Chapter 2) regardless of the nature of the animal. However, in some cases, special approaches are required, and these are detailed below. 3.4.1. Preparation of isolated hepatocytes from mouse

In recent years, mice have been found to be very useful models for

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a number of disease states relevant to man. The development of techniques for the preparation of isolated hepatocytes in mice therefore allows the opportunity to study hepatic function in such disease models. Although the yield of hepatocytes from a mouse liver will inevitably be small, a more serious concern is the metabolic adequacy of the final preparation. Many variations of the original one-step method of Berry and Friend (1969) and two-step procedure of Seglen (1972, f976) have been applied to the isolation of mouse hepatocytes. A substantial number of workers report methods that yield cell suspensions showing levels of trypan blue exclusion below 90'%1,suggesting that mouse hepatocytes may possess increased fragility when compared with cells from the rat where, with the recommended techniques, less than 5% of the cells may take up dye. Because of the small size of the liver, retrograde perfusion of the organ via the thoracic portion of the inferior vena cava, as introduced by Renton et al. (1978), has been extensively employed. These authors reported dye exclusion by about 95% of the cells, and a typical recovery from a 30-g mouse (liver weight 1.6 g) of 100 x lo6 cells, i.e. a 50% yield. Although, more recently, Soley and Hollenberg (1987) achieved comparable results employing a similar technique, other laboratories have been less successful, obtaining cell suspensions with percentage dye exclusion ranging from 80 to 90% (Edstrom et. al., 1983, Harman et al., 1987; Richieri and Buckpitt, 1988). Although technically more difficult, perfusion through the portal vein does not require an inordinate degree of skill. Klaunig et al. (198la) compared factors influencing the yield and integrity of mouse hepatocytes in detail, including a comparison of portal vein and inferior vena cava perfusion. These workers concluded that the former gave higher yields (77 x 106 cells/liver for the portal route compared with 69 x lo6 cells/liver for the retrograde method using mice weighing 30-35 g), provided that the inferior vena cava was occluded intermittently during the perfusion via the portal route, a technique that caused the liver to swell to twice the unoccluded size. Although 92% of the cells prepared by either method showed trypan

66


blue exclusion, in our experience techniques that cause the liver to swell abnormally during perfusion can produce inferior preparations and are to be discouraged. It is noteworthy that electron micrographs of cells isolated by this approach demonstrated features similar to those seen in damaged rat hepatocytes. Relatively few other descriptions of methods using portal perfusion for mouse hepatocyte isolation have been reported. Maslansky and Williams (1982), using a two-step portal perfusion technique, were able to obtain a yield of only 41 x lo6 cells from the livers of mice (20-30 g) with an average dye exclusion of 89%. Using similar techniques, Ciccia-Torres and Dellacha (1985) obtained a comparable yield of cells with a dye exclusion of 80-90%. One point of note is that Maslansky and Williams (1982), Renton et al. (1978) and Klaunig et al. (1981a) used two-step methods in which the initial perfusion time with Caz+-freemedium was reduced to 2 4 min before the addition of Ca2+and collagenase. Although direct experimental evidence is lacking in mice, the possibility exists that yield and integrity could be improved with a lengthening of this step as employed for rats (see Section 2.3.2). Elliott et al. (1976) used a procedure only slightly modified from the original one-step method to prepare mouse hepatocytes where dye exclusion was routinely at least 95%, but actual yields were not documented. Likewise, Mandl et al. (1979) used a comparable technique where livers were perfused with 0.02% collagenase solution for 30 min. These workers reported dye exclusion of 85-90%. In summary, the attainment of good yields of metabolically competent, undamaged mouse liver cells should be possible by both the retrograde and portal perfusion methods if care is taken to optimize conditions of isolation as described for the rat in Chapter 2. In general, lower flow rates (8-10 ml/min) consistent with the small size of the mouse liver and a similar collagenase concentration, are employed over a somewhat shorter perfusion time than for the rat (Harman et al., 1987), without recirculation of the perfusate. The mouse, unlike the rat, possesses a gall bladder which should be removed before freeing the hepatocytes from the liver capsule.

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3.4.2. Preparation of isolated hepatocytes from guinea pig

Hepatocytes from guinea pigs have been prepared by both the oneand two-step procedures (Elliott and Pogson, 1977; Burger et al., 1985; Dodgson and Forster, 1986). Elliott and Pogson (1977) perfused in a retrograde manner without cannulating the portal vein, on the grounds that this allowed more rapid surgery. Subsequent workers have chosen to employ portal vein perfusion. A preliminary flush of the liver with a solution containing EDTA, as used by some workers, does not seem necessary. Hence the preparative technique for guinea pig hepatocytes is virtually identical to that for isolated rat hepatocytes. 3.4.3. Preparation of isolated hepatocytes from ruminants Ash and Pogson (1977) prepared isolated hepatocytes from adult sheep liver by incubating surviving slices with collagenase. However, a more desirable approach is that adopted by Clark et al. (1976). These workers used only the caudate lobe which represents about 4.5% of the total liver weight. The lobe was excised from the liver in situ by a single transverse cut, weighed and placed on a tared polypropylene tray with the portal vein uppermost. The tray and lobe were positioned in the perfusion cabinet and a tapered cannula fitted into the portal vein. Venous outflow drained directly into the perfusion system. Median lobe liver cells were prepared by sectioning the median lobe. Other details were as for the one-step procedure. More than 90% of isolated hepatocytes excluded trypan blue, and gluconeogenic rates and adenine nucleotide content showed good agreement with levels in the perfused and freeze-clamped liver, respectively. Lomax et al. (1983) used a modification of the two-step method to prepare cells from the caudate lobe of sheep. Following a 5-min initial perfusion with Ca2+-freebuffer containing 0.5 mM EGTA, the lobe was perfused with medium containing collagenase and 2.3 mM CaCl,. Dispersion and washing steps were carried out with buffer to which Ca2+and 2% albumin had been added. Aiello and Armentano (1987) have used a similar method to prepare hepatocytes from the caudate

68


lobe of goats and obtained dye exclusion values greater than 90%. In 1985, Shull and co-workers developed a technique for the preparation of isolated hepatocytes from cattle (Forsell et al., 1985). This technique, as later improved by the same group (Shull et al., 1986), involves perfusion of the caudate process of the caudate lobe excised from anaesthetized cattle, which eventually make a full recovery. In these workers’ hands, optimal results were obtained with the two-step method, supplementation of media with volatile fatty acids and the use of Worthington CLS type I1 collagenase. The average value for dye exclusion by the cells was 85%, which is lower than that usually obtained with the rat, probably due to the longer perfusion times required (Shull et al., 1987). 3.4.4. Preparation of isolared hepatocytes ,from avian species

The anatomy of birds, particularly the lack of a diaphragm, is not conducive to portal perfusion. Accordingly, most workers have used retrograde perfusion of the liver. The technique described by Mapes and Krebs (1978) involves anaesthetizing a chicken with pentobarbital. To minimize the anoxic period between the exposure of the liver and the start of the perfusion, the body cavity is opened and a cannula passed through the right atrium of the heart into the inferior vena cava, and tied so that it rests just proximal to the entry of the hepatic veins. Perfusion of oxygenated medium is commenced immediately after insertion of the cannula, and the portal vein is cut to allow the perfusion medium to flow through the liver. The gizzard and intestines are then removed to form a cavity into which the perfusion medium drains. The first 80-100 ml is discarded and the remaining perfusion medium recirculated, as for the one-step procedure. A similar approach by Burns and Buttery (1984) has employed a two-step procedure. Schultz and Mistry (1981) have described a one-step procedure in which chicken liver is perfused in the normal direction, through the anterior coccygeomesenteric vein. The cells were shown to be capable of gluconeogenesis, but no comparisons in regard to cell quality were

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made with suspensions prepared using retrograde perfusion. In the light of anatomical considerations, the retrograde approach would seem to be the method of choice. More recently, Cross and Dodds (1988) have used the portal route, after cannulating the trachea and employing a respirator. They used a two-step procedure, but reported poor yields unless 0.1% hyaluronidase was included. Even so, the perfusion needed to be continued for 60 min. The investigators used a discontinuous density gradient formed with Nycodenz (Nyegaard) to separate ‘fatty’ from ‘non-fatty’ hepatocytes.

3.4.5. Preparation of isolated hepatocytes from fish A review of publications concerning the preparation of fish hepatocytes indicates that workers have favoured the two-step approach, though the reasons for this are not discussed. In the majority of species used such as the European eel (Anguilla unguilla L.) or the rainbow trout (Salmo gairdnerii),the anatomy of the species allows portal perfusion SO that the technique employed is essentially that for the preparation of rat hepatocytes. The fish is anaesthetized in a solution containing MS 222 (tricaine methanesulfonate). Perfusion is carried out at about 20°C. In general, HEPES buffer is preferred to bicarbonate. and 0, rather than carbogen used as the gas phase. Inclusion of 2 mM EDTA in the pre-perfusion medium has also been favoured (Jankowsky et al., 1984), but no systematic study of the need for this appears to have been carried out. For teleosts, a lower NaCl content (teleost Ringer solution of Young, 1933) is used in the perfusion medium (Renaud and Moon, 1980). In some species perfusion via the portal vein or even retrograde perfusion of the liver is anatomically not feasible. To overcome this difficulty, perfusion through the bile duct has been successfully employed (Morrison et al., 1985). Other workers have incubated liver fragments from various fish including the coho salmon (Oncorhynchus kisutch) (Plisetskaya et al., 19841, goldfish (Cavassius auratus) (Birnbaum et al., 1976)and carp (Cyprinus carpio) (Saez et al., 1982), an approach to be avoided where possible for the reasons stated in Section 3.2.

70


3.4.6. Preparation of human isolated hepatocytes

In studying the mechanisms of hepatotoxicity induced by drug administration, a dilemma is posed by the fact that certain forms of toxicity are confined to only a few species and sometimes only to man. Because such forms of toxicity can be satisfactorily studied only in man, a number of groups have endeavoured to prepare isolated hepatocytes from human liver. There are, of course, considerable limitations to obtaining high quality specimens of sufficient size for perfusion techniques. Bojar et al. (1976) perfused the whole liver of a renal transplant donor, maintained on a support system, and obtained cells of high quality. Other less ambitious approaches have limited the perfusion to a single lobe, thereby greatly reducing the amount of collagenase required (Guguen-Guillouzo et al., 1982). Smaller yields are obtained if biopsy specimens are taken and the samples cut up into small fragments, which are then incubated with collagenase (Maekubo et al., 1982). While this may be the only feasible method for dealing with a very small biopsy specimen, for instance from foetal liver (Section 11.11.2), the perfusion technique is to be preferred when larger samples are available. Ideally, these should be from organ donors maintained on life-support systems, or patients undergoing partial hepatectomy where the deterioration associated with an autopsy specimen can be avoided. Tee et al. (1985) provide a comprehensive account of the preparation of human hepatocytes by a three-step procedure, from patients undergoing partial hepatectomy. One or more branches of the portal vein are identified and cannulated and the sample flushed immediately with ice-cold saline medium (Dulbecco and Vogt, 1954). Perfusion is then carried out with Ca2+-and Mg2+-freeEarle’s balanced salt solution (Earle, 1943) containing gentamicin, bicarbonate and 0.5 mM EGTA, the gas phase being carbogen. In the first stage of the perfusion, the perfusion medium, which is not re-circulated, is passed through the liver for 5-10 min at a rate of 5-10 ml/min per g. This rate is substantially higher than that employed for the rat.

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In the second stage of the perfusion, the EGTA is flushed from the liver by a 7-15 min perfusion, without recirculation, of 1-2 1 of Earle’s medium, devoid of EGTA. Following Seglen (1976), a Ca2+ concentration of 5 mM is recommended by the authors for inclusion in the collagenase-containing medium in the final stage. As discussed earlier (Section 2.3.1), 1 mM Ca2+is considered adequate. Tee et al. (1985) employed foetal calf serum in their dispersion and incubation media, although the advantages of this are not discussed. One obvious disadvantage is that such media are not characterized, and hence it is difficult to attribute changes in metabolic behaviour to specific components. At the end of the perfusion the liver tissue is transferred to a petri dish, any capsule is broken open and the dissociated cells are teased from the specimen by stroking the surfaces of the sample with a glass rod. Although a substantial amount of hepatic tissue remains undissociated, an adequate yield can usually be obtained. The crude hepatocyte suspension is filtered through nylon bolting cloth and the cells washed in the usual manner. The yield and viability of human hepatocytes resulting from this technique is very variable. The data of Tee et al. (1985), for preparations from six donors, indicate that in only one case did less than 30% of the hepatocytes stain with trypan blue, and in fact in two preparations, over 60% of the cells were sufficiently damaged to take up the dye. It would seem highly desirable, therefore, to subject such suspensions to Percoll purification (Protocol 2.5), before using them for metabolic studies. Reese and Byard (1981), employing a similar technique on large biopsies from two donors only, obtained dye exclusion values of 75% and 95%. Houssin et al. (1983) perfused a portion of the left lobe of the livers of three kidney donors, according to Guguen-Guillouzo et al. (1982), and reported average values for dye exclusion of 85%. A major factor influencing the success of the reported techniques is the actual configuration of the sample used, including the sites that are available for cannulation (Reese and Byard, 1981). Fabr6 et al. (1988) have described a method for preparing isolated hepatocytes from whole adult liver. The perfusion is performed by us-

72


ing a Travenol peristaltic pump capable of a flow rate of 0.5-6 I/min, a recirculating water-bath at 38°C and a membrane oxygenator. The method employed was essentially a two-step procedure, with HEPES (10 mM) serving as buffer. Livers were obtained from organ donors maintained on life support systems, and the livers flushed in situ through the portal vein and aorta with 6-10 1 of sterile ‘Eurocollins’ medium (a solution used to preserve the function of the extirpated kidney prior to its transplantation) at CL4”C. After removal of the organs to be transplanted, the liver was washed with a further 2 1 of cold ‘Eurocollins’ medium through the portal vein to improve the elimination of erythrocytes. With this protocol the ischaemic period never exceeded a few minutes, and the time taken for transportation of the liver from theatre to laboratory was never greater than 15 min. Under these circumstances, it is perhaps not surprising that the investigators were able to obtain suspensions of hepatocytes, of which more than 85% excluded trypan blue. The facilities required for this approach are obviously beyond the reach of any group not connected with an organ donor program. However, it is by no means certain that whole liver perfusion is the best approach. since the cost of the collagenase required is substantial and the quantity of cells obtained is excessive, so that most will be discarded. In any case, the increasing demand for human livers for therapeutic transplantation is likely to diminish greatly their availability for experimental purposes! 3.4.7. Preparation of isolated hepatocytes from other species

It is not feasible to discuss all the methodologies relating to the various different species that have acted as donors for isolated hepatocytes. The livers of species as diverse as the wallaby (Janssens et al., 1977) and the frog (Wangh et al., 1979) have successfully been used for this purpose. A list of examples other than those already discussed in the text is given in Table 3.1 together with references that, as far as can be ascertained, contain definitive methods.

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TABLE3. I Additional species from which hepatocytes have been isolated Species Monkey

Rhesus Cynomolgus

Pig

References

Comments

Glinoer et al.(1976) Poole and Urwin (1976) Seddon et al. (l989b)

Whole liver,

Makowa et al. (1980) Nordinger et al. (1985)

Whole liver Portion of liver Whole liver, newborn pigs

Pegorier et al. (1982) Rabbit

Zaleski and Bryla (1977) Maslanksy and Williams ( 1982)

Hamster

Blaich et al. (1987) Rognstad and Wals (1976) Maslansky and Williams ( 1982)

Cat

Silva and Mercer (1985)

Dog

Bonnevie-Nielsen et al. ( 1982) Martin and Baverel (1983)

Molgolian gerbil African rodent

Mrriones

Portion of liver

Whole liver Single lobe Whole liver, PUPS

Munoz-Clares et al. ( 198I )

unguiculutus Thamnomys

Mazier et al. (1982)

garellur Ferret Wallaby Frog Tadp oIe

Kao et al. (1978) Tummur

Janssens et al. (1977)

Xenopus luevis

Wangh et al. (1979)

Runu cutesbeiuna

Kistler et al. (1975)

Pouch young

Liver minced prior to collagenase treatment

14


3.5. Preparation of hepatocytes from foetal or neonatal animals The adaptation of enzymic techniques for the preparation of isolated hepatocytes, designed for the adult animal, to the foetus and the newborn has provided valuable information on developmental aspects of metabolism. The major determinant of the method used is, of course, the size of the donor animal. In the case of larger animals, such as the newborn pig (Pegorier et al., 1982), the procedures recommended in Chapter 2 can be used with few, if any, adaptations. However, with very small animals such as the foetal rat or mouse, perfusion techniques are not technically feasible. Examination of the literature reveals that the methods most commonly used to isolate hepatocytes from foetal and neonatal rats are modifications of the in vitro digestion techniques of Devirgiliis and co-workers (Devirgiliis et al., 1981; Autuori et al., 1981). In the method of Devirgiliis and co-workers, livers were removed from foetuses or neonatal rats (< 15 days old), chopped with a blade, and then incubated in Ca2+and Mg2+-freeHanks' solution containing 0.015% collagenase, 0.005% DNase and 2% albumin. The incubation of the tissue fragments, which was carried out for 15 min in a rotating flask held at 39"C, was repeated three times, each time using fresh enzyme-containing medium. Hepatocytes were then purified by repeated centrifugation at 40 x g . The average hepatocyte yield from foetuses of 16 days gestation (term for the rat being 22 days) was reported to be 25 x lo6 cells/g liver and for 17-21-day foetuses, 50 x lo6 cells/g liver. Trypan blue exclusion for these two groups was 90% and 95%, respectively. In an earlier but quite similar method, Yeoh et al. (1979) used a 0.05% solution of collagenase without DNase to prepare cells from rat foetuses of gestational age 17 days and greater. This technique failed to produce good yields of cells from 15-16-day foetal rats. To produce cells from such animals, livers were dispersed using repeated

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passages through a pasteur pipette, without the use of collagenase. Unfortunately, actual values for yield and integrity were not reported. Fletcher et al. (1988) successfully modified Yeoh’s technique for 15day foetuses by treating tissue fragments with collagenase and then dispersing the tissue with a pasteur pipette. A similar procedure was also adopted by Germain et al. (1988), who successfully isolated epithelial cells, which are probable progenitors of hepatocytes, from livers of 12-day foetuses by using a mixture of collagenase, dispase and hyaluronidase (see also Section I 1.10.1). Other workers have incorporated an initial step, prior to incubation with collagenase, in which the sliced tissue is incubated for 5-30 min in a Ca2+-and Mg2+-freesalt solution containing 0.5 or 0.6 mM EGTA. Ca2+ (2.5-5 mM) is then added to the collagenasecontaining medium, in a manner analogous to the two-step procedure for adult rats (Section 2.3.) (DeSante et al., 1984; Gressner and Vasel, 1985; Lorenzo et al., 1986). Lorenzo et al. (1986) used a two-step technique that involved incubating liver pieces in 0.05% collagenase without DNase to prepare 15 x lo6 cells/g of foetal liver (at day 22), more than 95% of which excluded trypan blue. DeSante et al. (1984) used a similar method, but included DNase, to obtain between 30 and 50 x lo6 dye-excluding hepatocytedg from livers of foetuses of 19 or 20 days gestation. Ferre et al. (198 1 ) successfully prepared hepatocytes from rats, delivered by caesarean section immediately prior to term, by retrograde perfusion of the liver with a collagenase-containing medium. The technique has also proved satisfactory for preparing hepatocytes from newborn rabbits, or from rabbit foetuses removed 3 days prematurely (El Manoubi et al., 1983). On the other hand, at least two groups of workers (Deschenes et al., 1980; Nakamura et al., 1988) have successfully prepared hepatocytes from neonatal rats as early as 1 day after birth by perfusing the livers via the portal route. During foetal life, the liver contains large numbers of haematopoietic cells. The proportion of these, relative to other cells, decreases slowly towards term, and then rapidly following birth, to the adult value of

76


less than 1% of the liver by volume. For example at day 15 of gestation, haematopoietic cells comprise 61% of the total cell population dropping to 46% prior to birth. Because of their relatively small cellular volume, however, these cells comprise only 37% of the total volume of the liver at 15 days of gestation and 10%just before birth (Greengard et al., 1972). As a consequence of this, preparations of foetal hepatocytes purified by low speed centrifugation techniques are contaminated with much higher numbers of haematopoietic cells than if an adult rat liver is used. For example, after 3 washes, each followed by centrifugation for 5 min at 50 x 8, the final preparation of Hommes et al. (1971) contained 75'51 hepatocytes (by cell number) when foetuses were used 4 days before term, and 92% hepatocytes when animals were used the day before term. Increasing the number of washing steps will decrease contamination but at the expense of cell yield. Trypsin and lysozyme have also been used to isolate foetal hepatocytes. Hommes et al. (1971) used lysozyme to isolate foetal hepatocytes but rather poor yields were obtained. Trypsin has, in general, been avoided for the production of isolated cells because of the possibility that this enzyme may damage the cell surface. However, Dargel and co-workers (Schulze et al., 1984; Dargel et al., 1987) used trypsin to prepare foetal parenchymal cells with yields of 1 1 x IOVg liver wet weight and were able to demonstrate that these cells were responsive to a- and 0-agonists and to insulin.

3.6. Preparation of hepatocytes f r o m abnormal animals It has proved possible to prepare suspensions of intact hepatocytes not only from normal animals but also from a wide range of animals subjected to experimental manipulations. These include aged animals (van Bezooijen, 1978), animals treated with drugs such as phenobarbital (Moldeus, 1978; Berry et al., 1980) that induce cytochrome P-450-dependent monooxygenases, or clofibrate which increases perox-

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77

isomal @-oxidation of fatty acids (Yamada et al., 1986; Bergseth et al., 1986). Cells can be prepared from animals with genetic disorders, for example the obese Zucker rat (Triscari et al., 1981: McCune et al., 1981) and rats with glycogen storage disease (Clark et al., 1981; Blackmore and Exton, 1981). Hepatocytes have also been prepared from animals treated to create an endocrine disturbance such as diabetes induced by alloxan (Bollen et al., 1983; Miller et al., 1984) or steptozotocin (Golden et al., 1979). Cells can also be obtained from hypo- or hyperthyroid rats (Berry, 1974b; Werner and Berry, 1974). Rats fed abnormal diets including a ‘cafeteria diet’ (Barber et al., 1985; Berry et al., 1985), a high fat (Malewiak et al., 1985; Pegorier et al., 1988) or high alcohol diet (Berry et al., 1980; Kondrup et al., 1980), or a low protein diet (Harris et al., 1985; Ekanger et al., 1988) have also acted as donors. In addition, isolated hepatocytes can be prepared from cirrhotic livers caused by exposure of animals to phenobarbital and carbon tetrachloride (D’Arville et al., 1989) or thioacetamide (Dargel et al., 1987). In general, yields of cells from all such animals are good, although the fat-rich liver of the alcohol-fed animal gives poor yields (Berry et al., 1980). When pre-treated animals are used, the cells produced must be carefully assessed for integrity (as discussed in Chapter 4) to avoid the criticism that the experimental results are artefacts of poor cell preparation.

3.7. Preparation of suspensions of damaged hepatocytes Prior to the introduction of the use of collagenase, many attempts were made to prepare isolated hepatocyte suspensions by perfusion of the liver under pressure (Section 1.3.). Although high yields of cells were frequently obtained by this approach, the cells were invariably damaged and proved to be completely unsuitable for metabolic studies involving cytoplasmic processes (Berry, 1962). The reason for this was a loss of integrity of the plasma membrane that allowed leakage not only of small molecular weight constituents but also of cytoplasmic

78


enzymes. In effect, the techniques produced a particulate preparation from which most of the soluble constituents extraneous to the intracellular organelles (‘cytosol’, see Section 12.1) were lost. Moreover, the organelles themselves underwent dramatic changes in morphology, adopting the appearance normally seen after isolation following cell fractionation (Berry and Simpson, 1962; Berry, 1974a). Nevertheless, the preparation of cells by high-pressure perfusion of the liver should not be completely ignored, even though virtually every cell released stains with trypan blue, for it allows the rapid production of a particulate preparation that can be readily washed free of contaminating soluble components. Hence it may be regarded as a liver homogenate from which the soluble 100,000 x g supernatant fraction has been removed. The mitochondria of this preparation have been shown to behave very similarly to isolated mitochondria (Berry, 1965) and the preparation has proved a useful adjunct for mitochondrial studies. Purified cytosolic components can be recombined with the particulate system, to provide a more refined means of studying certain aspects of interactions between the cytoplasm and organelles such as the nucleus, mitochondria or endoplasmic reticulum. A simple method for the preparation of cells by high-pressure perfusion of rat liver is given in this Section. Again, it must be emphasized that the preparation is completely unsuitable for studying the properties of intact cells. 3.7.I . Reagents and apparatus

The only reagent required is a medium containing 0.25 M sucrose, buffered to pH 7.4 with 0.02 M Tris-HC1. The apparatus is also extremely simple, comprising a 50-ml syringe and a water bath, held at 37°C in which a wide-mouthed flask containing 250 ml of medium is placed. A loose-fitting glass and teflon homogenizer (Fig. 3.1) is also suspended in the bath, or alternatively and more conveniently, held in an incubator at 37°C.

Ch. 3.

OTHER METHODS

79

Fig. 3. I . Glass and teflon homogeniser.

Protocol 3.2 Preparation of damaged cells by high-pressure liver perfusion (i) Follow steps (i) to (iii) of Protocol 2.2. (ii) Fill the syringe with sucrose medium and attach it to the portal

80


cannula. Gently at first, and then more forcefully, flush the liver with the sucrose solution until the inflow rate exceeds the outflow to such an extent that the liver distends fully. Detach the syringe, refill it and repeat the procedure. At the end of the high-pressure perfusion, all lobes of the liver should be blanched and fully distended (‘Sydney Opera House effect’). (iii) Now gently tear the liver from its ligamentous attachments. A few minor cuts with scissors may be necessary to free the organ completely. Cut the excised liver into pieces small enough to be placed in the homogeniser tube. Flush them into the bottom of the tube with about 20 ml of sucrose medium. (iv) With the hand-held pestle, break up the liver. Generally twelve up-and-down strokes of the pestle are sufficient to obtain complete disintegration, provided that an adequate degree of perfusion and distension has been previously achieved. (v) Dispersion and washing are carried out at 0 4 ° C . Pour the dispersion into two ice-cold 50 ml centrifuge tubes and centrifuge at 50 x g for 3 min. Decant each supernatant and resuspend the pellets in fresh ice-cold medium. Centrifuge at 40 x g for 3 min, wash at least twice more and finally combine the pellets. The pelleted material, almost entirely comprising damaged cells, can be used as an experimental system in the same manner as other types of preparations of isolated intracellular organelles.

3.8. Preparation and purification of non-parenchymal cells A half-century-old method for preparing Kupffer cells involves loading them with colloidal iron in vivo and separating them magnetically (Rous and Beard, 1934). More recently, several methods have been described for the purification of Kupffer cells from the unwashed cell suspension obtained following collagenase digestion. One of the earlier methods (Berg and Boman, 1973) described the preparation of Kupffer cells by incubating the suspension with pronase. Pronase destroyed all parenchymal cells while leaving Kupffer cells relatively intact.

Ch. 3

OTHER METHODS

81

Some workers have preferred to avoid pronase (Bodenheimer et al., 1983)as Kupffer cells exposed to this enzyme have been shown to lack certain receptors (Crofton et al., 1978). Instead, they have used centrifugal elutriation (see Section 12.13.2) with or without loading of iron or Triton X-100 to increase Kupffer cell density (Praaning-Van Dalen and Knook, 1982). Bodenheimer et al. (1983) used a series of low speed centrifugation steps, including one involving a continuous density gradient of Percoll. The resultant suspension consisted of Kupffer cells and endothelial cells. Kupffer cells were purified further by their ability to adhere to tissue culture plates. Endothelial cells (Smedsrod et al., 1990),lipocytes (Friedman et al., 1989)and bile ductular epithelial cells (Mathis et al., 1989) have been isolated by a variety of enzymatic techniques. Most of these methods employ collagenase but contain a step in which hepatocytes are removed or destroyed.


CHAPTER 4

Assessment of integrity of isolated hepatocytes

4.1. Introduction It is often not easy for a newcomer to the field of hepatocyte isolation to decide whether the quality of a suspension of isolated cells is suitable for experimental purposes. A decision about this may be particularly difficult if the study being carried out has not been performed previously by other workers. An obvious approach is to choose a set of experimental conditions that seem likely to give the desired performance, and to examine the ability of the cells to function in this test system. If satisfactory results are obtained, much time and effort has been saved. However, if the outcome does not meet expectations, it may not be easy to determine whether it is the cells, or other features of the experimental protocol, that are at fault. For this reason, it is recommended that beginners in the field of isolated hepatocyte preparation gain confidence that their preparative technique is satisfactory by using a number of different measures of cellular integrity in their initial experiments. After it becomes apparent that high quality

84


preparations are regularly being produced, the use of trypan blue staining is likely to prove adequate for routine purposes. In the majority of papers, the term ‘viable’is used to describe isolated hepatocytes that exclude trypan blue. Thus, a preparation, in which 15% of the cells take up the dye, would be described as 85% viable. This term, with its implication of capacity to live, may give the beginner in the field a deceptive impression of the biochemical and physiological capabilities of the isolated hepatocyte. We therefore prefer to describe cells excluding trypan blue as ‘intact’. However, this term could be considered equally misleading. In truth, liver cells separated from their neighbours can hardly be expected to perform as they do in situ, and, furthermore, there is considerable evidence that the anatomy of the liver greatly influences the metabolic behaviour of the hepatocytes (reviewed by Jungermann and Katz, 1989). The presence of reticulo-endothelial cells may also have an influence on hepatocyte behaviour (Lowitt et al., 1981; McCallum, 1981; Keller et al., 1985; Fisher et al., 1989). Employment of the term ‘viable’ is now so commonplace that it is unlikely to cease. Nevertheless, the use of a simple statement of the degree of dye uptake or exclusion would be preferable. The assessment of the integrity of the hepatocyte preparation must be a largely empirical operation, depending to a considerable extent on the goals of the experimenter. Even so, experience has shown that a relatively limited number of procedures are sufficient to reveal whether or not a preparation is likely to have metabolic capabilities similar to those observed in vivo. The simplest procedure is to look at the cell suspension with the naked eye. If large clumps or strings of gelatinous material are present, it does not augur well, though on occasion, removal of this material by filtration through nylon mesh may yield a satisfactory suspension. Under the light microscope it is relatively easy with practice to distinguish intact and damaged cells even without application of a stain (Section 5.1.2). The intact cells tend to be rounded and refractile, the damaged cells, irregular, flattened and granular. For quantifying cell damage, however, more elaborate

Ch. 4.

ASSESSMENT OF INTEGRITY

85

methods are required. The most useful are those that rapidly draw the attention of the research worker to serious problems, and thus provide an opportunity for the experiment to be aborted. Three such approaches are described here. For individuals new to isolated hepatocyte preparation, additional but less immediate means of assessing integrity that supply evidence of the cells' ability to respond to ATP demand are suggested.

4.2. Measures of cellular integrity 4.2.1. Trypan blue staining

The use of trypan blue has been discussed in some detail in Section 2.5.1 and the related Protocol 2.3. This organic amine dye, M ,961, is excluded from hepatocytes with intact membranes, whereas damaged cells readily take it up. I t is not necessary for gross damage to be present, in that the plasma membranes of staining cells frequently appear intact when viewed by electron microscopy. Trypan blue is negatively charged and it seems likely that the dye is excluded as a result of an energy-dependent maintenance by the hepatocyte of a plasma membrane potential, negative inside (Claret and Mazet, 1972; Berry et a]., 1988). Loss of this potential due to cell injury may allow penetration of trypan blue. The 'supravital' stain, Janus Green B, M r 51 I , which is positively charged, also stains damaged hepatocytes on immediate exposure. However, in contrast to trypan blue, intact hepatocytes incubated with Janus Green B at 37°C over a period of about 5 min, take up the dye, the mitochondria showing the earliest signs of staining. The mode of uptake is probably analogous to the active concentration, within the mitochondria of intact cells, of other organic cations such as triphenylmethylphosphonium ion (TPMP') as a function of mitochondria1 membrane potential (see Section 12.9.2). A number of workers have stressed that trypan blue exclusion is a relatively crude measure of cellular integrity. Krebs et al. (1979a)

86


pointed out that hepatocytes, stored at 22°C for 3 h lost much of their gluconeogenicactivity, but most retained their ability to exclude trypan blue. They also demonstrated that certain surface-active agents abolished gluconeogenesis whilst having little effect on the degree of dye exclusion. In fact, many cellular poisons behave in this manner, since particular metabolic pathways may be far more susceptible to such agents than are the processes which maintain plasma membrane integrity (and potential). Hence trypan blue exclusion cannot be regarded as a firm indication that the hepatocytes are metabolically intact. However, we can say with some confidence that the converse is true. Thus, if a large percentage of the cells are perceived to take up the dye, it is a safe conclusion that the preparation is irrevocably damaged. The next question to be addressed is what constitutes a satisfactory preparation. Scrutiny of the literature indicates that most workers consider that at least 90% of the hepatocytes should be seen to be excluding trypan blue prior to the commencement of the experimental incubation. Some, however, appear content with an 80%exclusion level, but this would not be considered acceptable in our laboratory. Dye uptake of this degree is a warning sign indicating the possibility of some general damage to the hepatocyte suspension during the preparative procedures. The anabolic activities of such preparations are usually depressed proportionately more than the level of trypan blue staining would indicate, and a further substantial increase in trypan blue uptake can be anticipated during the experimental incubation. Indeed, even in a satisfactory preparation in which the degree of cell staining is less than 10% at the end of the preparative procedures, an additional 5-15”/0 of the cells may take up the dye during a 40-60-min incubation, depending on the experimental conditions. The technical problems that must be considered when using trypan blue staining as a measure of cell integrity are discussed in Section 2.5.1. 4.2.2. Succinate oxidation

Isolated liver mitochondria oxidize succinate vigorously. In fact, this

Ch. 4.


87

substrate is metabolized by hepatic mitochondrial suspensions more rapidly than any other physiological fuel. However, the oxidation is largely incomplete, the major end products being not CO,, but malate and fumarate. Mapes and Harris (1975) observed that the less the degree of damage to isolated cells, as measured by trypan blue exclusion, the lower the rate of succinate oxidation. They concluded that the integrity of isolated cell preparations could be judged on the basis of their inability to oxidize added succinate. However, some workers have erroneously used succinate oxidation as a positive indicator of metabolically active, and therefore intact, isolated cells. While there is widespread agreement that hepatocyte succinate oxidation is an indicator of cell damage, there is no evidence that lack of ability of cells to oxidize succinate is a better indicator of their integrity than is dye exclusion. The measurement of the ability of the hepatocyte suspension to oxidize succinate will not by itself give the information required to determine the degree of cell damage. In addition, it is necessary to know the rate of succinate oxidation that would occur were all the cells to be damaged. This can be achieved by measuring the rate of succinate oxidation in a small sample of cells that have been exposed to digitonin. (For a discussion on the mode of action of digitonin on cell membranes, see Section 12.3.1). Our data indicates that if the concentration of digitonin is increased above the minimum level required to cause all of the cells to stain with trypan blue, there is a decrease in the rate of succinate oxidation. This may be due to digitonin damage to the mitochondrial membranes. Thus, when either the digitonin added is insufficient to make all of the cells permeable to succinate, or is excessive and causes mitochondrial damage, there will be an underestimation of the rate of succinate oxidation that should occur when all the cells are damaged. This in turn will result in an overestimation of the percentage of cells permeable to succinate. Mapes and Harris (1975) observed a small unexplained discrepancy between the percentage of cells staining with trypan blue and the apparent percentage of cells permeable to succinate, the latter being greater. Likewise we have ex-

nn


perienced a difference between estimates of cell damage, determined by lactate dehydrogenase (LDH) leakage and permeability to succinate, the latter again being greater. It is possible that failure to use the optimal concentration of digitonin may account for these discrepancies. We have also observed that the amount of digitonin required to cause all cells to stain with the dye increases with the length of time that the digitonin solution has been stored. Hence, for reproducibility, the digitonin solution should be titrated against each cell preparation to obtain an accurate estimate of the maximum level of succinate oxidation over the titration range. Zuurendonk et al. (1979) recommend that a fresh solution be made every 3 h, but even with this precaution, it may prove necessary to use different concentrations of digitonin to lyse completely cells sampled at different times during a prolonged incubation. It is recommended therefore that, to avoid the errors introduced by the use of too little or too much digitonin, monitoring of the suspension for the degree of staining with trypan blue be carried out on a routine basis during the digitonin titration procedure. Krebs et al. (1979a) suggest that the combination of succinate and iodonitrotetrazolium is a useful marker since in damaged cells mitochondria1 succinate oxidation generates a red formazan dye within a few minutes. This approach has the advantage of obviating the need for an oxygraph or digitonin permeabilization of the cells. In any case, the results appear virtually identical to those obtained with trypan blue. 4.2.3. Equipment and solutions for measuring succinate oxidation

An 0,-electrode, monitor and incubation chamber capable of holding at least 4 ml of medium are required for the measurement of succinatestimulated respiration, but this apparatus is likely to be available in any laboratory equipped for undertaking metabolic studies. If no system is available, Yellow Springs Instruments (see Section 6.5.1) offer a relatively inexpensive apparatus. Other requirements are a recirculating constant temperature water supply and a cylinder of carbogen.

Ch. 4.


89

The incubation medium can be identical with that used for standard cell incubations, e.g. Krebs bicarbonate-saline (Krebs and Henseleit, 1932). Because the K,,, for succinate is relatively high, a succinate concentration 1 10 mM is required to achieve maximal rates of respiration. Hence a concentrated substrate solution of 0.4 M is necessary. Prepare a 2% (w/v) digitonin solution by carefully heating the digitonin in bicarbonate-free phosphate saline (Protocol 2.1) in a hot water bath until it dissolves. The reagent is stored on ice.

Protocol 4.1 Determination of proportion of cells oxidizing succinate (i) Add 2.5 ml of bicarbonate-saline to the incubation vessel and gas it with carbogen while stirring with a magnetic bar. Calibrate the monitoring equipment against the carbogen saturated medium. Add 1 ml of 9"h BSA and 0.4 ml of cell suspension, 30 mg dry weight/ml, to the incubation vessel. (ii) Monitor the 0,-consumption until a linear rate of respiration has been achieved. At this time add 0.1 ml of 0.4 M succinate, and again measure the rate of 0,-consumption. The rate of succinatestimulated oxidation by the hepatocyte preparation is calculated from the difference between these two rates. (iii) Titrate a sample of cells (0.5 ml) with digitonin solution in a step-wise manner, 5 pl at each addition, keeping both the suspension and solution ice-cooled. Monitor the effects of the digitonin by testing the cells for dye-exclusion after each addition of digitonin. Determine the lowest concentration of digitonin that results in virtually all cells taking up trypan blue, Only about 20 pl will be required. (iv) Take another cell sample (0.95 ml) and mix well with sufficient digitonin solution to give a concentration in the mixture equal to that previously observed to bring about total trypan blue staining. In our experience an appropriate final digitonin concentration is about 0.08%, but this may vary according to the quality of the digitonin.


90

(v) Add 0.4ml of the digitoninized cell suspension to an incubation chamber containing 2.5 ml of bicarbonate-saline, that has been gassed with carbogen, and 1 ml of BSA. Measure the rate of 0,uptake. Add 0.1 ml of 0.4 M succinate and measure the new rate of 02-consumption. Calculate the maximum rate of succinate-stimulated oxidation by the digitonin-treated cell preparation as in (ii). (vi) Calculate the percentage of damaged cells on the basis of the following equation: % damaged cells =

rate of succinate-stimulated oxidation by untreated hepatocyte preparation x 100

maximum rate of succinate-stimulated oxidation by digitonin-treated preparation 4.2.4. Enzyme leakage from isolated hepatocytes

Surprisingly little is known about the mechanisms whereby intracellular enzymes pass into the extracellular medium, even though the importance of this phenomenon in clinical diagnosis has encouraged much research in the area. The main conclusions from these investigations are that cytoplasmic enzymes escape much faster than do mitochondrial ones, and that membrane-bound enzymes are hardly released at all. As might be expected the amount of enzyme discharged is a function of its concentration within the cell, while the larger the molecule, the slower its release (Schmidt and Schmidt, 1967). For quantitative assessment of the degree of plasma membrane damage, the cells must be lysed to release all cytoplasmic enzyme activity. If the enzyme under study is confined entirely to the cytoplasm, Triton X-100 is a convenient disrupting agent. If, however, the enzyme is located in part within the mitochondria (or other organelles), Triton X- 100 is unsatisfactory, since it will bring about the release of enzyme from the mitochondria1 and other compartments also.

Ch.4.


91

LDH (EC 1.1.1.28) is an enzyme located almost entirely in the cytoplasm. Its use as a measure of cell integrity has some advantages over the trypan blue staining technique. In contrast to the situation when assessing the degree of trypan blue staining (Section 2.5.1), it does not matter when measuring LDH leakage if cell lysis has occurred, since an accurate estimate of cell damage can be obtained even if some of the damaged cells have disintegrated. Moreover, the statistical problems with the trypan blue technique, arising from the very small number of cells examined, are avoided. It is of practical importance that, in contrast to trypan blue exclusion measurements, which are tedious, time consuming and must be completed within 30 min of sampling, samples for LDH measurement may be set aside and accumulated until a more convenient time for their assay. Cell samples to be used for the estimation of total LDH activity should be lysed within 24 h to avoid loss of activity during storage. The LDH in diluted supernatant obtained from this lysate, or from samples of cell-free incubation medium, is stable during storage for at least 1 week at -10°C. It will be apparent that during the preparative procedures associated with obtaining a suspension of isolated hepatocytes, the cells undergo several washes, during which the LDH released from damaged cells is likely to be lost (Jauregui et al., 1981). Conversely, a substantial amount of LDH will be released from any cells completely disrupted following the final cell wash. It follows that the measurement of LDH cannot be used to ascertain accurately the degree of cell damage in the initial suspension. Rather, the technique is best suited for estimation of cellular damage occurring at measured time intervals over a set incubation period. Generally, the level of LDH leakage under these circumstances appears to correlate well with the degree of trypan blue staining (Kreamer et al., 1986) and succinate oxidation (Mapes and Harris, 1975).There are exceptions as described in Chapter 12, where treatment with agents such as filipin, saponin or digitonin, or electroporation can be used to dissociate the relationship between LDH leakage and trypan blue staining.


92

LDH catalyses the reversible reduction of pyruvate to lactate, the last step of anaerobic glycolysis. L-lactate

+ NAD'

L-LDH

-pyruvate

+ NADH + H+

The quantity of enzyme in a solution may be estimated by measuring the decrease in optical absorbance at 340 nm when NADH is oxidized to NAD'. Details of the assay are given in Bergmeyer (1983). To estimate the percentage of LDH leakage from cell suspensions the following procedure is followed.

Protocol 4.2 Measurement of enzyme leakage (i) Centrifuge a sample (0.2 ml) of a cell suspension (15 mg dry weight/ml) in a microcentrifuge (see Section 6.2.2) at 12,000 X g for 20 s at room temperature. (ii) Transfer a portion of the supernatant (approximately 0.1 ml) to a clean tube. When convenient, dilute it 1:20 with 0.1 M TrisHCI buffer (pH 7.4), if LDH leakage is estimated to be I 60% or dilute 1:50 for enzyme leakage > 60% Keep at CL4"C. Assay for LDH and calculate its total activity in the supernatant. (iii) To measure the total LDH present in the incubated hepatocyte suspension, transfer 0.2 ml of a thoroughly mixed portion of the suspension to a microcentrifuge tube. Add an equal volume of Triton X-100 (I%), v/v) to lyse the cells and mix well several times during the next 10 min. Keep at 0 4 ° C . Centrifuge the mixture at 12,000 x g for 20 s, transfer a portion of the resultant supernatant to a clean tube and dilute 150, as soon as convenient, with 0.1 M Tris-HCI. Assay for LDH and calculate the total activity of the hepatocyte extract. Stored samples of diluted supernatant or cell extract should be kept at -10°C and assayed within a week.

Ch. 4.


93

(iv) The percentage LDH leakage is calculated as follows: YOLDH leakage =

activity of supernatant activity of cell extract

x 100

Alanine aminotransferase (EC 2.6.1.2) leakage may also be used as a measure of cellular plasma membrane damage. However, in this case the enzyme is located in the mitochondria as well as the cytoplasm of the cell. When determining the percentage of damaged cells, the plasma membrane alone, and not the mitochondria1 membrane must be damaged by careful digitonin treatment, to release only the enzyme located in the cytoplasm, (cf. succinate oxidation, Section 4.2.2). 4.2.5. Choice of a method for determining cellulur integrity

In view of the relatively imprecise nature of all the above approaches to determining cellular damage, it is encouraging that such a good correlation between them is usually found. It might be anticipated, therefore, that any of the techniques, described in this Chapter, would be equally suitable for assessing cell integrity. In fact, each technique has certain advantages and disadvantages that limit its usefulness and determine under what circumstances it is optimally employed. Determination of trypan blue exclusion is the method of choice for assessing the quality of the initial suspension (see Section 2.5.1). I t is without doubt the most rapid means of identifying damaged cells and, as a first approximation, the proportion damaged can be readily ascertained by counting the number of stained cells and the total number in the field. However, error may result from a number of factors, such as differences in technique between different operators, uneven cell distribution over the slide, statistically small samples and albumin binding of the dye, which may give misleadingly low values for percentages of cells staining with trypan blue. Phillips (1973) showed that the inclusion of foetal calf serum (S%,v/v) in the medium

94


can lower the proportion of cells staining with trypan blue by 50'%). Since increasing the dye concentration is not practical because the serum takes up stain to such an extent that the cells are no longer visi(v/v) foetal ble, he suggests using the stain erythrosin B. Up to 10'%1 calf serum in solution (equivalent to about 7 g/I protein) had no effect on the percentage of cells staining with erythrosin B. When trypan blue is used to determine the degree of cellular damage in numerous samples, difficulties also arise due to the relatively short time available to count the cells before they deteriorate. In addition, during long incubations, particularly in the presence of a noxious agent, there is potential for disintegration of damaged cells, which must lead to an underestimation of the degree of damage by the dye-exclusion method. As explained previously, measurement of LDH activity can provide no insight into the quality of the initial suspension, but it is the method of choice for monitoring damage during long-term incubations. In theory, the measurement of succinate oxidase activity should be useful for assessing cellular damage both in the initial suspension and during long-term incubations. In practice measurement of succinate oxidase activity, which is considerably more complex than measurement of trypan blue uptake, does not appear to offer any advantages over the dye-exclusion method. Its value in the assessment of damage during long-term incubations is unfortunately limited by the time required to perform multiple analyses with the 0,-electrode. In summary, trypan blue uptake is the most rapid method for identifying cell damage during the preparative procedures and in the initial suspension. Measurement of LDH activity is the method of choice for assessing damage during long-term incubations. Under appropriate experimental conditions, all three methods give virtually identical results. Nevertheless, it is desirable on occasion to confirm the results from one type of test with those obtained with a different method.

4.3. Additional methods for determining cellular integrity The methods for assessing cell damage, described in the previous Sec-

Ch. 4.


95

tions, do not yield any direct information in regard to cell function. They are not capable of providing the research worker with insight as to whether or not the preparative techniques being used are likely to be satisfactory for the desired experimental studies. They do, however, provide useful information on the presence and degree of significant cell injury. Once the worker is confident that the preparative techniques do not induce serious cell injury, as measured by one of these methods, an evaluation must be made of the suitability of the hepatocyte suspension for the task the research worker has in mind. If the property to be studied has not been examined previously in isolated hepatocyte suspensions, the definition of optimal incubation conditions may not be easy. In such circumstances, it is probably desirable to ascertain that the total incubation system (apparatus and cells) is functioning well, before proceeding to the specific problem under investigation.

4.3.1. Functional measures of cellular integrity For cells from fasted rats, the simplest test of function is probably gluconeogenic capability, whereas for cells from fed animals the rate of urea synthesis is a useful indicator of the cells’ capacity for ATP synthesis. Further details about these major metabolic pathways are provided in Chapter 6. Another useful indicator of cell quality is ATP content (Table 6.3). ATP breakdown to ADP occurs rapidly in cells exposed to hypoxia or to inhibitors of respiration (Cornell, 1983; Baur et al., 1975). The conversion of ADP to AMP is then catalysed by adenylate kinase, and AMP in turn is broken down, leading to a fall in total adenine nucleotides (van den Berghe et al., 1989). Following recovery from hypoxic insult, cellular ATP levels, as well as total adenine nucleotides, will be perhaps only 60-80% of normal, although ATPIADP ratios may be within normal limits. The response to glucagon of hepatocytes from fed or fasted animals is a delicate measure of cellular integrity. Since the peptide hormones exert their effects through receptor sites on the plasma membrane, it

96


can be expected that damage to this membrane may diminish glucagon sensitivity. A good cell preparation should respond, by increased glycogenolysis (fed) or enhanced gluconeogenesis (fasted), to levels of glucagon as low as 10-*-10-9M (Garrison and Haynes, 1973; Seglen 1973). Further details of responses to hormones are described in Chapter 8. Other authors have used rates of protein or glycogen synthesis as measures of cellular integrity. These, however, are more difficult to measure and therefore are best used as indicators of damage caused by specific toxic agents (see Chapter 7). 4.3.2. Other measures of cellular integrity

Several groups (e.g. East et al., 1973; Sturdee et al.,1983; Krebs et al., 1979a) have suggested that respiratory rate may serve as a measure of cell quality. If cellular respiration is significantly below the levels reported by other workers, this is a useful indicator that a deficit in cell quality may exist. Generally, however, such a problem would be accompanied by a marked increase in the proportion of cells staining with trypan blue. The possibility also exists that the respiratory rate may be increased as a consequence of uncoupling of mitochondria1 respiration, which might occur, for example, as a result of exposure of cells to traces of detergent. Even so, the rate of respiration can be considered a useful measure of cellular integrity, and if facilities are available, beginners in the field are encouraged to determine the respiratory behaviour of their cell preparations. An important cellular function is the maintenance of transmembranous ion gradients. Hence it can be anticipated that measurement of these gradients should be a useful indicator of cellular integrity. That this conclusion is correct has been shown by various studies on hepatocytes depleted of Ca2+(Barnabei et al., 1974; Kolb and Adam, 1976; Thomas and Reed, 1988a,b). Ca2+-depletedcells are unable to attain or even maintain normal gradients of Na+ and K' across the plasma membrane, and the ability of such cells to regain normal

Ch. 4.


91

cation distribution is irreversibly lost when Ca2+-depletionpersists for 2 h (Kolb and Adam, 1976). The importance of this phenomenon in relation to hepatocyte preparation is discussed in Section 2.3.2, and further details are given in Section 6.3.1. Whilst in theory, measurement of Na+ and K+ gradients across the cell membrane should be a valuable tool, there are a number of factors which limit its usefulness. In the first instance, the maintenance of ion gradients requires a continuous energy flow. Since many of the manipulations during cell preparation, such as breaking up the liver or centrifuging the cells, inevitably must lead to some degree of hypoxia, the normal gradients are at least partially lost during the preparative procedures, so that the hepatocytes require a period of aerobic incubation at 37"C, before normal gradients are regained. Reversible loss of gradients can also be caused by storage at low temperatures. Secondly, the measurements require extensive manipulation of the cells (see Protocols 6.3 and 6.4), and the use of 'H,O and [14C]inulinto determine extracellular and intracellular water. Thus, the measurement of ion gradients may well be a useful endeavour for the beginner wishing to characterize the cell preparation, but it is not recommended for routine assessment of cell integrity.

4.4. Assessment of cellular integrity following cell incubations Attention has already been drawn (Section 4.2.1) to the cell damage which inevitably occurs during experimental incubations of hepatocytes. In an incubation medium free of toxic agents about 10% of cells that are apparently undamaged while held at 0 4 ° C take up trypan blue on subsequent incubation at 37°C. Most of the damage usually occurs within the first 10 min of incubation. Over the next 3 h, cell integrity tends to be maintained. If potent respiratory inhibitors are present, considerably more damage may ensue, particularly in the case of cells derived from fasting animals. It is desirable, therefore, to sample the hepatocytes at the end of the incubation period to deter-

98


mine the degree of damage. The relative merits of trypan blue exclusion and LDH activity for this purpose have already been discussed in Section 4.2.5. Kreamer et al. (1986) report that preparations in which damaged cells are removed by treatment with Percoll maintain high levels of integrity and dye-exclusion during incubations lasting many hours. It seems likely, therefore, that the susceptible cells liable to take up trypan blue during the first 10 min of the incubation are removed by prior treatment of the preparation with Percoll (see Section 2.8). There are many agents which in low concentrations cause specific biochemical disturbance, but when present in too high a dose bring about general non-specific damage, reflected by cellular LDH leakage or trypan blue uptake. Scrutiny of cellular integrity following incubation is the best means of ensuring that the concentration of the agent under study is optimal. Of course, the metabolic activities of the cells during the experimental incubation will also provide a good indication of their integrity. Useful parameters include levels of ATP, ATP/ADP ratios and synthetic rates for the processes under study. Note however that certain substrates such as fructose and glycerol can deplete ATP, but otherwise appear to have no effect on cell integrity. It is important to include in the experimental design time-courses that will reveal any decrease in metabolic rates during the period of study and thus may provide an indication of progressive cell damage.

CHAPTER 5

Microscopy of isolated hepatocytes

5.I . Light microscopy of isolated hepatocytes 5.1.1. Appearance of cells during and immediately after isolation

During perfusion of the liver with a medium containing collagenase, the hepatic sinusoids become distended so that the hepatocytes appear as naked cords, maintaining their contiguity with adjacent cells only at their pericanalicular junctions. Those portions of the cells bulging into the sinusoid tend to adopt a rounded appearance, but the hepatocytes generally retain the polygonal shape observed in vivo (Berry and Friend, 1969). However, by the end of the preparative procedures, the majority of cells have adopted an oval or spherical shape. If held standing on ice, some of the cells develop blebbing of the plasma membrane (see below). Irreversibly damaged cells rapidly take up trypan blue, the staining of the nucleus being particularly intense (Fig. 5.1). Most of the cells are single units, but not infrequently a pair of cells may remain associated to form a doublet. Occasional triplets are also encountered. Size distribution has been examined in detail by Bernaert et al. (1979) and Otto et al. (1983) who found that a majority of

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Fig. 5. I . Isolated hepatocyte suspension stained with trypan blue. Some of the cells show small blebs (-). A doublet (d) is present. Light microscopy. (Bar 100 pm, mag. x205).

hepatocytes from the livers of fed rats range in size from I8 to 24 pm in diameter, with a mean of 21 pm. Cells from fasted rats have significantly smaller diameters - a mean, approximately, of 17 pm, as determined by Otto et al. (1983). 5.1.2. Features of intact and damaged cells

Features of the cells can be seen by transmitted light microscopy but are best revealed by interference contrast microscopy. Undamaged hepatocytes are generally oval or spherical in shape and show obvious brightness contrast between the nucleus and cytoplasm, together with a highly refractile sharp perimeter at the cell membrane (Fig. 5.2a,e). The appearance of damaged cells depends on whether or not the damage is reversible. Mild reversible damage is frequently observed in hepatocytes held at ice-cold temperatures or subjected to hypoxia.

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Fig. 5.2. (a), (c), transmitted light microscopy; (b), (d), (e), interference contrast microscopy. (a) Oval-shaped intact hepatocyte. (Bar 10 pm, mag. X 1400). (b) Hepatocyte with single bleb. (Bar 10 pm, mag. x 1000). (c) Hepatocyte showing general swelling. (Bar 10 pm, mag. x 1400). (d) Multiple blebs induced by treatment with menadione. (Bar 10 pm, mag. x 1000). ( e ) One irreversibly damaged and two intact spherical hepatocytes. (Bar 10 pm, mag. x 1000).

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The characteristic feature is a blebbing or blistering of the plasma membrane (Fig. 5.2b). Often more than one bleb will appear on each cell. The blebs, which are a consequence of fluid uptake by the hypoxic or anoxic cell, are translucent and typically free of organelles. Large blebs may be formed from the coalescence of smaller blebs (Herman et al., 1988). Subsequent incubation under aerobic conditions at 37°C results in a rapid reversal of bleb formation. It is inferred that these fluid-tilled blebs lack a cytoskeletal framework, as isolated mitochondria occasionally seen in such blebs exhibit distinct Brownian movement. It is possible to distinguish clearly individual blebs, to assess their spatial distribution on the cell surface, e.g. by changing focus, and to follow their individual change in size. Cooled hepatocytes may also undergo a more general swelling (Fig. 5. lc) that also is reversed by aerobic incubation. Another form of blebbing occurs in response to certain drugs, particularly those which induce redox cycling (Kappus and Sies, 1981) and deplete the hepatocyte of glutathione. This type of bleb formation is thought to be caused by disturbances in thiol and Ca2+ homeostasis that lead to disorder of the cytoskeleton (Jewel1 et al., 1982; Orrenius et al., 1987;Thor et al., 1988; Thomas and Reed, 1989). Cells incubated with the noxious agent become covered with small blebs (Fig. 5.2d). This type of bleb formation is not reversible, although during the incubation the blebs detach from the cell surface membrane, which reseals. Irreversible damage to the cell is associated with discontinuities of the plasma membrane (Herman et al., 1988). By interference microscopy, damaged cells can readily be distinguished from intact cells. Damaged cells show a highly granular cytoplasm and an illdefined irregular granular boundary with the surrounding medium whereas the intact cells have a well-rounded appearance (Fig. 5.2e). There is a loss of contrast between the cell and medium and also a marked reduction in brightness contrast between the nucleus and surrounding cytoplasmic organelles in the damaged cells.

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5.2. Morphology of isolated hepatocytes by transmission electron microscopy 5.2.1. Introduction

The biochemical behaviour of the isolated liver cell suspension resembles in so many ways the behaviour of the intact organ that there is a natural tendency to regard the isolated hepatocyte as being identical to the corresponding cell in situ. In fact important structural differences are revealed, particularly by electron microscopy. Since these changes have not been systematically set out elsewhere, they are described in some detail here.

5.2.2. Features common to both hepatocytes in situ and isolated hepatocytes

Fine structural features, common to both the hepatocyte in the intact liver and the isolated cell, include the presence of one or two central nuclei with numerous nuclear pores, abundant rough-surfaced endoplasmic reticulum, a moderate amount of smooth-surfaced endoplasmic reticulum and plentiful mitochondria, peroxisomes, lysosomes and glycogen particles (Fig. 5.3). Following glutaraldehyde fixation, the appearance of the hepatocyte is virtually indistinguishable between the two states, although slight vacuolization of the Golgi apparatus is more common in the freshly isolated cell (Berry and Friend, 1969). In both types of preparation, the actin-based cytoskeleton appears concentrated around the tight junctions of the bile canaliculi (Montesano et al., 1975; Wanson et al., 1977; Hubbard and Ma, 1983). This relationship lasts only as long as the junctional complexes are in their native position.

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Fig. 5.3. Hepatocytes removed for fixation during collagenase perfusion. The cell is loosened from its neighbours, but its junctions are probably still intact. Notice the integrity of the nucleus, rough surfaced endoplasmic reticulum, glycogen, mitochondria and plasma membrane. (Bar 5 pm, mag. ~3750).

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5.2.3. Differences between the hepatocyte of the intact liver and the isolated cell In the intact liver the hepatocyte has a clearly polarized architecture (Berry and Friend, 1969; Groothius et al., 1981; Roman and Hubbard, 1983; Hubbard et al., 1985; Simons and Fuller, 1985; Chapman and Eddy, 1989). Bile canaliculi constitute apical surfaces which have wellformed microvilli, tight, intermediate and gap junctions, and desmosomes clearly delineating their boundaries. The other facets of the polygonal hepatocyte abut adjacent hepatocytes and the resident cells of the mononuclear phagocyte system, the Kupffer cells. The bile canalicular and the polygonal surfaces facing the space of Disse have distinctive cytochemical, immunological and freeze-fracture characteristics (Roman and Hubbard, 1983; Hubbard et al., 1985; Maurice et al., 1985; Stevenson et al., 1986; Chapman and Eddy, 1989). There is also a subjectively perceived polarity attributable to the maintenance of this architecture by the location of the Golgi apparatus relative to the bile canaliculi. Features indicative of polarity are altered in the isolated cell. The majority of cells that maintain their structural integrity on separation, tear off, from contiguous cells, blebs of cytoplasm adjacent to the tight and gap junctions (Berry and Friend, 1969) (Figs. 5.4,5.5a,b). Filamentous actin seems condensed in cells near these junctional complexes. Gap junctions are not effectively open to the extracellular environment. All elements of the junctional complex (the tight junction, the intermediate junction, gap junction and desmosome) are endocytosed within an hour and appear in the walls and contents of endosomes and secondary lysosomes. At this time, the cytoskeletal network within the hepatocyte is no longer concentrated in paracanalicular areas. Indeed, it seems to unravel and become dispersed near the lysosomes where the junctions are dismantled. With this transformation, the Golgi apparatus appear more randomly distributed throughout the cytoplasm. The endocytosis and digestion of junctional complexes is accom-

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Fig. 5.4. At the completion of collagenase perfusion, cell junctions have been cleaved and some endocytosed. A few cells show common signs of damage. The hepatocyte in the centre of this micrograph has a lipid droplet in the nucleus and swelling of vacuoles, perhaps part of the endosomal system. Otherwise, all organelles are intact and display their usual in situ morphology. (Bar 5 fim. mag. x 3750).

Ch. 5.

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Fig. 5.5. (a) A small portion of cytoplasm from a formerly contiguous cell is still adherent via a cell junction, probably a gap junction in this instance, to an otherwise fully isolated cell. (Bar 1 pm, mag. x 35.000). (b) Cell junctions are incorporated into endocytic vacuoles. Later the vacuoles will be joined by primary lysosomes to be degraded within the secondary lysosomes in the hepatocyte. (Bar 0.5 pm, mag. x 54,000). (c) After junctions are endocytosed, the entire hepatocyte surface, be it bile canulicular or the portion lining the space of Disse, has a fairly uniform smooth appearance with occasional, short microvilli. Cells without junctional complexes and a uniform surface retain their morphology and constitutive functions, but show no morphologic evidence of polarized cell activity such as bile secretion. (Bar I p n , mag. X 16,500).

panied by a loss of surface polarity. Plasmalemma markers distinguishing the bile canalicular surface disperse into the remainder of the membrane. The plasma membrane confining this now spherical cell has scattered short microvilli, no obvious cytoskeletal framework

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beneath it, and dispersion of the lectin, immunologic and enzymatic markers that formerly identified the different domains of the plasmalemma (Fig. 5.5~).This loss of domain structure in the plasma membrane occurs concomitantly with the loss of regulated secretory function by the hepatocyte. For example, there is no longer packaging of synthetic products such as albumin, bile and lipoproteins for targeted transport. Lipoproteins, present in the liver cell before its isolation, persist in small vesicles of trans-Golgi reticulum, and in other parts of the smooth-surfaced endoplasmic reticulum (Figs. 5.6, 5.7b). While constitutive secretion may still occur, vectorial transport to surface domains seems disrupted (Simons and Fuller, 1985). The isolated cell is still metabolically active, maintaining its content of mitochondria, vesicles, endoplasmic reticulum, Golgi apparatus, peroxisomes and nucleus, all with apparently full integrity (Figs. 5.6, 5.7). When these hepatocytes are cultured as a monolayer (Chapter 1 I), cell junctions re-establish and the polarities visible in the native cell, including the disposition of its cytoskeleton, can return (Wanson et al., 1977; Marceau et al., 1986a; Meredith, 1988). 5.2.4. Signs of injury

The morphological indicators of cell injury are the same for cells in the intact liver and in suspension. In disrupted cells the integrity of the plasma membrane is destroyed if the tear in the region of the junctional complex is greater than 1 pm. Under these circumstances, mitochondria quickly condense (Fig. 5.8a). In less damaged cells that are capable of recovery, some mitochondria may swell (Fig. 5.6), accumulate proteinaceous droplets, and subsequently develop calcium hydroxyapatite crystals. In irreversibly injured hepatocytes, the endoplasmic reticulum and Golgi apparatus swell, large fluid-filled vacuoles appear in the cytoplasm, glycogen becomes depleted, and the cell in toto loses its former aesthetically pleasing structural organization (Fig. 5.8a). Virtually all cells prepared by mechanical techniques show the damaged appearance observed in Fig. 5.8a.

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Fig. 5.6. As seen at higher mag., fully isolated cells retain all the morphologic features of an hepatocyte in situ. The normal configuration of the rough surfaced endoplasmic reticulum ( R E R ) , smooth surfaced endoplasmic reticulum (SER), clusters of glycogen ( G ) ,a lipoprotein particle (-) and mitochondria are readily recognizcd.One mitochondrion is swollen (S), a common feature in early anoxia and preceding physiologic turnover (autophagocytosis) of organelles. (Bar 0.5 pm, mag. ~ 2 7 , 0 0 0 ) .

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Fig. 5.7. Electronmicrographs of organelles in isolated hepatocytes. (a) A mitochondrion sur. Golgi apparatus with retenrounded by clusters of glycogen. (Bar 0.5 pm, mag. ~ 4 0 . 0 0 0 )(b) tion of lipoprotein particles (-), a coated vesicle (CV) and a tubular crystal in a peroxisome (P). (Bar 0.5 pm, mag. x 26,000). (c) The nucleus and the nuclear pores also retain their normal morphology. (Bar 0.5 pm, mag. ~ 2 1 , 0 0 0 ) .

Ch. 5.

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Fig. 5.8. (a) Irreversibly damaged hepatocyte showing asymmetric swelling of the outer mitochondrial compartment. Condensation of the mitochondria1 matrix and vesiculation of the endoplasmic reticulum. (Bar I pm, mag. x 18,000). (b) Portion of an isolated hepatocyte exposed to pentachlorophenol. The cell is extruding large blebs of endoplasmic reticulum-enriched cytoplasm. Compare Figs. 5.2d and 5.10. (Bar 2 p n . mag. x 6000).

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5.3.Techniques This Section deals with techniques that have been found optimal for transmission electron miscrscopy of the isolated hepatocyte suspension. Basic electron microscopy techniques common to all tissues are not covered. 5.3.1. Fixation

The foregoing description of the hepatocyte is based on electron microscopic examination of aldehyde and osmium fixed, plastic embedded, thin-sectioned material. All general principles of good fixation apply to the isolated cell. In our original publication (Berry and Friend, 1969), isolated cells were rapidly fixed at &4"C for 90 min in 0.1 M Sorenson's phosphate buffered 1% osmium tetroxide (pH 7.4). This procedure preserved the glycogen well (Figs. 5.3, 5.7a). In several experiments, the tissue was also prefixed for 15 min in a lo/" paraformaldehyde, 3% glutaraldehyde mixture buffered with 0.7 M sodium cacodylate (pH 7.4). While these fixation procedures remain excellent, contemporary technologies have led to modification of the fixation procedure for thin sections and other types of ancillary studies. 5.3.2. Current procedure for thin-sectioned material

Fixation is initiated by the immersion of 0.1 ml of the hepatocyte suspension (106 cells) in 0.1 M sodium cacodylate buffer containing 1.5% glutaraldehyde for 1-2 h at room temperature. The cells are then washed in 0.1 M sodium cacodylate buffer (pH 7.4) containing 7%)sucrose, followed by fixation for 2 h at 0 4 ° C in Veronal-acetatebuffered 1% OsO, (pH 7.4) containing 5% sucrose. The hepatocytes are then treated for 1 h at room temperature with 0.5% uranyl acetate containing 470sucrose, quickly dehydrated in a graded series of ethanol solutions, embedded in Epon, and cut with diamond knives (PorterBlum MT-2 microtome). Sections stained with alkaline lead and 5%

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uranyl acetate are examined in a transmission electron microscope at 80 kV (JEOL lOOCX). Cells prepared with this fixation protocol are illustrated in Fig. 5.8b. They are derived from an experiment where cells were intentionally injured with pentachlorophenol (Berry and Friend, unpublished observations; see also Fig. 5.2d). For even greater enhancement of membranes, cells may be immersed in 1 % buffered tannic acid for 30 min following the OsO, fixation. 5.3.3. Ultrathin cryosections for immuno- and leetin-labelling

For cryo-immunoelectronmicroscopy, cells are fixed in 8% paraformaldehyde in 0.1 M phosphate buffer (pH 7.4) for 3 4 h at room temperature, impregnated with 20% polyvinylpyrrolidone (average M , 10,000) in 2.0 M sucrose overnight at M " C , then frozen in liquid nitrogen and stored until taken for ultrathin cryo-sectioning (Reichert FC-4 Ultracut microtome). Sections are picked up on carboncoated copper grids for immunolabelling as follows: Grids are incubated in 2% fish gelatin in phosphate buffered saline (PBS, see Section 11.12) for 10 min, washed in PBS 3 x 2 min each, and the grids transferred to drops of a specific primary antibody, usually an IgG. A range of dilutions of antibody is made (l:lO, 1:50, 1:100, 1:500) in PBS supplemented with 0.5 M NaCl (pH 7.4) and each grid is exposed to a single dilution for 60 min. Grids are then rinsed with PBWO.5 M NaCl and then with 1% fish gelatin plus 0.1% bovine serum albumin (BSA) dissolved in PBS (pH 8.0). Following reaction with a secondary antibody, usually a 10 nm gold-labelled mouse antibody suspended in PBS, containing 0.1% BSA (pH 8.0), tissues are postfixed in 1.S% glutaraldehyde in 0.1 M sodium cacodylate buffer containing 1% sucrose (pH 7.4) for 10 min. After rinsing in double distilled water (2 x 5 min), the cells are stained with 2% uranyl acetate (pH 7.4) for 5 min, rinsed again, and counterstained with a mixture of 1 ml of 2% methyl cellulose and 0.1 ml of 3% uranyl acetate (2 x 5 min on ice), then dried overnight, and

I I4


viewed in an electron microscope. The outlined prototype procedure is highly suitable for localizing antigenic determinants in the isolated cell. While some soluble components are lost, membranes are generally well-preserved and membrane-associated antigenic determinants remain confined to their native position. Labelling of single or oligosaccharide moieties on cell surfaces, for example in membrane-associated glyco-conjugates, is undertaken in an essentially analogous manner. Following cryosectioning, grids are exposed in a two-step labelling procedure, first to one of a panel of biotinylated lectins, using an appropriate range of dilutions, and then to an avidin-labelled marker. This latter marker can either be a directly visualizable label - e.g. gold or ferritin, or else may be labelled with an enzyme (e.g. glucose oxidase, peroxidase, etc.), whose location will be revealed by subsequent incubation with appropriate substrate; post fixation and staining may then be carried out as for immuno-labelling (Horisberger, 1984, 1985). Lectin binding sites on isolated liver cell plasmalemma are well revealed by this approach. Alternatively, a onestep labelled lectin procedure (gold, ferritin or peroxidase etc.) can be used. The two-step procedure has a greater sensitivity, due to the amplification available in the biotin-avidin step. 5.3.4. Freeze-fracture

To see large expanses of the hydrophobic interior of the plasma and internal membranes, freeze-fracture serves as a valuable tool in examining isolated liver cells (Montesano et al., 1975; Friend, 1982; Pinto da Silva, 1987; Severs, 1989). In freeze-fracture membrane replicas, in general, proteins appear as 8-nm particles, and the smooth intervening domains represent cholesterol and acid tails of phospholipids. With this technique, we have confirmed the endocytosis and degradation of tight and gap junctions and desmosomes (unpublished observations) previously seen in thin-sectioned material. For freeze-fracture, cells are fixed in 1.5%glutaraldehyde buffered at pH 7.4 with 0.1 M sodium

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cacodylate at room temperature. After 1 h, the fixed specimens are immersed in 20% glycerol in distilled water or 0.2 M cacodylate buffer. Tissue is mounted on cardboard or standard freezing disks, depending on the particular freeze-fracture apparatus to be used, frozen on semisolid freon cooled to liquid nitrogen temperature, and fractured in a freeze-fracture device (JEOL JFD 9000). When deep-etched specimens are desired, particularly to examine cytoskeletal/membrane relationships or the surface of organelles, unfixed, non-cryoprotected cells are rapidly frozen by one of several techniques. One procedure we use is placing the cells on a thin slice of fixed lung secured to the plunger of a slam freeze apparatus (Heuser, 1989). The plunger is released, pressing the hepatocytes onto a polished copper block cooled by liquid helium. The slice of fixed lung tissue acts as a cushion, thereby preventing specimen damage. The preparation is then immersed in liquid nitrogen and stored until freeze-fracture is achieved in a standard manner. After fracturing, the tissue remains under vacuum in the freeze-fracture apparatus for 1-1 5 min before rotary or unidirectional shadowing. This procedure applied to the isolated liver cell permits better visualization of the cytoskeletal/membrane relationship, allows views of organelle surfaces, reveals the contours of glycogen clusters, and enhances viewing of intramembranous particles. In the native cell, there is a difference in intramembranous particle number between the bile canalicular domain and other domains of the membrane; in the isolated cell, the membrane is uniform in intramembranous particle distribution. Any other contemporary morphological preparative procedure is appropriate for application to the isolated hepatocyte preparations. The cells are amenable subjects for standard cytochemical, in situ hybridization, and high voltage electron microscopic techniques.

5.4. Morphological studies yet to be exploited Isolated hepatocytes in suspension and hepatocytes in tissue culture

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should prove valuable as tools for answering a number of questions that would benefit from morphologic approaches. Amongst them are the relationships between the cytoskeleton, cell junctions, the position of the Golgi apparatus, and vectorial transport. With immunochemical markers available for all of the components in this inter-related system, the hepatocyte offers special advantages over other cells where such relationships might be studied. A major advantage of the hepatocyte is that the cell loses certain structural features during isolation that re-emerge upon culture. The sequence and nature of these changes can therefore be studied. Also ready for current application are the multiple freeze-fracture cytochemical techniques, both for membrane proteins and certain lipids (Friend, 1982; Pinto da Silva, 1987; Severs, 1989). Do distinctive intramembranous particles, known antigenic determinants and glycoconjugates (Odin et al., 1986) disperse in the plane of the membrane concomitantly, or is there a difference in their temporal sequence for dispersion and re-formation? Can any direct association be established by freeze-fracture cytochemistry and rapid-freezinddeep-etching to show which intramembranous components are associated with the cytoskeletal elements? Applied to the nucleus, freeze-fracture could answer whether loss of polarity and secretory function in the isolated cell is paralleled by a decrease in number, or changes in spatial distribution of nuclear pores. Can the various biochemical studies on interaction with cholera toxin be complemented by morphologic examination (Janicot et al., 1988)? What are the changes in the plasma membrane that occur during binding, internalization, generation of A 1 peptide, and activation of adenyl cyclase during cholera toxin action? Is there an alteration in the bile canalicular and other domains of the cell membrane with respect to lipid distribution as probed by freezefracture cytochemistry with filipin, polymixin B, gold conjugated adriamycin and neomycin, and other lipid probes (Friend, 1982)? Are the changes in redistribution of membrane proteins synchronous with the redistribution or modification of membrane lipids?

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5.5. Summary of studies by transmission electron microscopy It has been well-established that the ultrastructural morphology of the isolated hepatocyte is similar to that of the cell in the intact organ. The major differences relate to the endocytosis of the junctional complex with its secondary effects on the relocalization of actin and lost orientation of the Golgi apparatus on hepatocyte isolation. Loss of vectorial transport appears to coincide with the structural changes. The isolated hepatocyte system, extremely useful to many investigators through the years, has yet to be fully exploited with current electron microscopic techniques that incorporate freeze-fracture, rapid-freezing and deep-etching, freeze-fracture cytochemistry and immunocytochemistry of frozen thin sections. I t is probable that the application of these techniques to the isolated hepatocyte to answer basic questions of cell biology would bring about a burst of new and exciting information.

5.6. Scanning electron microscopy To date, information gained about isolated liver cells by scanning electron microscopy has largely been supplemental to that gained with the higher resolutions achievable by transmission electron microscopy. Nevertheless, the use of the scanning microscope has proved popular because of the striking three dimensional images of the cell that are revealed (Fig. 5.9). The isolated hepatocytes appear rounded and, in the case of intact cells, virtually covered with microvilli; such a pattern of spatial distribution of microvilli over the irregular cell surface is virtually impossible to assess efficiently by transmission electron microscopy. Scanning microscopy (Fig. 5.10) has also proved a useful technique for revealing blebbing on the cell surface due to cellular injury (Jewel1 et al., 1982; Orrenius et al., 1983) for similar reasons the ability to survey at relatively high resolution large expanses of an irregular surface.

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Fig. 5.9. Scanning electron micrograph of intact isolated hepatocyte. Note the carpet of microvilli and the spherical shape, quite distinct from the shape of liver cells in situ. (Bar 8 pm, mag. x 3000).

Application of the newer high resolution field emission scanning electron microscopes to image the surface of isolated hepatocytes may allow the efficient detection of isolated plasma membrane-associated defects in the cells not likely to be found by transmission electron microscopic sections. In addition, imaging of the cell interior by field emission scanning microscopy of fractured cells from which the cytosol has been leached may provide, with greater efficiency, new insights into isolated hepatocyte structure and experimental structural disturbance, albeit with somewhat lower resolution than is achievable with

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Fig. 5.10. Scanning electron micrograph of an isolated hepatocyte that has developed multiple blebs as a consequence of exposure to pentachlorophenol. The blebs do not exhibit microvilli. (Bar 5 pm, mag. x 3750).

freeze-fracture and freeze-fracture/deep etching techniques, based on transmission electron microscopic approaches. 5.6.1. Technique

For scanning microscopy, cell suspensions can be fixed in 2.5% glutaraldehyde in phosphate buffer (pH 7.4) post-fixed in 1% osmium tetroxide, and then dehydrated in graded ethanols. The cells can be sedimented by gentle centrifugation (50 x g ) at the end of exposure to each solution, but should not be tightly packed. After transfer to

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amyl acetate, the resuspended cells are placed in small chambers made of two nucleopore filters clamped between three 1 cm outer diameter annular magnets. The suspensions are critical point-dried from the amyl acetate using liquid CO, (Pelco Model H unit).The nucleopore filters with adhering cells are mounted on stubs, sputter coated with gold and viewed and photographed at 5-20 kV (Siemens/E.T.E.C. Autoscan scanning electron microscope).

CHAPTER 6

Biochemical properties

6. I. Bcrsis ,fiw expressing cell activity Research workers are often faced with the need to compare experimental findings with data obtained by themselves or others in previous studies. Comparisons of this kind are facilitated if the results can be expressed on a common basis, for example dry weight, or protein content. In the case of cell preparations, such as the isolated hepatocyte suspension, results may also be expressed in terms of cell number. There is often a requirement to compare the behaviour of cell suspensions with the performance of the intact liver in vivo or with the perfused organ. This Section decribes various ways in which the function of isolated hepatocytes can be expressed and related to the performance of the whole organ. However, because of the difficulties arising from dynamic changes in relative numbers, ploidy and volume of hepatocytes during foetal and postnatal life, the following comments relate mainly to the livers of rats between 150 and 300 g body weight. Values given to convert activities in isolated hepatocytes to whole liver are not applicable to foetal and early postnatal liver.

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6.1.1. Hepatocyte dry weight

The cell solids form about 30% of the centrifuged pellet weight derived from a liver cell suspension and tend to be the most consistent component, but the ratio of pellet wet weight to dry weight varies under different metabolic conditions. In view of this variability, the causes of which are discussed in Section 6.3.1, metabolic results are probably best expressed on a dry weight basis. In order to avoid the variation introduced by differing glycogen content under various nutritional conditions, the acid-precipitated glycogen-free dry weight, which is mainly protein (Table 6.1), is preferred. Protein content in 24 and 48 h starved livers falls by approximately 3% and lo%, respectively (Harrison, 1953). It follows that there will be a similar decrease in the acid-precipitable dry weight.

Protocol 6.1 Determination of acid-precipitable pellet dry weight This is obtained by washing and drying the PCA or trichloroacetic acid (TCA) precipitated pellet from a volume of the cell suspension identical to that used for the experimental studies (e.g. 1 ml). No acidprecipitable material, such as albumin, should be present in the cell suspension. (i) Quickly transfer 1 ml of well-mixed cell suspension to each of three tared glass tubes and add 1 ml of cold 1 M PCA or 10% TCA. (ii) Mix well and stand the tubes on ice for 10 min. Centrifuge at 1500 x g for 3 min at 0 - 4 " C . (iii) Pour off the supernatants and resuspend the pellets in 2 ml of 1% TCA. Centrifuge again, drain and dry the pellets at 98°C for at least 3 h. Increasing the temperature to 105°C causes charring of the pellets as does insufficient washing of the pellets with 1% TCA. (iv) Weigh the tared tubes and pellets as soon as possible after cooling to avoid uptake of atmospheric moisture, but not before the tubes

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are completely cooled. Calculate, by difference, the dry weight of cells per ml of suspension. 6.1.2. Relationship between isolated cell dry weight, cell wet weight and whole liver wet weight

The metabolic function of whole liver is frequently expressed on the basis of activity per g wet weight. It might be anticipated, therefore, that measurement of the wet weight of a cell pellet might provide a better comparison with the metabolic activity of whole liver than determination of dry weight. However, this comparison is not as straightforward as might be thought, because a substantial portion of the mass of the intact organ is not comprised of hepatocytes. The liver from an adult 24-h fasted rat is comprised of 16% extracellular volume, which includes the lumina of sinusoids, the space of Disse and the bile canaliculi. Non-hepatocytes occupy a further 6% of liver volume. While only 6 0 4 5 % of the cells found in the adult rat liver are parenchymal cells, i.e. hepatocytes, they are much larger cells than any others present and represent approximately 8OYo of the total volume of the organ (Weibel et al., 1969; Blouin et al., 1977). It follows that, in the fasted state, a gram of liver may contain only 0.8 g of hepatocytes. On the other hand an isolated cell suspension, prepared under standard conditions, will comprise 95-98% hepatocytes. It might seem, therefore, that the activity of a pelleted isolated cell suspension could readily be converted into a corresponding activity for whole liver by using a multiplication factor of about U0.8. However, as discussed in Sections 2.6.2 and 6.3.1, the water content of isolated hepatocytes will vary considerably depending on experimental conditions. Moreover, to estimate isolated cell wet weight, it is not sufficient merely to centrifuge the cell suspension and determine the pellet wet weight. The pellet will contain a substantial amount of entrained medium, and this too will vary depending on the experimental circumstances. The volume and thus the weight of this entrained fluid must be known if net cell

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wet weight is to be accurately determined. This involves incubation with an extracellular marker, such as (hydr~xy['~C]methyl)inulin (Section 6.2.5). Thus, cell wet weight, estimated by centrifugation of a cell suspension, is not a parameter that can be assumed to bear a constant relationship to a given quantity of whole liver and in any case is not readily determined. Nevertheless, because much metabolic data relating to hepatic metabolism is expressed in terms of liver wet weight, it is desirable to have some means of comparing results from isolated hepatocyte studies with this large body of data. A generally accepted solution to this problem is to use an empirical factor to convert cell dry weight to whole liver wet weight. Using livers from adult rats, Berry and Kun (1972) determined that the ratio of fresh liver weight to dry weight of TCA-insoluble material was 3.77. Krebs et al. (1974) suggested a ratio of 3.7 after examining wet and dry weights (not acid-precipitated) from fed and fasted rat livers. These values do not allow for the contribution to the dry weight of the whole liver made by non-parenchymal cells, nor for the presence in the suspension of damaged and therefore inert cells, but these two complicating factors tend to cancel each other out. Hence, the use of a factor to convert isolated cell dry weight to equivalent wet weight of liver provides a reasonable mechanism for expressing results in terms of whole liver activity, and at the same time allows comparison of data from different experiments or even different laboratories. In most of our publications, the term 'wet weight', used as a basis to express isolated cell activity, is a derived term obtained by multiplication of the measured acid-precipitated dry weight by the factor 3.77. However, data in Table 6.1 are calculated using recent results obtained for the wet weight/dry weight ratios of whole livers from hooded Wistar rats. Ratios may vary depending on animals strain and experimental conditions. In this book, 'pellet wet weight' refers to the actual weight of a pellet of centrifuged cells, and includes the weight of entrained water. The pellet wet weight, obtained under standardized conditions, is useful as a guide to the total cell yield (Protocol 2.4).

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6.1.3. Protein and D N A content of heputocytrs

A number of workers in the field express cellular activity on the basis of protein or DNA content. As mentioned in Section 6. I . I , the protein content of the rat hepatocyte is dependent on the nutrition of the donor animal and can decrease by 3--10'%! during a 24-48 h fast (Harrison, 1953). In adult mouse, rat and human hepatocytes, Epstein (1967) found that cell mass is almost directly proportional to cell ploidy. Also, the DNA content of the liver is not altered by fasting (Harrison, 1953). Many studies have found a direct correlation between cell ploidy and metabolic activity, e.g. albumin synthesis, leucine or methionine incorporation into protein (Le Rumeur et al., 1983), glucose-6-phosphatase and glycogen synthetase activities (Bernaert et al., 1979), NADPH cytochrome c reductase and succinate dehydrogenase activities (Tulp et al., 1976), and the quantity of some nuclear membrane-bound receptors for dexamethasone (Tulp and Sluyser, 1977). However, LDH (Tulp and Sluyser, 1977) and cytochrome P-450 concentration (Sweeney et al., 1978) were found to be unrelated to DNA content. In most cases therefore, the use of either protein or DNA as a basis of expression of cell activity would appear satisfactory. 6.1.4. Cell number

The technical aspects of cell counting are discussed in Section 2.6. While the counting of cells in a Coulter counter is relatively accurate and convenient (Stewart, 1989), damaged and clumped cells are not differentiated and the equipment is expensive. The counting of cells in a haemocytometer chamber is liable to considerable inaccuracies. but enables cellular activity to be expressed on the basis o f cells that exclude trypan blue, thereby removing damaged cells from consideration. The expression of activity on the basis of cell number is popular in part because of its convenience. Furthermore, cell numbers do not change on fasting of the animal. However, since it is known that fol~owing weaning in the rat ( g 3 weeks, 50 g) hepatocytes increase in ploidy

126


and volume rather than in number (Section 2.6.3), and many cellular processes are directly related to ploidy (see Section 6.1.3), a serious error will be introduced when comparing, on the basis of cell number, metabolic activities of cells from the livers of rats of body weight 150 g ( z 6.5 weeks) with those obtained from livers of 300 g ( P 14 weeks) animals. 6.1.5. Choice of measure for expressing cellular activity

From this discussion it will be clear that none of the bases chosen for the expression of cellular activity is entirely satisfactory, though for general purposes, we favour the dry weight after acid-extraction, converted to a whole liver equivalent wet weight by means of the factor 3.77. The technique is quick and simple, gives reproducible results and, as has been discussed, is at least as valid as other measures commonly used. For any particular series of studies where constant experimental conditions are to be maintained, any of the above references will prove satisfactory. Often, however, an experimenter will wish to compare results with those obtained by other workers, and will find that an entirely different means of expressing cellular activity has been used. In order to overcome this difficulty, conversion factors have been compiled in Table 6.1. It is obviously desirable that workers publish the weight (and preferably age), sex, strain and nutritional state of experimental animals, as well as the exact method used to express results (e.g. how the wet weight of liver is determined).

6.2. Measurement of cellular composition 6.2.1. General considerations

The measurement of the intracellular content or concentration of metabolites or ions is a frequent feature of experimental studies. For solid organs, such as the liver, a relatively simple approach can be

Ch. 6.


127

TABLE 6. I Bases for expression of hepatocyte activity

Parameter

Nutritional status Fed

Reference

Fasted

Hepatocyte number

per g whole liver per g whole liver per g wet weight* of isolated hepatocytes per g dry weightt of isolated hepatocytes

95 x 106 100 x 106

130 x lo6 140 x lo6

Greengard et al. (1972) Weibel et al. (1969) Our data

377 x 106

465 x 10'

Our data

Meosured we1 weight/dry weight ratio

Whole liver (no acid precipitation) Whole liver? Isolated hepatocytest

3.95

3.58

Krebs et al. (1974)

3.75 3.82

3.35 3.36

Our data Our data

Protein content ( m g )

per g whole liver per lo6 isolated hepatocytes per g wet weight* of isolated hepatocytes per g dry weightt of isolated hepatocytes

I80 I .7 I69

222 I .5 206

Harrison (1953) Our data Our data

633

689

Our data

DNA content ( m g )

per g whole liver per lo6 isolated hepatocytes per g wet weight' of isolated hepatocytes per g dry weight? of isolated hepatocytes

2.1 0.017

2.7 0.015

Harrison (1953) Our data

1.7

2. I

Our data

6.3

7. I

Our data

*Wet weight calculated from our data for wet weightldry weight ratios of whole liver. ?Dry weight obtained by PCA precipitation (Protocol 6.2). SHepatocytes incubated at 37°C for 15 min. An aliquot was taken for a 12,000 x g pellet wet weight (Protocol 2.4) and a second was taken for a PCA-precipitated dry weight. Our data was obtained from hooded Wistar rats. Protein was measured using the Fohn procedure, as described by Lowry et al. (1951). after precipitation of the protein with 5% TCA. BSA fraction V was used as a standard. DNA was measured using the Burton method (Burton. 1956) and dAMP as a standard.

I28


adopted. A biopsy specimen can be taken if the metabolites under study are non-labile. On the other hand ‘freeze-clamping’ of the tissue with pre-cooled aluminium tongs (Hohorst et al., 1959; Wollenberger et al., 1960) can be undertaken if the lability of the metabolite requires this approach. For isolated cells in suspension, where the ratio of extracellular to intracellular volume may be 20-fold greater than in the whole liver, these relatively simple measures are precluded, particularly if the medium in which the cells are suspended contains the same substances as are to be measured in the cells. Fortunately, the widespread use of isolated liver cells for experimental studies has resulted in the development of a number of satisfactory techniques for separation of hepatocytes from the suspending medium and their subsequent analysis. If the suspending medium is free of the cellular consitutent under study, and all that is required is expression of the amount present on the basis of cell dry weight (or derived wet weight), a separation step may be unnecessary, provided that the intracellular content of the constituent is relatively high. In any incubation mixture the volume of intracellular water is likely to be less than 5% of the total volume of the suspension. Hence, the concentration of any totally intracellular metabolite will be diluted to micromolar levels on PCA extraction, even though it is present in millimolar concentrations within the cell. For many cellular metabolites, therefore, centrifugation of the cells will be required to achieve the necessary analytical sensitivity (Section 6.2.2). If cell centrifugation is not needed, a measured portion of the cell suspension is rapidly mixed with an equal volume of icecold 1 M PCA, and the precipitated protein centrifuged. Care must be taken to ensure that complete extraction of the metabolite or ion takes place, for example by measuring recovery of added standards and that no losses occur during the extraction procedure, particularly if the constituent to be analysed is labile. The same protocol may be used if the total intracellular plus extracellular amount of a substrate or product is required. A detailed discussion of analytical procedures is given by Bergmeyer (1983).

C h . 6.


129

For lipid-soluble substances such as fatty acid or triacylglycerol, extraction with heptane or, if lipid subfractionation is required, more complex extraction and separation procedures, must be employed (Borgstrom, 1959; Hara and Radin, 1978; Kaluzny et al., 1985). At the end of the incubation, a measured sample of the suspension is removed for lipid extraction and the remainder precipitated with PCA in the usual manner.

Protocol 6.2 Protein precipitation and neutralization of hrpatocyte suspensions (i) The following equipment is required: ( a ) Calibrated glass centrifuge tubes. ( b ) Refrigerated bench centrifuge, e.g. Heraeus Christ Minifuge T. ( c ) Optional: automatic titrator, e.g. Radiometer TTT60. ( d ) Optional: autoburette, e.g. Radiometer ABU.

(ii) Precipitate protein by adding an equal volume of cold I M PCA to each volume of incubation mixture. Mix thoroughly and stand tubes on ice for 10 min. (iii) Pour each solution into a plastic tube and centrifuge at 1500 x g for 3 min at 0 4 ° C . Pour the supernatant into a calibrated glass tube and record the volume (V,). (iv) Using an automatic titrator and burette (or by hand), neutralize the sample with 2 M KOH/0.2 M KCI to pH 6-6.5 (if the metabolite under study is unstable in this range, titrate to an appropriate pH). Record the volume of KOH/KCI used (V,). (v) Allow the sample to stand on ice for at least 30 min to precipitate potassium perchlorate. Centrifuge again at 1500 x g for 3 min at 04°C. (vi) The neutralized supernatant can be assayed for water-soluble metabolites. Make allowance for dilution of the sample during precipitation and neutralization as in Equation 1.

130


CF = CM x 2 x

VI + v2 VI

Where C, = concentration of metabolite in initial sample. C, = measured concentration of metabolite in the neutralized extract. 6.2,2. Separation of stable intracellular and extracellular components by microcentrijiugation of hepatocytes If, on occasion, interest centres round the constitution of the medium rather than the cells, rapid filtration of the medium is feasible (Mauger et al., 1984; Hummerich et al., 1988). Generally, however, centrifugation rather than filtration of the suspension is favoured as the means of separation since this allows the analysis of cell constituents as well. To avoid, or at least diminish, breakdown or leakage of intermediates during the centrifugation procedure, a high centrifugal force (110,000 x g ) is recommended, so that centrifugation time can be reduced to a fraction of a minute. Usually, the amounts of tissue available for analysis are small, so use of a microcentrifuge is necessary. Moreover, the microcentrifuge has other desirable features such as rapid acceleration and braking, portability (allowing transfer to the cold-room) and convenience of use, all of which are important in reducing the time during which cells may be exposed to ‘warm-hypoxia’, a condition highly conducive to the breakdown of labile metabolites. A number of microcentrifuges are available commercially. We have had experience with both the Beckman Microfuge 11 and the Eppendorf model 5415 and found each to be satisfactory. For the rapid centrifugation techniques described in this Section, a single speed and centrifugal force (usually 12,000 x g ) is adequate. A staggered start and finish is necessary if multiple incubations are being carried out. Following centrifugation, the pellet as well as the supernatant can be extracted with PCA, neutralized with 2 M KOW0.3 M morpholinopropane sulphonic acid (MOPS)/0.2 M KCl to pH d 6 . 5 and water-soluble metabolites measured. The inclusion of

Ch. 6.


131

MOPS, when titrating small volumes, reduces the risk of overshooting the end-point. Where necessary, contamination of the pellet with entrained supernatant can be measured by including 3H,0 and (hydr~xy['~C]methyl)inulin in the incubation (Protocol 6.3). This simple method of separation of cells and medium by microcentrifugation is only suitable for non-labile intermediates, and in any experimental situation, preliminary recovery studies are essential to determine its validity. The values obtained with this method should be compared with those obtained using techniques needed for measurement of labile metabolites (Protocol 6.5). If the results are the same with both methods, the less complicated approach may be used. However, there is a large amount of literature relating to metabolite stability, and the likelihood is that for many substances of interest, the preliminary effort to ascertain whether or not the compound under scrutiny is sufficiently stable to employ this approach may well have already been carried out. 6.2.3. Separation of labile intracellular and extracellular components

When there is a possibility that the metabolite may be modified by enzyme action in the time taken to form the pellet and decant the supernatant, it is necessary to centrifuge the cells into a solution such as ice-cold PCA, that virtually instantaneously destroys all enzymatic activity. Separation of the PCA from the suspending medium is achieved by interposing a layer of silicone oil between the two solutions (Zuurendonk and Tager, 1974; Zuurendonk et al., 1979). In this three-layered system, the concentration of PCA must be sufficient to achieve rapid and complete protein precipitation and its density must be greater than that of the silicone oil fraction, which in turn must be denser than the incubation mixture. The density of the silicone oil mixture is selected so that it effectively partitions the cells and their suspension medium. The composition of the medium into which the cells are centrifuged can be varied to suit the subsequent analytical procedure. For

I32


example, the lowermost layer could be a sucrose solution containing Triton X-100if the enzyme content of the pellet was of interest. Contamination of the bottom layer (including the pellet) by suspension medium adhering to cells during centrifugation through the silicone oil layer, can be estimated by including (hydr~xy['~C]methyl)inulin with or without 3 H 2 0in the medium (see Section 6.2.5 and Protocol 6.3). Other oil layers including brominated hydrocarbons (Cornell, 1980) and dibutyl phthalate (Danon et al., 1966; Meredith and Reed, 1982) (also see Section 7.3.2) have been used in similar procedures. Fariss et al. (1985) found that centrifugation through these oils does not significantly change the intracellular concentration of K', Ca2+, LDH, aspartate or glutamate but that during centrifugation through brominated hydrocarbons, 20% of GSH was oxidized. Also, Katz and Wals (1985) noted a loss of glucose-6-phosphatase and adenylate kinase activity after centrifugation through brominated hydrocarbons. 6.2.4. Measurement of' the intracellulur constituents qJ cen trifugrd hepatocytes

If it is desired to express the quantity of a metabolite or ion in terms of the actual (rather than derived) wet weight of the cells or as a concentration, a centrifugation step is essential so that the volume of intracellular water can be determined. Centrifugation is also necessary when the intracellular constituent under examination is also present in the suspending medium in an amount likely to cause interference with accurate measurement of the intracellular content. I t must be appreciated, however, that the pellet water is not synonymous with the intracellular water. In fact, a substantial quantity of suspending medium is trapped within the centrifuged pellet. Thus, either to allow for contamination, or so that cell content can be expressed in terms of actual concentration, measurement of the entrained medium is essential to determine the true intracellular water content. The theoretical estimate of the extraparticulate space for the closest

Ch. 6.


133

possible packing of spheres is 26%)(Conway and Downey, 1950).Since the hepatocytes undoubtedly undergo some deformation on centrifugation, it is not possible to assume this value for pelleted hepatocytes. Moreover, the cell surfaces are not smooth, but virtually completely covered with microvilli that may prevent close intercellular contact and serve to increase the amount of entrained medium. I t must also be remembered that a proportion of the cells in the suspension will be sufficiently damaged to allow the penetration of extracellular medium, thereby contributing to the apparent extracellular space. In our experience the volume of entrained medium may contribute 25--60% of the pellet water, depending on the centrifugal force employed to form the pellet. Obviously such a major contribution cannot be ignored if a realistic measurement of the water content of intact hepatocytes is to be made. 6.2.5. Use of inulin and other agents j o r measuremrnt yf'c~?rtracellular water

A wide variety of molecules that do not penetrate intact hepatocytes have been used to determine the volume of entrained medium in a hepatocyte pellet. Probably the most widely used is inulin, a molecule of approximate M r 5000, which is able to pass through the plasma membrane of damaged cells but which is excluded by those that are intact. Thus, its concentration in a pelleted fraction will allow measurement of the total volume of entrained medium, including the intracellular volume of damaged cells. 'H,O added to the same cell suspension equilibrates with both intracellular and extracellular water and provides a measure of the total intracellular water of intact and damaged cells plus the entrained medium. Although it is feasible to use unlabelled inulin (Beutler, 1984), the use of radiolabelled marker greatly facilitates the procedure. The researcher has a choice of tritiated or ''C-labelled inulin derivatives. In studies where total pellet water is being determined simultaneously in a double-labelling procedure using >H,O (Protocol 6.3), there is

134


no alternative but to use a I4C-labelled derivative. In other circumstances, [3H]inulin might seem equally satisfactory. However, in practice we find this is not the case. Chemiluminescence induced by compounds in pellet extracts is more troublesome when counting tritium. Moreover, inulin derivatives break down on storage to compounds of smaller M , that can enter the hepatocytes and produce an overestimate of the amount of entrained medium. This decomposition of inulin can be minimized by its storage at -70°C but preparations of tritiated inulin seem more susceptible to degradation. For these reasons the use of (hydr~xy['~C]methyl)inulin is recommended, even though it is more expensive than tritiated inulin. A possible concern in the use of inulin or other macromolecular markers of entrained medium is that they are slowly taken up by intact hepatocytes, a process that must lead to overestimation of the extracellular space. In experiments involving hepatocyte incubation, it is good practice to add the labelled inulin no more than 2 min before termination of the incubation. Kolb and Adam (1976) used a variation of the method described in Protocol 6.3. By centrifuging cell suspensions containing in~lin-[~~C]carboxylic acid through a layer of medium containing 4% Ficoll (Pharmacia), they were able to estimate binding and uptake of the label by the cells. Using this method, intracellular Na' and K' concentrations were calculated to be 42.1 mM and 138.1 mM, respectively, after incubation of cells at 37°C. This correlates closely with values obtained by our laboratory using Protocol 6.3 and cells isolated under similar conditions (Table 6.2). A number of other agents have been suggested as markers of entrained medium. These include [I4C]dextran (Baur et al., 19751, [14C]poly-dextran(Hoek et al., 1980), [45Ca]EGTA(Hughes et al., 1987), 1251-labelledalbumin (Krebs et al., 1974), [WrIEDTA (Stacy and Thorburn, 1966) and [14C]sucrose(Zuurendonk et al., 1976). In most instances, comparisons were made with values obtained using either 3H- or I4C-labelled inulin compounds and results were found to be identical. The exception was [14C]sucrose,which was found to

Ch. 6.


135

permeate the plasma membrane of cells at stages where they were still impermeable to large molecules. Gordon et al. (1985) found sucrose can diffuse through surface blebs on freshly isolated hepatocytes suggesting that [14C]sucroseshould not be used as an extracellular marker. lZsI and W r are both gamma-particle emitters and require more stringent safety precautions. It is recommended therefore, that [3H]inulin should be used in preference to lZSI or S'Cr as an external marker in incubations where the presence of 4sCa or I4C is also necessary. An accurate method of determining the intracellular and extracellular water content of a centrifuged cell pellet is to include both 'H,O and (hydroxy[14C]methyl)inulin in the incubation. By measuring the ratios of 'H to I4C counts in both the supernatant and pellet fractions, it is possible to calculate the intracellular volume of intact cells per mg dry weight at various times during an incubation. To minimize problems of isotope overlap between 3H and I4C during dual channel counting of the samples, the starting isotope ratio should be at least 1O:l (Lund and Wiggins, 1987).

Protocol 6.3 Measurement of extracellular and intracellulur water in a cell pellet (i) Add 400,000 dpm of jH,O to 2 ml of incubation medium containing approximately 30 mg dry weight of cells. Proceed with a normal incubation. (ii) Two min before sampling the incubated suspension, add 40,000 dpm of (hydr~xy[~~C]methyl)inulin in a small volume (20 pl) of 1 pM inulin. If inulin is added for the entire incubation period, there is some risk of cellular uptake due to pinocytosis by undamaged cells. (iii) At predetermined times, quickly pipette 1 ml of the well-mixed suspension into a microcentrifuge tube and centrifuge at 12,000 x g for 20 s. The remaining incubation mixture may be analysed for substrates, products, etc.

136

ISOLATED H EPATOCYTES

(iv) Pipette a portion of the supernatant (S) into a glass scintillation vial. A convenient volume is 200 pl. Add 5 ml of scintillation fluid (e.g. ACSII, Amersham), cap tightly to prevent loss of 3H,0, mix well and analyse in a counter (e.g. Beckman LS 3801) that has been preprogrammed to discriminate 3H and I4C and allow for sample quenching. (v) Using a fine needle and syringe, remove the remaining supernatant from the microcentrifuge tube and dry the sides of the tube with tissues or cotton buds. (vi) Resuspend the pellet with a vortex mixer in a measured volume of H,O,(Vw),e.g. 500 pl. Lyse the cells by freezing and thawing three times using liquid nitrogen. Centrifuge at 12,000 x g for 5 min and then pour the pellet extract (P) into a clean microcentrifuge tube. (vii) Pipette a portion of the pellet extract (200 pl is convenient) into a glass scintillation vial and count as in (iv). (viii) Measurements and calculations. First step: Determine values for H, H, C, C,

= dpm per ml for 3H in S. = dpm per ml for 3H in P. = dpm per ml for I4C in S. = dpm per ml for I4C in P.

Second step: Calculate volume of water in whole pellet (V,) (cells plus entrained medium). The value of H, is a function of the relative magnitudes of Vw and V,. i.e.

Solving for Vp vp

=

HP x v w Hs - HP

Ch. 6.


137

Third step: Calculate total volume of entrained water in pellet, including the cytoplasmic water content of damaged cells (VE). Total I4C dpm in P = VE x CS

c p

=

VE

cS

vw

+ VP

Solving for VE

Fourth step: Calculate volume of intracellular water of intact cells (V,)

(ix) The dry weight of a measured portion of cells should be determined as in Protocol 6.1. V, can then be expressed as ml/g dry weight of cells. Note,. Protocol 6.4 contains additional steps that are required to obtain values for the concentrations of intracellular Na' and K ' or metabolites.

Protocol 6.4 Measurement of intracellular concentration of ions or metabolites (i) At step (iv) of Protocol 6.3 transfer a second 200-pl portion of supernatant (S) to a small tube. Stopper tightly to prevent evaporation and concentration of the analytes. Following step (vi). transfer an appropriate measured amount of pellet extract (P) to another tube. (ii) Analyse samples of S and P for Na+ and K+ concentrations,

I38


using a vented flame photometer (e.g. Instrumentation Laboratory Model 943). (iii) Calculate Na; (intracellular Na+ concentration) and K; (intracellular K+ concentration). Let: Naa = Na+ concentration in S . K; = K+ concentration in S . Na; = Na+ concentration in P. K; = K+ concentration in P. Total Na' in P = Na; (V,

+ V,,).

The Na+ in P that is within entrained medium and damaged cells = V,Na; It follows that: Na; =

Na; (V,

+ Vp) - VENa{ VI

Similarly: K: =

K+P(V,

+ Vp) - VEK{ VI

Intracellular concentrations of stable metabolites are calculated similarly. In these circumstances, the pellet is extracted with PCA instead of H,O (Protocol 6.3(vi)) to stop further enzyme activity. A measured portion of supernatant is also acidified. Both fractions then require neutralization with 2 M KOH/0.3 M MOPW0.2 M KCI and removal of potassium perchlorate before analysis of metabolites by appropriate analytical procedures. The above equations can be modified to allow for dilution of the samples during acidification and neutralization.

Ch. 6.


139

Protocol 6.5 Centrifugation of hepatocytes into PCA through a layer of silicone oils (i) Prepare microcentrifuge tubes (e.g. Eppendorf 38 12, capacity 2.2 ml) by overlaying 0.2 ml of 8.5% (w/v) (approximately 1.5 M) PCA with 0.65 ml of a silicone oil mixture (Dow Corning 550 and 200, 1.5 centistokes oils, in the ratio 7: 1 (v/v), respectively ( p z 1.03 at 25°C). Centrifuge the tubes for 15 s in a microcentrifuge to consolidate the layers. Store the tubes at 0 4 ° C in an ice slurry. (ii) To allow sufficient time for processing of each sample, incubations are commenced at 2-min intervals. At appropriate times, hold the microcentrifuge tubes vertical and carefully but rapidly pipette 1.4 ml of incubation mixture down the side of the tubes to form a layer on top of the silicone oil layer. (iii) Centrifuge the tubes for 20 s in the microcentrifuge. The cells will pass through the silicone oil layer into the PCA. If a layer of material forms on the top surface of the oil, this will represent some damaged cells which are not sufficiently dense to pass through. If desired, a less dense silicone oil layer, containing relatively more 200 oil, can be made which allows these cells to pellet in the bottom layer as well. However, it will then be more difficult to add the top layer without disturbing the oil. (iv) Transfer 1.2 ml of each of the uppermost aqueous layers (representing extracellular medium) into microcentrifuge tubes containing 0.2 ml 50% (w/v) PCA. Mix thoroughly and store on ice. When convenient, centrifuge in order to pellet precipitated protein and then neutralize the supernatants with 2 M KOH/0.3 M MOPY0.2 M KCl and record the amounts required. Store the neutralized samples on ice for 30 min and then remove the precipitated potassium perchlorate by centrifugation. (v) Carefully remove the remaining upper and silicone oil layers and discard. Wipe the walls of the tubes clean, using cotton buds. Alternatively, the upper aqueous layer, only, may first be removed, and water added above the silicone oil to dilute the residual fluid. Both

140


washing and silicone oil layers may then be removed and discarded. (vi) Vortex the pellet/PCA layer vigorously. (vii) Centrifuge the tubes to re-pellet precipitated protein and transfer portions of supernatant for neutralization with KOH/MOPS/KCl followed by removal of potassium perchlorate as in step (iv). (viii) Analyse the neutralized and clarified supernatant and pellet extracts for water-soluble metabolites and correct for the concentration and dilution steps. Calculate the amount of extracellular and intracellular metabolite(s) per mg dry weight of hepatocytes. (ix) Check that all metabolites have been recovered by comparing their total sum in supernatant and pellet extracts with that obtained by PCA-precipitation of untreated cells. Note. This is the method of choice for the determination of unstable metabolites or where for reasons other than instability, a reaction needs to be stopped quickly.

Protocol 6.5 may be modified: (a) to allow the measurement of intracellular enzymes by substituting a 0.5 M sucrose/l'%,Triton X-100 mixture for 8.5'%, PCA acid. In this case, acidification and neutralisation steps are omitted; (b) to determine the amount of entrained medium which has passed through the silicone oils layer, by including 3 H H , 0and (hydr~xy['~C]methyl)inulin in the incubation. This will enable measurement of intracellular metabolites to be expressed in terms of concentration, since the intracellular volume can now be calculated (Protocols 6.3, 6.4). Under these conditions P comprises the volume of the lower PCA layer (0.2 ml) plus V,, the volume of water contained by the cells and entrained medium.

6.3. Cellular composition 6.3.I . Water and ion content

The water and ion content of hepatocytes depends to a large extent on their metabolic activity. For example, liver cells lose 2.4 g of water

Ch. 6.


141

for each gram of glycogen lost during fasting (Puckett and Wiley, 1932). Hepatocytes are capable of achieving and maintaining substantial concentration gradients of Na+ and K+ across their plasma membranes (Table 6.2). Because the gradients are maintained by energy-dependent mechanisms, it is not surprising that the cell isolation procedures, in which cells are cooled and subjected to hypoxia, lead to a loss of ion gradients, i.e. an inflow of Na+, outflow of K+ and an uptake of water (Table 6.2). Cells prepared according to the one-step method (Protocol 2.2) with restoration of Ca2+to the medium at the end of the perfusion phase, or by the two-step procedure of Seglen ( 1976). recover rapidly from the trauma of the preparative procedures and can achieve a normal water content and Na+ gradient across the plasma membrane within 10 min of incubation at 37°C. In contrast, we find cells freshly isolated and washed in Ca?+-freemedia are more swollen and have higher Na+ and lower K+ intracellular concentrations (Table 6.2). They take longer to regain normal levels of intracellular Na+ and K+ when incubated at 37°C with Ca” and do not achieve normal intracellular Na+ and K+ concentrations when Ca?+ is not replaced. However, after 30 min incubation at 37”C, the intracellular volume of the hepatocytes is independent of the method of cell isolation provided Ca2+is present in the incubation medium. Edmondson and Bang (1981) compared cells prepared by a twostep procedure in which 2.5 mM Ca” was either added together with collagenase at the beginning of the second step or was omitted. Intracellular Mg2+ concentrations did not differ between the two methods, being 37 nmol/mg dry weight after preparation and dropping slightly to a stable value of about 30 nmol/mg dry weight after a 30-min incubation. However, the intracellular Ca2+concentration was significantly lower in the Ca”-deficient cells, 4.5 nmolimg dry weight compared with 10.5 nmol/mg dry weight in Ca”-replenished hepatocytes. The intracellular distribution of Ca” between mitochondria, endoplasmic reticulum and cytoplasm, which appears to play an important role in metabolic regulation, is discussed in more detail in Chapter 9.

TABLE 6.2 Na' and K and water content of isolated hepatocytes from fasted rats Ca" added to perfusion Ca" added t o 10-min preincubation Ca" added to washing buffer lntracellular K' lntracellular K' lntracellular K' (without Ca"

(mM) at 0 ° C (mM) at 37°C (mM) at 37°C in incubation)

lntracellular Na' (mM) at 0 ° C lntracellular Na' (mM) at 37°C lntracellular Na' (mM) at 37°C (without Ca" in incubation) Pellet wet wt/dry wt (g/g) at 37"C** lntracellular H,O/dry wt (mI/g) at lntracellular H,O/dry wt (mlig) at 37°C lntracellular HzO/dry wt (ml/g) at 37OC (without Ca" in incubation)

+

+

-

-

-

+

+

+

-

79.8 f 0.86 133.3 f 3.8

81.8 f 6.4 150.7 f 4.4 120.0 f 3.2

51.7 f 2.6 137.6 f 2.3 112.1 f 5.5

35.5 f 3.4 134.3 rt 10.8 116.7 r t 1 1 . 4

111.4 rt 4.5 34.2 rt 7.9 56.7 rt 4.8

122.7 f 6.1 32.4 f 2.1 64.3 f 7.0

134.8 f 2.6 31.4 f 7.0 63.2 f 3.4

+*

-

78.1 rt 5.02 28.0 f 5.1 -

3.24

f

0.09

3.25 rt 0.03

3.53 f 0.05

0.06

2.13 f 0.08

2.48 f 0.10

2.04 f 0.07

1.86

1.64 f 0.02

1.48 f 0.04

1.56

f

0.03

1.50 f 0.06

1.60 f 0.13

1.65

f

0.12

1.67 f 0.05

f

'Perfusion media and technique were as described by Seglen (1976). In all other preparations. media (with slight modifications) and technique were as described in Protocol 2.2. **Pellet was obtained by centrifuging a suspension incubated for 35 min at 12.000 x g for 2 min.

Ch. 6.


143

Any metabolic inhibitor is likely to cause water uptake and swelling as a consequence of a reduction in the capacity of the cells to extrude Na'. The most potent agents in this respect are uncouplers and ionophores, whereas respiratory inhibitors, such as rotenone or antimycin, are considerably less effective in inducing swelling. The inner mitochondrial membrane is sensitive to valinomycin at concentrations as low as 15 nM (indicated by a loss of mitochondrial membrane potential), whereas the plasma membrane is less responsive, so that valinomycin concentrations as high as 50 nM are required to lower intracellular K+ content substantially. On the other hand, the plasma membrane appears highly sensitive to gramicidin which dissipates transmembrane Na+ and K+ gradients at concentrations similar to those capable of uncoupling mitochondria in isolated cells. An agent frequently used in the study of Na+ and K+ metabolism is ouabain, a specific inhibitor of the Na+/K+ATPase. This enzyme appears to be responsible for transport of Na' and K+ across the plasma membrane against the concentration gradient. Rossier et al. (1987) review the regulation of the Na+/K+ ATPase and ouabain binding to the enzyme. In the presence of ouabain at a concentration of 1-2 mM, the hepatocytes are unable to achieve their normal monovalent cation gradients, and any established gradient is dissipated. Some caution should be employed in using ouabain, since its effects are not entirely limited to the Na+/K+ATPase. For example, ouabain inhibits gluconeogenesis from some substrates (unpublished observations). This is an important consideration when ouabain is used to inhibit the ATPase in attempting to measure the energy requirement for Na' and K+ transport. However, the use of ouabain in hepatocyte suspensions is far more satisfactory than its use with the perfused organ or surviving slices, where cell swelling causes major problems in terms of 0,-delivery (Claret et al., 1973). 6.3.2. Metabolite content

Procedures related to the extraction of cellular metabolites are listed

144


in Protocols 6.1-6.5. A complicating feature that has not been discussed is the presence in the suspending medium of metabolites that are generally considered to be entirely intracellular. It appears that these substances escape from damaged cells and are then not further metabolized by the intact hepatocytes. A good example of such a metabolite is malate. Up to 30% of this intermediate can be found in the extracellular fluid. Van Schaftingen et al. (1987) report similar findings in relation to hexose phosphates and glycerol 3-phosphate. These intermediates can be released early in the incubation period at a time when their intracellular concentrations are much higher than later in the experiment. It is apparent that determination of these intermediates, following simple PCA treatment of the hepatocyte suspension at the end of the incubation period, could lead to an overestimation of the content within the intact cells, as could the determination of metabolite concentration in the intracellular water of a cell pellet if measurement of the amount present in the entrained medium was omitted. 6.3.3. Lipid, protein and enzymes

The lipid, protein and enzyme content of the isola ed hepatocyte are similar to those of the hepatic parenchymal cell in situ (Seglen, 1976; Berry, unpublished observations). 6.3.4. Adenine nucleotides

The adenine nucleotide concentrations provide a useful indicator of cellular integrity, since damage to the cells invariably brings about a fall in ATP, and usually a decline in the total nucleotide content. At the end of the isolation procedure the ATP concentration and the ATP/ADP ratio are likely to be low, due to the inevitable hypoxia to which the cells are exposed during .washing and centrifuging. However, within 10 min of commencing cell incubation, normal values should be found. A low total nucleotide and ATP content, observed

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145

at this time, suggests a problem during preparation of the cells, even though the ATP/ADP ratio may be normal. We have noted that cells prepared using early restoration of Ca?+to the medium show higher levels of ATP. It is probable that, following incubation, a lowered total nucleotide and ATP content of a poor quality preparation reflects the increased number of damaged cells, as determined with trypan blue. Studies using Percoll to separate damaged from intact cells, have shown that the former are devoid of ATP and ADP (Berry et al. 1988). It can be anticipated, therefore, that the levels of total nucleotide and of ATP will be inversely proportional to the percentage of damaged cells in the preparation. In a preparation containing a relatively high proportion of damaged cells, a normal ATP/ADP ratio may still be observed, as was found by Baur et al. ( 1975) in cells aged for 2 1 h, reflecting the apparent integrity and potential metabolic competence of those cells that still excluded trypan blue. As shown in Table 6.3, under normal circumstances the principle adenine nucleotide is ATP, the hepatocytes maintaining an ATP/ADP ratio of about 4:l. Only trace amounts of AMP are present. When cells are poisoned with inhibitors of respiration there are falls in total adenine nucleotide content, in ATP and in the ATPIADP ratio, whereas the level of AMP rises substantially (Fig. 6.la,b). The TABLE 6.3 Adenine nucleotide concentrations Substrate

Nutritional state

[ATP] mM

[ ADPI [ ATPlI[ADPl mM

Endogenous

Fed Fasted Fasted

2.42 2.17 2.23

0.42 0.57 0.51

5.8 3.8 4.4

Fasted Fasted Fasted

2.45 0.48

0.50 0.51 0.49

4.9 0.9 2.1

Palmitate (2 mM) + lactate (10 mM) Glucose (40 mM) Fructose (10 mM) Glvcerol (10 m M )

1.01

I46


0

1 2 3 4 Antimycln Concentration ( pM )

0

1 2 3 4 Antirnycin Concentration ( pM

5

5

4

9 3 \

5 2

1

Fig. 6.1. Hepatocytes (30 mg dry weight) were incubated in a final volume of 2 ml Over a range of antimycin concentrations for 40 min at 37°C. ( * ) , Total adenine nucieotides; ( A ), ATP; ( ), ADP; ( 0 ), AMP; ( T ), ATP/ADP.

Ch. 6.


147

mechanism of nucleotide depletion has been explained on the basis of the activity of AMP deaminase (van den Berghe et al., 1989). The nutritional state of the animal has some impact on nucleotide levels. Thus, the nucleotide content and ATP levels of cells from fed rats tend to be slightly greater than those of cells from fasted animals (Table 6.3). Incubation of cells from fasted animals with glucose (> 10 mM) raises ATP concentrations some 10-15% by the end of a 40-min incubation. In contrast, some carbohydrates or related substances greatly deplete hepatocyte nucleotide levels. The best known of these is fructose, addition of which can lower the intracellular ATP concentration by 50% within 2 min (Woods et al., 1970; van den Berghe, 1977; Des Rosiers, 1982). Glycerol also is effective (Table 6.3) (Des Rosiers et al., 1982). The fall in ATP is associated with accumulation of phosphorylated metabolites within the cells (Woods et al., 1970; van den Berghe, 1977), but the reasons for this accumulation of phosphorylated intermediates has not been unequivocally established. It is of interest that despite this depletion of ATP the hepatocytes remain capable of the rapid conversion of fructose to glucose. Gluconeogenesis from glycerol is also not limited by lack of ATP (Berry et al., 1973).

6.3.5.Glycogen Levels of glycogen depend very much on the nutritional state of the donor animal. The level is highest in fed animals and is very low in the cells of rats fasted for 18 h. The levels in isolated cells follow those in the intact organ, and the highest cell content of glycogen is observed in cells obtained from meal-fed animals (Katz et al., 1975). However, to retain cell glycogen during the preparative procedures, it is necessary to ensure a level of at least 15 mM glucose in the perfusion and washing media. If, in the subsequent experimental study, it is desired to incubate the cells in the absence of added glucose, it may be omitted from the last washing solution. Under these circumstances, it is particularly important to ensure that a temperature close to M " C

148


is maintained during the washing procedures or else considerable glycogen breakdown will occur at this stage.

6.4. Experimental conditions Many thousands of studies of isolated liver cells have been published over the past twenty years and it is obviously not feasible to discuss them in detail here. Instead, general principles relating to the use of isolated hepatocytes for metabolic studies will be discussed, and optimal conditions for examining the major metabolic pathways of the liver cell will be delineated. It is well established that during the preparative procedure, isolated hepatocytes lose important metabolites (Pogson et al., 1984). These include dicarboxylic acids (Cornell et al., 1974), GSH (Viiia et al., 1978) and carnitine (Christiansen and Bremer, 1976). In the case of carnitine, the loss does not appear to affect metabolic behaviour, but in other instances the effect is noticeable. For example, the loss of intermediates of the malate-aspartate shuttle initially limits the rate of glucose synthesis from lactate, but this limitation can be overcome by addition of lysine or pyruvate to the cells (Cornell et al., 1974). No comprehensive set of rules covering every experimental circumstance can be given concerning the desirability of fortifying incubation media with metabolites anticipated to have been lost from the hepatocytes during the preparative procedures. Decisions about this need to be based on individual experimental studies. During incubations of more than 60 min (see Chapter lo), the cells suffer a substantial decline in cytochrome P-450, a decline which is not readily reversible. This must be taken into account when performing studies on drug metabolism (Chapter 7). Furthermore, it is reported that polysomal disaggregation is a continuing phenomenon during hepatocyte incubation (Dickson and Pogson, 1980), suggesting the possibility of the development of impaired protein synthesis. As discussed previously (Section 4.2.1 ), some cell damage during incuba-

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149

tion is inevitable. Nevertheless, metabolic rates observed with isolated hepatocytes in suspension compare favourably with those determined in the isolated perfused liver and, also for many processes, with those measured in the intact animal. 6.4.1. Incubation time and temperature

In general, the incubation time can be varied to suit the nature of the experiment. If the cells are taken from storage in the cold to be incubated at 37"C, it can be anticipated that at least 2 min will be required for stabilization at the new temperature. If accurate initial metabolic rates are required, the substrate or inhibitor under study should be added to the incubation vessel in a small volume after this brief stabilization period. At the other extreme, experience has shown that the cells can be incubated for at least 3 h at 37°C (Section 10.6.2) without incurring so much damage that results become invalid. This statement must be qualified by pointing out that much depends on the nature of the experiment. If studies are being carried out with a noxious agent such as a potent respiratory inhibitor, it is possible that all hepatocytes will be damaged and become permeable to trypan blue within 30-60 min. Long term experiments can be successfully carried out only on non-poisoned cells, incubated under optimal conditions (Section 10.6.2). For general metabolic experiments where measurement of metabolite concentrations is performed at a single time-point, an incubation period of 40 min is satisfactory provided that substrates do not become limiting within this time. This interval allows respiratory rates to be measured accurately, and is sufficiently long for metabolites to accumulate to a level that can be readily measured. In many experiments, however, we incubate cells for more than twice this length of time, using a series of flasks so that we can sample at regular time intervals. The value of such time-course experiments cannot be over-emphasized. Often, research workers choose an arbitrary time-point and make the assumption that this is all that is needed to characterize the behaviour


150

of the cells. In fact, studies over many years have revealed that in many instances the hepatocytes endeavour to establish steady states. Usually, at the beginning of an experimental incubation, the physiological steady states will be considerably perturbed, so that metabolism will be rapid as the cells endeavour to correct the distortion. Once the normal steady state is reached, metabolic flux is likely to decline. A single time-point measurement will fail to reveal this complexity. An example of this pattern is the accumulation of lactate during glycolysis shown in Fig. 6.2. For the worker interested in studying maximal rates of metabolism, it is important to ensure that experimental conditions are such that approach to steady-state is avoided. For this purpose, experimental conditions should be arranged so that the hepatocytes function far from steady state over the whole length of the experiment. In these circumstances, the metabolic flux is linear and a single time-point is adequate.

0

50 100 Incubation Period

150 (

200

min

Fig. 6.2. Hepatocytes (30 mg dry weight) were incubated at 37°C with 40 m M glucose. Total incubation volume was 2 ml ( A ) or 4 ml ( 0 ).

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151

The standard temperature for incubation of rat hepatocytes is 37°C. Temperatures of only a few degrees higher rapidly cause cell damage (Bowers et al., 1981). Cells from fed rats maintain their integrity longer than those from fasting animals (unpublished observations). On the other hand, the cells tolerate lower temperatures well although metabolic activities are depressed. Na+ and K+ gradients across the plasma membrane can be maintained at temperatures as low as 25°C (unpublished observations). 6.4.2. Cell mass, incubation volume and oxygenation

There seems little doubt that the reason the isolated hepatocyte suspension has become by far the most popular in vitro system for examining hepatic metabolism, relates to the fact that the preparation may be divided into numerous aliquots, each of which may have the potential to reproduce the pattern of activity of the whole organ. Thus, the limitations of the perfused liver preparation, where perhaps only one set of conditions can be tested in a single experiment, are overcome. An average yield of hepatocytes from the 6 g liver of a 200 g rat is about 2-3 g wet weight of cells. In deciding how many incubations can be carried out with this cell mass, the experimenter faces a number of constraints. Perhaps the most important of these relates to 0,-supply. The basal rate of 0,-uptake for cells incubated without substrate is about 7 pmol/g dry weight per min, but this can rise to 60 pmol/g dry weight per min in cells supplied with substrate and incubated with valinomycin. Studies with surviving slices (Umbreit et al., 1964) indicate the potential risk of hypoxia if cell mass is too great, although in these circumstances, 0, diffusion through the tissue slices is a limitation. We find that it is satisfactory to use a 2-ml cell suspension containing 30 mg dry weight of cells in a capped vessel in which the surface area of the incubation medium is 4-5 cmz. For convenience, we often use 20-ml glass scintillation vials as incubation vessels, but it is desirable that researchers compare the performance of such vials with that of conical flasks, before using them consistently for

152


any particular study. A dry weight of 30 mg of cells generally supplies sufficient material for most of the analyses of metabolic intermediates that may be required, without lowering the 0,-tension of the medium to hypoxic levels. I t is technically difficult to determine directly by measurement of 0,-tension in the incubation medium whether or not the hepatocytes are in a hypoxic state. In many circumstances, the finding of a steady state ratio of lactate/pyruvate in the incubation medium greater than 1O:l provides a useful indication of hypoxia. Note, however, that this ratio can be raised in the absence of hypoxia under a number of physiological conditions, such as during the metabolism of fatty acids. We have also chosen a water bath shaking speed of 120 cycledmin with an amplitude of U . 5 cm, and a gas phase of carbogen (O,:CO,,955). However, experience suggests that a gas mixture of 5% CO, in air is satisfactory for incubations of up to 60 min. A 20-ml incubation vessel, containing 2 ml of fluid will hold about 750 pmol O,,when filled with carbogen. If gassed with air, the vessel will contain about 150 pmol 0,. Maximal 0,-consumption rates for hepatocyte preparations, incubated at 37"C, are of the order of 60 pmol/g dry weight per min, so that an incubated cell suspension, containing 30 mg dry weight of cells, could be anticipated to consume, maximally, 2 pmol/min. Since the 0,-tension in the medium does not appear to become limiting for respiration until it falls to levels below a partial pressure of 10 mmHg (No11 et al, 1986), flasks gassed with carbogen should be able to supply. sufficient 0, for at least 4 h to cells respiring at maximal rates. Over a period of this length, other factors such as mechanical damage to the cells, accumulation of products, or pH changes are more likely to cause concern than any limitation in 0,-supply. For very long-term experiments there is no reason why the incubation cannot be briefly halted and the vessels re-gassed. An alternative approach is to use a gassing manifold and supply a continuous fine stream of gas to the cell suspension. Obviously, this technique is likely to promote some evaporation, but this can be minimized by first bubbling the gas through saline at 37°C. In experiments where fre-

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153

quent additions must be made to the vessels, continuous gassing may well be preferable, but, generally, a single gassing procedure followed by capping of the vessels at the commencement of the incubation proves adequate. Most of our experiments are carried out with cells suspended in 2 ml of incubation medium. One reason for choice of this volume is that an incubation vessel that we often employ is specifically designed for measuring O2 in the presence of COz and is not well-suited to larger incubation volumes (Fig. 6.4). However, a more important reason for using a small volume is to provide the opportunity for the cells to regain normal metabolite gradients as rapidly as possible. Generally, the incubation medium in which the cells are suspended will be devoid of many nutrients found in the blood. Many of these nutrients are derived from the liver, so that the isolated hepatocytes can be anticipated to secrete them into the medium. Obviously, for any given mass of hepatocytes, the smaller the incubation volume, the lesser the amount of secretion required of them before physiological concentrations are attained. This is not merely an academic point but has important practical implications. Gluconeogenesis from lactate, for example, does not reach its maximum rate in liver until a normal lactate/pyruvate ratio in the medium of about 10: 1 has been attained. In liver cells incubated with a given concentration of lactate, but no pyruvate, it is found that maximal rates of glucose synthesis are achieved 1&20 min earlier in cells incubated in 2 ml rather than 4 ml of medium. It may be asked why the incubation volume should not be reduced to 1 ml. Unfortunately, there is a serious difficulty in making such a reduction. Hepatocytes are very active towards many substrates, rates of removal often ranging between 7 and 20 pmol/g dry weight per min. It follows that 30 mg dry weight of cells may remove 0.4 pmol of substrate/min, equivalent to 16 pmol during a 40-min incubation period. If the substrate was added at a concentration of 10 mM at the start of the incubation, it is evident that it would be totally depleted from a I-ml incubation volume and only 20% would remain if the incubation volume were 2 ml. This problem of substrate depletion is

154


a very serious one and is difficult to deal with. It is recognized that the rate of uptake and metabolism of many substrates, presented to the liver cells at physiological levels, is highly concentration-dependent. Hence, a continually falling concentration in the incubation medium will in turn lead to a continually falling rate of metabolism. The obvious answer might seem to be to load the incubation medium with much higher initial substrate levels. However, exposure of the hepatocytes to non-physiological levels of substrate not infrequently leads to inhibition of the metabolic pathway under study. As yet there is no entirely satisfactory answer to this problem. The best solution would be a sensor that detected changes in the level of substrate and triggered a mechanism for restoring its level. Such devices exist for 0, and for glucose, but they are not generally available for other metabolites. Substrate levels can however be held constant by perifusion techniques (Section 12.8), but these too have a number of limitations. The volume of the incubation medium is also important in regard to the rate of production of certain metabolites that tend to accumulate to a steady state (Section 6.4.1). When hepatocytes are incubated with glucose at relatively high concentrations (> 20 mM) a steady-state level of lactate in the medium is reached in 60-80 min, if a standard incubation mixture of 30 mg dry weight of cells in 2 ml of medium is employed (Fig. 6.2). If however the same mass of cells is incubated in 4 ml of medium, the approach to steadystate takes substantially longer, because the cells have to accumulate twice as much lactate in the medium to achieve the steady-state concentration, Finally, it should be emphasized that when establishing an incubation system for an investigation that has not been carried out previously, it is desirable to examine the effects of cell dilution on the metabolic process under study. There is experimental evidence that rates of metabolism may vary considerably as a function of the cell concentration, even in the absence of any approach to steady-state (Jurin and McCune, 1985).

Ch. 6.


155

6.4.3. Incubation media

Balanced saline media, such as Krebs-bicarbonate saline (Krebs and Henseleit, 1932), that have been used for many years for incubation of surviving slices, or for liver perfusion, are also suitable for hepatocyte incubation. The hepatocytes tolerate physiological concentrations of Ca2+and, as has been previously noted, the presence of these ions is essential for optimal metabolic performance. However, if the permeability of the plasma membrane has been altered to allow Ca2+penetration, there will be much more damage induced in the presence of Ca2+than in their absence (Berry, 1962). A considerable difference of opinion appears to exist regarding the choice of a buffering agent. As has been discussed in Section 2.4.7, our preference is bicarbonate, because together with a gas mixture containing 5% CO,, it represents the physiological buffering system of the blood. However, it should be noted that the special properties of this buffer in the intact mammal depend on the fact that the concentrations of either or both components can be altered physiologically, something certainly not easy to do in the incubation vessel. Therefore, the bicarbonateKO, system is by no means a perfect buffer in vitro. Bicarbonate solutions should be gassed with carbogen to bring the pH to 7.4 prior to addition of the cell suspension. If marked changes in pH are expected as a consequence of the experimental circumstances, a supplementary buffer such as phosphate, or even a synthetic buffer such as HEPES, is desirable. Many metabolic processes, including gluconeogenesis or urea synthesis, require the presence of CO,, but in our experience a level of 5% in the gas phase is not necessary, 1-2% proving adequate for glucose synthesis. Once again, the best approach is to invest some time in exploratory studies before making a final choice. 6.4.4. Addition of albumin to incubation media

Another question that frequently arises is whether or not albumin

156


should be included in the incubation medium. There are many obvious disadvantages in regard to albumin use. These include expense, uncertainty in relation to composition and purity, the extra work associated with achieving preparations free of fatty acid (Chen, 1967) and the presence of much greater precipitate in the PCA-treated medium, which tends to reduce the volume of extract available for analysis. For these reasons it would seem appropriate to avoid inclusion of albumin in the medium, and for experiments in which cells will not be exposed to noxious agents, this is the recommended approach. However, if toxic agents are to be used, the presence of at least 1% albumin is highly desirable. Although this subject has not been explored in depth, it seems likely that damaged cells themselves release toxic products into the incubation medium. Albumin appears to be an effective sink for such substances. Whatever the explanation, empirical observation indicates that in the presence of a noxious agent, cells incubated in a medium containing albumin remain apparently intact much longer. Long-chain fatty acids are water-insoluble but are solubilized by complexing with albumin (see Protocol 6.6). Hence for studies of their metabolism, the presence of albumin is essential. A final albumin concentration of 2-2.5‘% is appropriate for a palmitate concentration of 2 mM. While fatty acids of C, or below are soluble in water, they qualify as possible toxic agents, so that inclusion of albumin in the medium is recommended. Because much of our metabolic work involves the study of fatty acids or inhibitors, we routinely include 2.25% BSA in the incubation mixture. Commercial albumin, even when designated ‘fatty acid-free’, frequently contains measurable quantities of fatty acids. The method of Chen (1967) is recommended for their removal. 6.4.5. Addition of gelatin to incubation medici

Gelatin was originally included in incubation media by Zahlten et al. (1973) to replace albumin, which seemed to inhibit gluconeogenesis. The inclusion of 1.5%;, gelatin was thought to provide viscosity and

Ch. 6.


I57

protection of cells, but no experimental evidence was presented to support these ideas. Some researchers still include gelatin in media. especially when long incubations are involved, but it is not clear that it is necessary. Preliminary results in our laboratory indicate that its addition to media slows the binding of hepatocytes to the glass walls of incubation vessels and may increase the rate of gluconeogenesis from added lactate in cells from fasted rats. On balance, however, no clear cut advantage of gelatin over albumin has been established experimentally. Gelatin has frequently been used in studies of hormone action (see Chapter 8). 6.4.6. Osmolulity of incubation mediu

For most studies it is desirable to maintain the osmolality of incubation media within the physiological range (280-300 mmol/kg). However, the addition of substrates may change the osmolality of the basic incubation media. I t is possible to compensate for this by omitting or including an equimolar amount of NaCI. We use the following incubation conditions. Cells are suspended in washing buffer (Protocol 2.1 (iv)). BSA preparations are dialysed against a solution containing in 1 I, 6.43 g NaCI, 0.78 g KCI, 0.39 g MgS0,.7H20 ( 1 10 mM Na+, 120 mM CI-, 10.4 mM K', 1.6 mM Mg2+,1.6 mM SO:-). A 2-ml standard incubation mix comprises:

ml

Hepatocyte suspension 0.2 M Na phosphate buffer (pH 7.4) 0.08 M CaCI, 1 M N~HCO, 0.4 M NaCl 9% BSA HI0

I .ooo 0.050 0.050 0.050 0.075 0.500 0.275

158


The volume(s) of H,O and NaCl are adjusted to allow for substrate additions. The final concentrations of ions (mM) in our basic incubation are: 140 Na+, 5.4 K+,0.8 Mg2+,2.6 Ca2+, 11.0 phosphate, 0.8 SO:-, 103 CI-, 25 HCO;. This medium has an osmolality of 290 mmol/kg. Various groups (e.g. Baquet et al., 1990; Haussinger et al., 1990) have demonstrated that changes in cell volume may alter metabolic rates, but the osmolalities that have been used to swell the cells lie well outside the physiological range. 6.4.7. Use of inhibitors

Inhibitors are frequently used in studies of the metabolic behaviour of hepatocytes. One procedure in such studies is to use the lowest concentration of inhibitor that gives the maximal inhibition of the process under study. In the course of our experiments, we have found that much more information can be gained by using a wide range of inhibitor concentrations, from the smallest level that will cause a detectable inhibition, to the lowest amount that brings about maximal inhibition. The hepatocyte suspension is well-suited to this type of experiment, in view of its homogeneity and ease of handling. Our own studies suggest that the inhibitory action of low concentrations of some antibiotic inhibitors, such as rotenone and antimycin, may decline during the experimental period, particularly when the cells are from well-fed animals, which have a high glycolytic capacity.

6.4.8. Solubility of experimental substances A problem that frequently arises in regard to the use of inhibitors or other experimental agents is that many of these substances are not water soluble. They must therefore be delivered to the incubation vessel in a non-aqueous form that nevertheless allows their uptake by the isolated hepatocytes. One possibility is the use of a relatively polar

Ch. 6.


159

solvent that is miscible with water. By far the most commonly used agent for many types of biological preparation is ethanol, but this is unsuitable for studies with isolated hepatocytes. Ethanol is rapidly metabolized by hepatocytes, and the metabolism of ethanol has profound effects on hepatic intermediary metabolism. On the assumption that the smallest quantity of ethanol that can be accurately pipetted is 5 pl, it can be calculated that the delivery to the medium of an agent dissolved in this solvent will result in the addition of almost 100 pmol of ethanol. Two alternative solvents that are metabolized much less readily than ethanol are acetone and dimethylsulphoxide (DMSO). However, as is the case with ethanol, substantial quantities of these agents will be delivered to the incubation mixture together with the substance under test. Our experiments have shown that both acetone and DMSO have significant effects on hepatocyte metabolism, and their use as solvents is not recommended, if they are to be added directly to the incubation medium. Fortunately, there is another approach that experience has shown to be more satisfactory. Advantage can be taken of the volatility of acetone to remove it from the incubation vessel. The substance under study is taken up in acetone and a measured amount is delivered to the empty and dry incubation vessel. The vessel is now placed in a fume hood at room temperature and the acetone evaporated. The remainder of the components of the incubation medium and the isolated hepatocytes are then added. It may seem surprising that isolated hepatocytes are capable of taking up substances added in this way, but our experience with various inhibitors such as rotenone, antimycin and oligomycin confirms this to be the case. 6.4.9. Availability of substrates to isolated hepatocytes

The ability of isolated liver cells in suspension to metabolize an added substrate will depend not only on the existence of appropriate intracellular enzymes for its metabolism, but also on the ability of the substance under study to penetrate the plasma membrane. Thus though

160


the liver is rich in enzymes of the Krebs tricarboxylic acid cycle, intact isolated hepatocytes fail to metabolize cycle intermediates added to the incubation medium. Nevertheless, some metabolism of added cycle intermediates can be expected, since even the best preparation of isolated liver hepatocyes is likely to contain damaged cells. These damaged hepatocytes have lost the ability to exclude dicarboxylic or tricarboxylic acids and hence are able to metabolize them to a significant degree. This difference in permeability properties between intact and damaged cells forms the basis for a method for measuring the percentage of damaged cells in a hepatocyte suspension (Mapes and Harris, 1975), in which the response of hepatocytes to added succinate is used to determine the degree of damage (Protocol 4.1). On occasion the inability of hepatocytes to metabolize an added substrate may reflect the lack of a key enzyme in the metabolic pathway. A good example of this is the pathway of leucine degradation. Minimal oxidation of leucine occurs when the amino acid is added to a suspension of hepalocytes. On the other hand the corresponding keto acid formed by the action of leucine aminotransferase (in muscle) is rapidly oxidized by hepatocytes (Williamson et al., 1979). Since leucine is readily incorporated into hepatic protein, the amino acid undoubtedly crosses the plasma membrane. Hence the lack of oxidative metabolism of leucine in the liver is apparently due to a low level of the aminotransferase rather than to a permeability barrier to the amino acid (Ichihara and Koyama, 1966). Protocols given in this chapter provide a mechanism for determining whether or not a putative substrate can enter the isolated hepatocyte.

6.5. Measurement of cellular 02-uptake Relatively few studies of hepatocyte metabolism include the measurement of 0,-uptake, yet in our experience this measurement can yield

Ch. 6 .


161

valuable insights into metabolic behaviour, as well as providing a continuous indicator of cellular integrity through the course of the experiment. A possible reason for the relative lack of enthusiasm for measuring 0,-uptake is the perception that such measurements are difficult and require complex equipment. There is some degree of truth in this, but we consider that the value of the information that can be obtained warrants the necessary effort. 6.5.1. Polurographic measurement qf’ cellulur 0,-uptake

The simplest technique for measuring 0,-uptake involves use of a polarographic apparatus. Systems are available commercially, comprising a thermostatted water bath into which four incubation chambers can be inserted (Yellow Springs Instruments). A portion of the cell suspension is placed in one of the chambers that is then sealed with a plastic plunger, through the body of which is inserted an 02-electrode (Fig. 6.3). The system is easy to calibrate and simple to use, but it has a number of minor disadvantages. The incubation chamber requires continuous stirring, to allow 0, to diffuse freely, so the design includes a magnetic stirring system. The stirring bar is liable to inflict damage on the cell suspension, unless it possesses a small girdle that raises the bar from the base of the chamber. The incubation medium should be gassed to achieve maximal 0, saturation before the cells are added in a relatively small volume. It is undesirable to gas the medium by bubbling gas through it once the cells have been added, because of the likelihood of damage induced as the gas bubbles pass rapidly through the medium. Even when the incubation medium is thoroughly oxygenated by gassing, the rate of 0,-uptake of 15 mg dry weight cellsiml is often sufficient to deplete the medium of 0, within 10 min. It is possible to extend this period by decreasing the cell concentration. However, this often results in metabolite levels that are at the limits of detection, and metabolic rates may be changed by dilution effects (Jurin and McCune, 1985). The cost of the equipment prohibits the running of multiple channels and therefore

162


limits the informatior. that can be obtained. Nevertheless, for shortterm and relatively simple experiments the polarographic system is ideal. A simple mechanism for re-oxygenating an 0,-depleted oxygraph vessel without dismantling the plunger and 0, electrode makes use of a loop of fine silastic tubing. The tubing is submerged in the incubation medium and its ends attached to cannulas which protrude from the vessel (Fig. 6.3). Gas may be passed through the tubing, which is readily permeable to 0,, thereby raising the 0,-tension of the incubation medium. 6.5.2. Manometric measurement of cellular O p p t a k e Manometry has been a useful tool for the measurement of tissue respiration in vitro since the time of Warburg. The early respirometers required accurate calibration, but more recent versions are designed to avoid the need for this. Some sophisticated pieces of manometric apparatus are presently available, including the differential respirometer (Gilson, U.S.A.) and the automated Warburg system manufactured by Plischke and Buhr K.G., which employs pressure transducers. These apparatus usually accommodate up to twenty vessels and hence enable a much greater number of experimental conditions to be explored. Thus, in contrast to polarographic techniques, manometric methods enable 0,-consumption to be measured for a large number of samples, at relatively constant pressures of 0,, over extended incubation periods (30-120 min, depending on the incubation conditions). However, certain limitations in their use will be encountered. Because of the sensitive nature of manometry, the vessels must equilibrate completely with the surrounding bath before accurate readings can be taken. Any subsequent alteration of environmental conditions will inevitably lead to inaccurate readings. The requirement for equilibration demands that the cells be incubated for at least 10 min before the first reading can be made. In the case of the polarograph a warm-up time of only 2 min is necessary.

Ch. 6.

BIOCHEMICAL PROPERTIES CROSS SECTION

Fig. 6.3. 0,-electrode assembly

163

164


The use of manometric techniques to measure cellular 0,-uptake is complicated by the production of CO, by animal cells. In standard methods, the CO, generated is trapped in KOH contained in a centre-well within the incubation vessel. If such methods are used the gas phase must be devoid of CO, and the medium free from added bicarbonate, This non-physiological system can be satisfactory for some types of studies, but is of limited use. The utility of the manometric approach has been greatly increased by the introduction of trapping agents for CO, that maintain a pC0, of about 40 mmHg in the gas phase of the incubation vessel. These ‘CO, buffers’ were first introduced by Pardee ( 1949)and contained diethanolamine, which reversibly binds to CO,. Subsequently, Warburg and Krippahl (1 959) modified the technique, including carbonic anhydrase in the C 0 2 buffers. The buffers permit the accurate measurement of 0,-consumption by maintaining physiological concentrations of C 0 2 in the medium and gas phase and are recommended when metabolic studies involving CO, fixation, such as gluconeogenesis or urea synthesis are being carried out. As descriptions of the technique have not been published in readily-available journals, an outline is given here. 6.5.3. Apparatus

Manometry can be carried out using a standard Warburg constant volume respirometer or a Gilson differential respirometer. Incubations are performed in a special manometer vessel (Fig. 6.4), consisting of a large centre-well, a main compartment and a side-arm. The centrewell is constructed with an inverted lip to reduce the possibility of spillage of the alkaline CO, buffer into the main reaction compartment. 6.5.4. Preparation and composition of CO, bujyer The CO, buffer is prepared daily from 3 M KHCO, and 3 M K,CO, in the ratio 3:l (v/v), a mixture that is at equilibrium with an at-

Ch. 6


I65

snorkel

side a n inverted lip centre well main comDartmenl

Fig. 6.4. Manometer vessel for measuring 0,-uptake in the presence of CO,.

mosphere of 5% CO,. The attainment of equilibrium is facilitated by the addition of carbonic anhydrase. The presence of the enzyme is essential because the buffering reaction

CO,

+ H,O + C0:-

-

2 HCO;

is too slow in its absence. The solution, once prepared, is stored at room temperature in a stoppered flask with an atmosphere containing 5% CO,. Some shaking and gentle warming is required to make up stock solutions of 3 M K,CO,. The K,CO, should be derived from a source that has been dried to constant-weight. The stock solutions must be stored in well-sealed containers. It has been recommended that the two components can be mixed and stored for several weeks at room temperature in a well-stoppered bottle with little air space, and that the buffer containing carbonic anhydrase can be prepared on a weekly basis (Krebs et al., 1974). However, in our hands, better results are obtained by mixing the two components of the buffer, and adding the enzyme, on the day of the experiment. We have obtained more consistent results by using a commercial source of carbonic anhydrase. However, the commercial preparation

I66


is sufficiently expensive to make preparation of the enzyme in crude form worthwhile (Tashian et al., 1966). An alternative means of reducing costs, at least for experiments where no radiolabelled compounds yielding I4CO, are present, is to re-use the CO, bufferkarbonic anhydrase mixture. This procedure succeeds only if there is no precipitation of the buffer components, so it is essential that the mixture is removed from the centre-well before the incubation vessel is stood on ice. We have established that 0.5 mg of lyophilized carbonic anhydrase (Boehringer Mannheim, Catalogue no. 103 187) in 2 ml CO, buffer is adequate to maintain linearity over a wide range of rates of 0,-consumption for a standard 40-min incubation period. 6.5.5. Incubation procedure

The main compartment is loaded with substrates, albumin, salts and any other experimental agents to a total volume of 1 ml, as described in Section 6.4. and the vessels stood on ice. Immediately before loading the cell suspension (1 ml) into the outer compartment, 2 ml of the pregassed CO, bufferkarbonic anhydrase mixture is added to the centrewell of each vessel. The vessels are transferred to the respirometer, gassed with carbogen for 5 min, equilibrated to bath temperature for a further 10 min and then incubated for the desired period during which measurements of 0,-uptake are made. Deviation from linearity of 0,-consumption during the early stages of recording normally indicates poor equilibration or faulty preparation of the bufferlenzyme mixture. At the completion of the incubation period, the length of which depends on the nature and amount of the substrate added, the pressure in the respirometer is equalized, the vessels removed and the main compartment acidified with an equal volume of ice-cold I M PCA. Further treatment is according to Protocol 6.2. 6.5.6. Measurement of radiolabelled carbon dioxide formation

In experiments where a I4C-labelled substrate has been added, it is

Ch. 6.


I67

relatively straightforward to determine the amount of radiolabelled CO, produced. It has been shown that 96”h of the I4CO, generated equilibrates with the buffedenzyme mixture (Krebs et al., 1974). At the completion of the experiment and before acid is added to the main compartment, a sample (0.5 ml) of the mixture is removed from the centre-well and transferred to a scintillation vial for counting. 6.5.7. Measuring 0,-uptake with an oxystat system

The majority of studies with isolated hepatocytes are conducted with a gas phase containing 95% 0,, representing a PO, of about 680 mmHg. A number of workers have expressed concern that this 0,-tension is unphysiological and have recommended that a tension commensurate with the content of 0, in air be employed, i.e. approximately 150 mmHg. In fact, the actual range of PO, values within the liver varies between 1 and 60 mmHg with a mean around 20 mmHg (Kessler et al., 1984). Nevertheless, there is no evidence that the metabolism of isolated hepatocytes differs whether they are exposed to an 0,-tension of 680 or 150 mmHg. For some experiments it may be desirable to simulate 0,-tensions actually encountered in liver. De Groot and co-workers (No11 et al., 1986; de Groot et al., 1988) have developed an oxystat system suitable for this purpose which can maintain constant 0,-tensions as low as 0.01 mmHg. Studies with the system have shown that isolated hepatocytes from fed rats can tolerate a PO, of only 0.3 mmHg for at least 4 h without increased staining with trypan blue, cells from fasted animals being only slightly less tolerant (Anundi and de Groot, 1989). However, there is a marked decrease in ATP levels in hepatocytes derived from fasted rats and exposed to PO, levels of 0.3 mmHg. The oxystat system has obvious merit for studies of the effects of hypoxia on hepatocytes, and has also proved of value in determining the levels of pOz that promote free radical generation and lipid peroxidation in the presence of toxic agents such as carbon

168

IS0 LATED H EPATOCY TES

tetrachloride (de Groot et al., 1988). However, the method has a drawback in that there is an inevitable dilution of the incubation medium by the 0,-carrying solution. A possible means of overcoming this is to use fluids with a higher 0,-content than can be achieved with saline. Whereas haemoglobin solutions prove unsatisfactory, the possibility of using fluorocarbons (Faithful1 et al., 1988; Tabayashi et al., 1988; Ballatori et al., 1989) or dilute hydrogen peroxide as the 0,-carrier remains to be fully explored.

6.6. Metabolic activities Isolated hepatocytes have been used to study many of the metabolic activities known to occur in liver, and it is not feasible to describe these in detail. A number of these activities such as gluconeogenesis, lipogenesis and ureogenesis require the presence of bicarbonate in the incubation medium and a gas phase containing CO,. 6.6.1. Gluconeogenesis

Measurement of the rate of glucose synthesis (Jg,,,os,)is frequently carried out during the study of metabolic regulation. However, it is also of considerable value as an indicator of cellular integrity. Fasting rates of glucose formation of 0.5-0.7 pmol/g wet weight per min from lactate or 1.3-1.5 pmol/g wet weight per min from lactate plus palmitate are suggestive of healthy (rat) hepatocytes. Gluconeogenic rates are best measured in cells prepared from livers derived from rats fasted for 18-24 h to deplete hepatic glycogen. An indication of Jglucose by cells derived from fed rats can be made by use of [I4C]lactate but, due to isotope exchange, the rate determined may prove to be a considerable underestimation. Numerous other substrates have been used as glucose precursors in the study of gluconeogenesis in isolated hepatocytes. A number of studies have employed inhibitors of gluconeogenesis such as

Ch. 6.


169

quinolinate, fluorosuccinate or amino-oxyacetate (Gregory and Berry, 1989). 6.6.2. Glycolysis

Although glycolysis in hepatocytes has received far less attention than has gluconeogenesis, it is an area well worthy of investigation. A difficulty with such studies is that the pyruvate formed from glycogen or glucose is likely to be further metabolized. Hence, the quantifying of glycolytic flux may prove difficult. Much of the earlier work on glycolysis utilized short-term incubations, and so it has only recently been recognized that the rate of glycolysis as measured by lactate accumulation (JlactaJ declines with time, frequently to zero (Fig. 6.2). Hence, accurate determination of maximum Jlactale requires measurements to be carried out within 10 min of commencement of the incubation. In the fed state, endogenous glycogen is an important precursor of lactate (and glucose). However, if glucose is not added to the incubation medium, the rate of glycogen degradation to glucose is considerable. In a typical incubation, some 50% of the glycogen will be lost within 45 min under control conditions, while the presence of an activator of glycogen breakdown may lead to the total loss of glycogen within that time. Glycogen is absent from hepatocytes obtained from fasted rats, and glycolysis must be studied by incubation of hepatocytes with glucose. Because of the high K , of hexokinase D (glucokinase) for glucose (Reyes and Cardenas, 1984) concentrations of the hexose of at least 20 mM are required to induce substantial glycolysis, and maximal rates for Jlilc,ate are not reached unless cells from fasted animals are incubated in a medium containing over 80 mM glucose. 6.6.3. Glycogen synthesis

Whereas the liver in vivo can synthesize glycogen at blood glucose levels of 10 mM glucose, in vitro preparations require the presence

170


of a much higher glucose concentration before net glycogen synthesis takes place. Isolated hepatocytes are no exception, and our unpublished observations indicate that the rate of glycogen synthesis by hepatocytes from fasted rats is dependent on the concentration of glucose in the incubation medium. The maximal rate of glycogen synthesis observed (0.74 pmol/min per g wet weight at 80 mM glucose) is close to that reported for hepatic glycogen synthesis under optimal conditions in vivo (Devos and Hers, 1979; Boyd et al., 1981; Youn et al., 1987). 6.6.4. Ureogenesis

The precursor of urea may be an amino acid such as alanine or an amine (e.g. glutamine). However, the most rapid rates of urea synthesis are induced by ammonium chloride added at supraphysiological concentrations of 10-20 mM. Indeed relatively short-term experiments, lasting no more than 30 min may be required to preserve a sufficiently high level of ammonia to maintain maximal rates of ureogenesis under conditions where other metabolites are in sufficient supply (cf. Wiechetek, 1986). Under these conditions, ornithine is limiting (Meijer et al., 1975; Stubbs and Krebs, 1975; Wojtczak et al., 1978) and, as originally noted by Krebs and Henseleit (1932), its addition (2-10 mM) substantially stimulates the rate of ureogenesis. If there is a deficiency in the supply of precursors of oxaloacetate, as in the fasted rat, the generation of aspartate is also a rate-limiting factor (Meijer et al., 1975, 1978; Briggs and Freedland, 1976). Krebs et al. (1979b) demonstrated that the provision of lactate (2 mM) stimulated urea synthesis from ammonia three-fold in cells isolated from fasted, but not from fed, rats. When 2 mM ornithine was present however, the addition of lactate also increased urea synthesis in fed rats. In the fasted state, the presence of an exogenous energy source such as oleate is required to achieve maximal rates of ureogenesis (Krebs et al., 1979b and our own observations). As might be expected, diet plays a major role in determining the activity of the urea cycle, maximal rates of urea

Ch. 6.


171

production being found when rats are fed high protein diets such as 70% casein (Briggs and Freedland, 1977). 6.6.5. Lipogenesis

Hepatocytes from fed rats are capable of rates of lipogenesis equal to those found with perfused liver. However, whether or not a lipogenic substrate is added, it is important that the cells be replete with glycogen. To ensure this, it is often desirable to resort to ‘meal feeding’ (Clark et al. 1974). Measurement of lipogenic rates is frequently carried out by determining the incorporation of 3H,0 or [I4C]acetateinto cellular lipid. Bottomley and Garcia-Webb (1987) compared lipogenesis in isolated hepatocytes using both of these compounds and found significant differences both quantitatively and qualitatively. Lipogenesis was greater when measured by 3H20incorporation than with [14C]acetate, irrespective of the exogenous metabolites supplied. One likely explanation for this is dilution of [I4C]acetate by endogenous sources of acetate or acetyl carbon. Apart from incubations in which aspartate and glutamate were added, where insulin actually inhibited lipogenesis as measured by 3H,0 incorporation, this hormone stimulated lipogenesis. This effect was, however, greater with [I4C]acetate than with ’H20. These results demonstrate that care must be taken when comparing results using these two tracers or when choosing a particular isotope and set of conditions for use in a particular study. For measurement of the rate of lipogenesis from endogenous substrates, the use of 3 H 2 0is essential. There appears to be a lag period of up to 30 min before maximal rates of lipogenesis are achieved (unpublished observations). 6.6.6. Lipid oxidation and ketogenesis

Hepatocytes oxidize short, medium and long chain fatty acids, the main end-products generally being 3-hydroxybutyrate and acetoacetate (frequently termed ‘ketone bodies’), although some CO, is generated via

172


the Krebs cycle. The short chain fatty acids (C,-C,) are somewhat soluble in water and concentrations of sodium butyrate as high as 10 mM can be obtained. Even octanoate is sufficiently soluble that concentrations of 4 mM can be achieved. However, all these fatty acids show some detergent properties, and experience leads us to recommend that albumin (2.25%) be included in the incubation medium when fatty acids are added substrates. With fatty acids of chain length greater than C,, the presence of albumin is essential (see Protocol 6.6). The use of albumin to bind fatty acids is not as straightforward as it might seem. Many commercial preparations of albumin, even those labelled 'fatty acid free', can contain substantial quantities of fatty acid, which need to be removed if the fatty acids under study are to be bound. We use the method of Chen (1967) to accomplish this. The rate of ketone body formation from fatty acid is dependent on free substrate concentration and declines with time. It is greatly influenced by the presence of other substrates such as lactate, glucose or fructose, which are quite strongly antiketogenic. In the absence of other exogenous substrates, cells from fasted animals oxidize fatty acids almost entirely to ketone bodies (&oxidation).

Protocol 6.6 Solubilization of long-chain futty acids (i) Weigh into a scintillation vial sufficient fatty acid to yield 20 ml of an 8 mM solution (e.g. 41 mg of palmitic acid). (ii) Slowly melt the fatty acid and neutralize with 1.6 ml of pre-heated 0.1 M NaOH. If there are still droplets of fat on the surface of the liquid, slowly and carefully add 1 M NaOH drop by drop until they disappear. (iii) Warm 18.4 ml of 9%)fatty acid-free BSA solution to 45-50°C, stirring constantly. Pour in the hot neutralized fatty acid. Take care. BSA denatures around 55°C. (iv) Stir for a few seconds to allow the palmitate to bind to the BSA, cool and adjust the pH to 7.4. (v) Filter through Whatman No. 1 paper. If the solution is cloudy, the fatty acid has not completely bound to the albumin and the procedure must be repeated with fresh reagents.

Ch. 6.


173

(vi) Store the BSA-bound fatty acid solution frozen in 20-ml aliquots. 6.6.7. Protein synthesis

Several difficulties arise when protein synthesis is studied in isolated hepatocytes. Since proteolysis is occurring simultaneously, the specific activity of any labelled precursor amino acid will be constantly changing during an incubation period (Mortimore et al., 1972; Seglen and Solheim. 1978; Rannels et al., 1982). Complications also arise because the intracellular amino acids do not completely equilibrate with the extracellular pool and, furthermore, there appear to be at least three separate intracellular amino acid compartments (Khairallah and Mortimore, 1976; Rannels et al., 1982). The branched-chain amino acids, valine and leucine, are most commonly used as precursor amino acids in liver protein synthesis studies. They are not synthesized by hepatocytes and their rates of catabolism are not significant. Both amino acids are commercially available in high purity and specific activity, and can be easily assayed. It seems that amino acids bound to t-RNA (the immediate intracellular precursor pool for proteir. synthesis) are preferentially, although not completely, derived from extracellular sources (Airhart et al., 1974, 1981). At concentrations of 5 mM or higher, it is possible to equate the extracellular specific activity of the amino acid with that of the aminoacyl-t-RNA pool (Khairallah and Mortimore, 1976; Flaim et al., 1982a) and ignore dilution of the isotope by endogenous proteolysis. The elevated concentrations of valine or leucine added do not alter rates of protein synthesis (Khairallah and Mortimore, 1976; McNurlan et al., 1979; Flaim et al., 1982a,b). Generally, label incorporated into acid-precipitable material is used as a measure of total protein synthesis. However, by using ratspecific antisera, individual proteins can be immunoprecipitated and their production and secretion studied (Koj et al., 1984; Fries and Lindstrom, 1986). Experimental protocols must be designed specifically to take into account the existence of two populations of proteins with markedly different average rates of turnover ( t , , ? of approximately 10

174


min (short-lived) and 40 h (long-lived), respectively). Wheatley (1984) found short-lived protein turnover accounted for one-third of protein turnover but that these proteins contribute only 0.6% of total liver protein. As well, it is thought that short-lived proteins may be acidsoluble peptides which are not acid-precipitated with general liver proteins (Solheim and Seglen, 1980). During the first 60 min of incubation at 37°C in unsupplemented media, protein synthetic rates in isolated hepatocytes are less than those observed in vivo. Supplementation with greater than physiological concentrations of amino acids or other substrates as energy sources increases protein synthesis (Seglen and Solheim, 1978; Flaim et al., 1982a). However, protein synthetic rates also increase during longer periods of incubation at 37°C in unsupplemented media (Woodside and Mortimore, 1972). Hormones also influence synthetic rates (see Chapter 8). For example, glucocorticoidsdecrease and insulin increases protein synthesis in cultured and isolated hepatocytes (Koj et al., 1984; Brostrom et al., 1986). Protein synthesis is pH-dependent with leucine incorporation into protein being depressed 30% by lowering the pH of the media from 7.4 to 7.0, or increased by about the same amount by raising the pH to 7.8 (Beloqui et al., 1986). Ethanol oxidation decreases protein synthesis (Mqrland et al., 1979) possibly by decreasing cytosolic pH (Beloqui et al., 1986). Hasselgren et al. (1988) have comprehensively reviewed methods of studying protein synthesis and degradation not only in isolated hepatocytes, but also in liver slices, perfused livers and in vivo. They detail other non-invasive techniques, methods of detecting secretory and non-secretory proteins, and influences Kupffer cells may have on hepatocyte protein synthesis. 6.6.8. Proteolysis

During early starvation (24-48 h), liver is a major source of amino acids for gluconeogenesis and other metabolic processes. However,

Ch. 6.


I75

even in the fed state, proteolysis is a continuous process within the liver, some proteins being turned over much more rapidly than others. Amino acids and hormones have a regulatory role in the proteolysis of long-lived (but less effect on short-lived) proteins. When hepatocytes are incubated in the absence of amino acids, or in the presence of glucagon, high rates of protein breakdown (&5% per h) of total intracellular protein, can be observed (Seglen et al., 1980a; Schworer et al., 1981). Most of this proteolysis occurs in the lysosomes due to autophagy (Mortimore and Poso, 1984; Seglen, 1987) which may specifically target some proteins (McElligott et al., 1985). Addition of certain, but not all, amino acid combinations can inhibit the lysosomal, but not the extralysosomal, component. Most effective are combinations of leucine with either alanine, asparagine or glutamine, added at concentrations of 5 mM (Seglen et al., 1980a; Car0 et al., 1989). However, when added singly, these amino acids are relatively ineffective. The production of free NH, from glutamine may decrease proteolysis rates by increasing intracellular (and intralysosomal) pH which directly inhibits lysosomal protein degradation (Poso et al., 1982, 1986). Glucagon and P-agonists stimulate proteolysis (Schworer and Mortimore, 1979) whereas insulin and other growth promoting factors inhibit (Ballard and Gunn, 1982). As in studies of protein synthesis, branched-chain amino acids are often used to measure long-lived protein degradation. However, unless allowance is made for re-utilization of these amino acids for protein synthesis, low rates of proteolysis are obtained. When rates of release of unlabelled valine or leucine are used to measure long-lived protein degradation, cycloheximide may be added to the incubation to inhibit protein synthesis. Unfortunately, after 10-1 5 min cycloheximide also inhibits proteolysis. Short-lived protein turnover rates must also be subtracted. Another approach involves prelabelling intracellular protein with ['4C]valineseveral hours before killing the experimental animal. Contamination by short-lived protein degradation can be eliminated by pre-infusion with unlabelled valine before measuring [I4C]valine

176


release from hepatocytes into the incubation medium. Isolated hepatocytes are incubated in the presence of 1 5 mM unlabelled valine to minimize re-incorporation of [I4C]valine(or cycloheximide can be used). In contrast to protein synthesis, leucine cannot be used in this technique as a high concentration of this amino acid inhibits proteolysis. The degradation of labelled RNA (Lardeux et al., 1987) and the volume of intracellular macroautophagic vacuoles (Schworer et al., 1981) have also been used as measures of protein degradation. Microinjection (Freikopf-Cassel and Kulka, 1981) and electropermeabilization (Seglen and Gordon, 1984) (Section 12.7.4) are other techniques used to study proteolysis. Because of the high rate of production of amino acids in isolated hepatocyte suspensions, large amounts of amino acids will accumulate in the incubation vessels, especially at high cell concentrations, and can affect several metabolic pathways. For example, gluconeogenesis from lactate is accelerated by amino acids derived from proteolysis (Crabb et al., 1980). As well, proteolytically derived arginine can supply ornithine for the urea cycle. The requirement for ornithine in urea synthesis from added ammonia is due to the fact that ammonia, a weak base, raises the intralysosomal pH and inhibits lysosomal proteolysis and thus arginine production (A.J. Meijer, personal communication). Mortimore et al. (1989) have recently reviewed the mechanism and regulation of proteolysis.

6.7. Transport activities 6.7.I . Transport across the sinusoidal membrane

The hepatocyte plasma membrane contains numerous transport systems for ions and metabolites, and many of these have been demonstrated with isolated hepatocytes. The liver cell appears freely permeable to small neutral molecules (Alpini et al., 1986), but many

Ch. 6.


I77

charged species require specific transporters. Measurement of the kinetics of transport systems is straightforward, but can be technically demanding when uptake of the intermediate under study becomes non-linear within seconds. Usually, radiolabelled metabolites are employed and this generally allows the degree of their uptake to be determined after an incubation time of 1 min, the measurement being carried out after cell centrifugation as when the intracellular concentration of Na' or K+ is measured (Protocol 6.4). Alternatively, the cells may be centrifuged through silicone oil (Protocol 6.5), or the suspension may be diluted with a large volume of washing medium at 0 4 ° C . 6.7.2. Secretion of biliary components

Within a relatively short time after isolation hepatocytes lose their polarity and the biliary region of the plasma membrane can no longer be identified (Chapter 5). Despite these morphological changes, the hepatocytes remain capable of biliary transport as demonstrated by their ability to secrete glutathione conjugates into the medium. Oude Elferink et al. (1990a) have demonstrated the conjugation of a model by isolated hepatocytes organic anion, 1-chloro-2,4-dinitrobenzene, and the subsequent secretion of dinitrophenyl glutathione into the medium. From studies with intact and Tr- mutant rats, they infer that the anion transport takes place across the canalicular membrane of the hepatocyte. The Tr- rat has a greatly impaired capacity to secrete organic anions such as conjugated bilirubin or dibromosulphthalein, or sulphated or glucuronidated bile acids, whereas the secretion of taurocholate and cholate is normal. This failure to secrete certain organic anions is believed to be the consequence of a defect in the canalicular transport mechanism. The secretion of dinitrophenyl glutathione is inhibited by the 3-sulphates of glycocholate and taurolithocholate, and even more strongly by the 3-0-glucuronides of cholate and lithocholate (Oude Elferink et al., 1990b). These workers postulate that the canalicular

178


membrane contains a multispecific organic anion transporter that also mediates the transport of oxidized glutathione (GSSG).

6.8. Analysis of radiolabelled compounds The isolated hepatocyte preparation lends itself well to the study of the transport and metabolism of radiolabelled compounds. For analysis of these substances, commercially avalilable liquid scintillation cocktails (e.g. ACSII, Amersham) are recommended where aqueous samples are involved. Whenever isotope has been incorporated into hepatocytes, it may be necessary to count cell pellet samples. Allowance for isotope associated with the entrained medium in the pellet must be made by including a radiolabelled inulin derivative in the incubation (Protocol 6.3). To prepare such material for counting, the pellet (up to 10 mg dry weight of hepatocytes) is solubilized using 0.5 ml of tissue solubilizer (NCS, Amersham) at 5 50°C until the sample is homogeneous. After solubilization, the digest should be neutralized to a pH of 6.7 with glacial acetic acid (0.034 ml/ml NCS) to minimize chemiluminescence.(The brochure provided with the NCS gives additional means of decreasing these spurious counts). The samples are counted after the addition of 5-10 ml of ACSII.

CHAPTER 7

Utilization of hepatocytes for drug studies

7. I. Introduction The liver is the chief organ involved in the metabolism of xenobiotics (foreign compounds), a wide variety of which are t.aken up by hepatocytes and converted to pharmacologically inactive, active or sometimes toxic metabolites. The complexity of these processes, involving both endogenous and exogenous factors, has encouraged investigators to turn to simpler models than the whole animal for the study of drug metabolism and toxicity. It is not surprising, therefore, that the isolated hepatocyte preparation has proved a popular tool. However, it should be noted that most of the studies have been carried out with rodent cells, and the results sometimes extrapolated to humans. The validity of such extrapolation may be questioned in view of the wide interspecies differences in drug metabolism known to exist. The use of isolated liver cells for the study of hepatic drug uptake, metabolism and toxicity has been extensively reviewed (Klaassen and Stacey, 1982; Smith and Orrenius, 1984; Suolinna, 1986). Uptake studies have been performed for a large number of drugs, and kinetic parameters have been measured. Isolated hepatocytes have proved very useful for the study of drug metabolism because the incubation medium

180


and cells can be easily separated for analysis of drug metabolites, following which the hepatocytes can be fractionated or extracted. Studies with isolated hepatocytes have proved particularly valuable for identifying metabolites that are short-lived or present in very small amounts. Hepatocytes can be used to examine partial reaction sequences as in spectral studies involving the combination of a drug with cytochrome P-450(Hirata et al., 1977). On the other hand, they can be employed to establish the overall metabolism of a compound. In this respect hepatocytes have a considerable advantage over microsomes, which often do not possess the complete drugmetabolizing sequence. The value of isolated liver cells for toxicity studies is widely recognized. Variables such as blood flow, nervous or humoral factors are eliminated, and the environment can be directly manipulated. Comparisons are readily made between hepatocytes exposed to the drug under study and control cells. The large number of hepatocytes that can be produced in a single preparation allows a much greater amount of data to be obtained in comparison to the isolated perfused liver or experiments in vivo. Acute toxicity can be examined by the effect of the drug on the rate of one or more of many cellular metabolic processes (Chapter 6). In addition, a special advantage of the hepatocyte preparation is the ease with which cells can be examined under the light or electron microscope, so that structural changes induced by exposure to a drug can be readily detected. However, it must be appreciated that the behaviour of the liver is to some extent dependent on its anatomy, and this cannot be mimicked by the isolated hepatocyte suspension. For example, normal concentration gradients of 0, are lost, and this may well have some bearing on sensitivity to certain toxins. Moreover, the disruption of lobular structure removes the hepatocyte’s polarity and largely prevents biliary secretion of any drug. There is good evidence that periportal and centrilobular hepatocytes show different metabolic behaviour (Jungermann and Katz, 1989),and these distinct patterns are lost when hepatic architecture is destroyed.

Ch. 7.

UTILIZATION

181

Gumucio et al. (1978) found that pretreatment with phenobarbital produced a substantial increase in the cytochrome P-450 content of centrilobular hepatocytes over the level in the same subpopulation prepared from untreated animals. No such increase was detected in periportal hepatocytes. The detection of differences of this kind is not feasible with the standard isolated hepatocyte preparation, in which periportal and centrilobular hepatocytes are inextricably mixed together (although an attempt has been made to separate the two classes by centrifugal elutriation (Sumner et al., 1983)). However, recent developments have enabled the preparation of suspensions enriched with hepatocytes derived from either periportal or centrilobular regions of the liver (Lindros and Penttilii, 1985; Quistorff, 1985), and these enriched preparations have been used to investigate regional hepatic drug metabolism (Suolinna et al., 1989; Gascon-Barre et al., 1989). Some of the difficulties relating to loss of cellular orientation and organization in the isolated suspension can be remedied by studying cells in adherent culture (Chapter 11). This approach also has its limitations, in that cultured hepatocytes prepared by collagenase perfusion may rapidly lose cytochrome P-450 and critical drug metabolizing enzymes (Sections 3.3 and 10.2). The extent and nature of this decrease are dependent on the culture conditions employed. Measures which have been used in an attempt to maintain levels of cytochrome P-450 in cultured hepatocytes similar to those found in vivo have been recently reviewed (Paine, 1990). Metyrapone has been found to maintain cytochrome P-450 levels in cultured hepatocytes (Paine, 1990), but this substance, however, inhibits cytochrome P-450-mediated oxidations and must be removed from the culture medium 24 h prior to addition of the drug under test. According to Paine and co-workers (Paine, 1990)levels of cytochrome P-450 are still maintained during this period. Hexobarbital has also recently been shown to maintain levels of cytochrome P-450 and other drug metabolizing enzymes in primary cultures of hepatocytes (Kim et al., 1988). It should be noted, however, that the term cytochrome P-450 encompasses a family of about 20

I82


haemoproteins (Gonzalez, 1988). Treatments such as those discussed may alter the relative distribution of these multiple forms (e.g. Steward et al., 1985; Schuetz et al., 1988). Metyrapone, for example, has been shown to increase those activities associated with polycyclic hydrocarbon-inducible cytochrome P-450 (Lake and Paine, 1982). Meredith (1988), using the EDTA perfusion procedure for the isolation of cells (Protocol 3. l), was able to culture cells which maintained cytochrome P-450and glutathione levels for 5 days (see Section 10.2). It would be of interest to determine whether the forms of cytochrome P-450present in cells prepared by EDTA perfusion are qualitatively and quantitatively identical to those found in vivo.

7.2. Drug uptake and metabolism by isolated hepatocytes 7.2.1. Measurement of drug uptake

The measurement of the rate of uptake of a drug (e.g. Blom et al., 1982; Iwamoto et al., 1986; Mol et al., 1988) is a procedure very similar to the determination of the rate of uptake of any normal metabolite. It requires a means of rapidly separating the hepatocytes from the incubation medium by centrifugation, and the subsequent determination of the intracellular concentration of the drug in the pellet after estimation of intracellular and entrained water. The procedures are similar to those described in Protocols 6.2 and 6.3, though the solution used to extract the pellet will, of course, depend on the nature of the drug under study. Studies of the uptake of drugs and xenobiotics by isolated hepatocytes have been particularly advantageous in elucidating the mechanisms of transport (Klaassen and Watkins, 1984). Relevant kinetic parameters can be readily determined, free from interference by non-specific binding to plasma proteins or the influence of haemodynamic factors. The isolated hepatocyte preparation facilitates

Ch. 7.

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the study of drug interactions, whereby one chemical may enhance or diminish the uptake of another. It is also feasible to examine how various pretreatments of the donor animal will affect hepatic drug uptake (Klaassen and Stacey, 1982). Such studies have suggested the existence of at least three different pathways involving carrier-mediated uptake mechanisms, each involved in the transport of a separate class of hydrophilic compound (Schwenk, 1980; Klaassen and Watkins, 1984; Mol et al., 1988). Lipophilic xenobiotics, on the other hand, appear to pass through the membrane by passive diffusion (Chen et al., 1980; Schwenk, 1980; Ziegler et al., 1988). Using representative compounds for each of the putative three carrier systems, Blom et al. (1982) found that although the clearance by uptake of d-tubocurarine (a member of the class of exogenous organic cations) by isolated hepatocytes was very similar to that of the isolated perfused liver, the clearances of ouabain (representative of hydrophilic neutral organic compounds) and dibromosulphophthalein (an organic anion) were each a factor of 2-3 lower in the isolated hepatocytes although the quality of the cell preparation was good. The reason for these differences has yet to be determined. Similar uptake studies have suggested the mechanism of the selective toxicity towards the liver of phalloidin and a-amanitin, two toxins isolated from the poisonous toadstools of the genus Amanita. It appears that these poisons are transported into hepatocytes by an energy-dependent, liver-specific, carrier system that is also involved in the uptake of other organic anions (Frimmer et al., 1980; Petzinger et al., 1983; Wieland et al., 1984; Kroncke et al., 1986). Wieland et al. (1984) and Kroncke et al. (1986) have used photoaffinity labelling techniques to provide evidence for common carrier-mediated transport processes. Hepatocytes are preincubated with photolabile derivatives of bile salts, or toxins such as phalloidin, the plasma membrane fraction is isolated, and polypeptides are identified that are capable of binding these compounds, and therefore presumably involved in their uptake.

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7.2.2. Study of drug metabolism

The reactions of drug metabolism are conveniently classified into two groups. The major phase I reactions involve oxidation reactions catalysed by the monooxygenases (mixed function oxidases). Other phase I reactions include reductive or hydrolytic processes. All these reactions are located within the endoplasmic reticulum. This cellular membrane system yields the microsomal fraction when whole liver is homogenized and fractionated. In phase I1 reactions, a small water soluble group is added to the molecule, often as a sequel to an oxidation reaction. Of the various conjugating reactions glucuronidation, catalysed by UDP-glucuronyltransferase, is the major process, but sulphation, acetylation and conjugation with glutathione are also important. Isolated hepatocytes have been widely used over the past 20 years to investigate drug metabolism. The behaviour of the cells can be studied in short-term or long-term suspension (Chapter lo), or in adherent culture (Chapter 11). Cells prepared by mechanical means have been employed for drug studies (Henderson and Dewaide, 1969), but such preparations represent particulate fractions of liver cells (Berry, 1961) and are likely to yield results similar to those found with microsomes. Whereas damaged hepatocytes immediately lose the majority of cytoplasmic functions, endoplasmic reticulum activity is to a considerable extent maintained, particularly if co-factors are provided. Hence in studying the metabolism of xenobiotics it is critical to ensure that the integrity of the cell preparation is satisfactory before conclusions can be drawn that the metabolic processes observed resemble those occurring in vivo. In keeping with findings in regard to drug uptake, rates of phase I drug metabolism by isolated hepatocytes are frequently, but not always, as high as rates in the perfused liver or in microsomal preparations (Klaassen and Stacey, 1982). The metabolism of hexobarbital (Stacey, 1978), of other barbiturates (Yih and van Rossum, 1977) and of benzo(a)pyrene (Vadi et al., 1975) and antipyrine (Aarbakke et al.,

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1977) by isolated hepatocytes has been shown to be similar to that of the perfused liver. Unsupplemented microsomes are incapable of carrying out phase 11 reactions, so that products of phase I reactions tend to accumulate, whereas in the intact cell phase I products generally undergo rapid further metabolism by a cytoplasmic phase I1 reaction (Klaassen and Stacey, 1982; Salonen and Suolinna, 1988). The ability of isolated hepatocytes to carry out phase I and phase I1 reactions has been demonstrated for a wide variety of drugs and donor species (Fabre et al., 1988; Salonen and Suolinna, 1988; Seddon et al., 1989a,b; Suolinna et al., 1989). A consequence of this is that in some cases the rate of production of an intermediate formed by a phase I reaction appears to be slower in isolated hepatocytes than in the microsomal preparation. For example, the concentrations of two intermediary metabolites of aflatoxin B1, aflatoxins QI and M I , were lower following incubation of intact hepatocytes with the parent compound compared to their incubation with a microsomal preparation (Metcalfe et al., 1981).The formation of polar metabolites of the aflatoxins, that were presumably products of phase I1 reactions, was detected in intact cells. The difference between the metabolite patterns in the intact cells and the microsomes was potentiated after pretreatment with an inducer such as phenobarbital or 3-methylcholanthrene, and was apparently due to the conjugation reactions, which were intact in the hepatocytes but not in microsomes. Although microsomes may be useful in the identification of potentially harmful metabolites, the behaviour of the isolated hepatocyte is more likely to reflect that of the liver in vivo, due to the fact that the isolated cells can perform both phase I and phase I1 reactions at rates similar to those of the intact organ (Wiebkin et al., 1976; Billings, 1977; Fabre et al., 1988). The profile of metabolites (usually by HPLC analysis) following incubation of isolated hepatocytes with the drug under study has frequently been found to be very similar to the profile of urinary metabolites (Green et al., 1986; Salonen and Suolinna, 1988) or blood metabolites (Seddon et al., 1989a.b)

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determined following oral administration of the drug. Comparisons between isolated hepatocytes and microsomes of the binding of drugs to cytochrome P-450, as determined by spectral analysis, have shown that the two preparations are similar, although the spectrum forms more slowly in hepatocytes, possibly due to a limitation in the rate of drug uptake (Moldbus et al., 1973; Hirata et al., 1977). Drug-induced spectral changes may offer a useful tool for studying drug uptake and metabolism, but it may be difficult to distinguish between spectral changes due to interaction with cytochrome P-450 and those due to interaction of drugs with other haemoproteins of the hepatocyte (Smith and Orrenius, 1984). The ability of isolated hepatocytes to perform various phase I1 reactions introduces difficulties in determining the overall rate of disposal of a substance by a phase I reaction, since the rate of product formation cannot readily be used unless all metabolites are identified and measured. Despite this, rates of isolated hepatocyte metabolism for a large number of drugs have been determined, frequently by substrate disappearance (e.g. Moldeus et al., 1974; Richard et al., 1989; Seddon et al., 1989b), and it has been found that many drugs are metabolized in isolated hepatocytes at rates similar to those observed for the relevant phase I reaction with microsomes (Moldeus et al., 1974; lnaba et al., 1975; Junge and Brand, 1975; Fabre et al., 1988). Thus, phase I1 reactions do not appear to be rate-limiting. Although intact hepatocytes can metabolize drugs without the need for added co-factors, the nutritional state of the cell is an important factor in determining the rate of drug metabolism (Guillouzo, 1986). NADPH is a crucial co-factor for monooxygenation of many drugs, and in the well-fed animal glycogen serves as a major reservoir of substrates for the NADPH-generating pentose phosphate pathway. However, in the starved state a high level of cytoplasmic NADPH cannot be maintained unless a source of mitochondria1 substrate such as lactate is provided (Smith and Orrenius, 1984). The isolated hepatocyte preparation has proved particularly useful for studying interspecies differences in drug metabolism (Moldeus,

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1978; Holme et al., 1986; Humpel et al., 1989; Seddon et al., 1989a). For example, 2 mM paracetamol (acetaminophen) increased the permeability of mouse hepatocyte membranes, as judged by NADH penetration, whereas incubation with the same concentration of paracetamol for 5 h produced no such increase in rat hepatocytes, even following pretreatment of the donor rats with phenobarbital to induce cytochrome P-450-linked oxygenases (Moldeus, 1978). This was in keeping with the increased resistance of rats to paracetamol in vivo. The rates of paracetamol glucuronidation in the mouse and the phenobarbital-treated rat were found to be very similar, whereas the rate of sulphate conjugation was ten-fold higher in the rat than the mouse. On the other hand, the formation of the glutathione conjugate by mouse hepatocytes was about twice that found in the rat. Species differences in the metabolism of the liver carcinogen 2-acetylaminofluorene by monolayer cultures of hepatocytes from mouse, hamster, rat and guinea pigs were found by Holme et al. (1986). The ratio of activation reactions, as measured by the amounts of covalently bound metabolites of 2-acetylaminofluorene, to detoxification reactions, as measured by the sum of C-hydroxylated and watersoluble metabolites, was highest in rats in keeping with the higher susceptibility of the rat to this agent. The possibility that drug metabolizing enzymes may vary with age and sex must also be considered when animals are chosen for such studies (Guillouzo, 1986; Iwamoto et al., 1986; Adamson and Harman, 1989). The uptake and metabolism of a drug by the human liver may exhibit marked differences to that observed with liver of experimental animals (Green et al., 1986; Fabre et al., 1988). Recent improvements in the isolation of human hepatocytes from large biopsy samples, or if available, whole livers (Section 3.4.6) have increased their utility as a model for studying hepatic drug metabolism (Green et al., 1986; Fabre et al., 1988; Seddon et al., 1989a). Where human hepatocytes are unavailable, hepatocytes prepared from the livers of cynomolgus monkeys are considered by some workers to show a pattern of drug

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metabolism very similar to that observed with human hepatocytes (Seddon et al., 1989b).

7.3. Isolated hepatocytes and drug toxicity 7.3.1.Introduction The literature on the toxic effects of drugs on isolated hepatocytes is very substantial and has been the source of a number of comprehensive reviews (Klassen and Stacey, 1982; Smith and Orrenius, 1984; Tyson and Green, 1987). The toxic effects of chemicals have been examined both biochemically and morphologically. Certain agents such as toluene (Hilderman et al., 1975) and haloalkanes (Ungemach, 1987) may act directly on the cell membrane by means of a solvent effect. Others such as quinones induce their effects through cellular reactions that generate active oxygen species (Thor et al., 1982; Smith et al., 1985), the metabolism of which may be accompanied by severe depletion of intracellular glutathione. Another group of agents including halothane and carbon tetrachloride (reviewed by Kappus, 1987) are thought to bring about cell damage by inducing lipid peroxidation and by covalent binding of reactive intermediates to macromolecules (de Ruiter et al., 1982; Recknagel, 1983; Santone et al., 1986; Gutteridge and Halliwell, 1990) although a direct causal relationship between the observed peroxidation and cell death is disputed (Ungemach, 1987). Isolated hepatocytes have also been used to assess the nature and extent of DNA damage caused by radiation, and carcinogens such as daunorubicin. and the ability of the cell to repair such damage (e.g. Sina et al., 1983; Howell et al.. 1986). Hepatocytes have been extensively used to assess metabolic disturbances caused by drugs, metals and other foreign compounds (Poul et al., 1979). The rates of various metabolic pathways can be measured as described in Section 6.6 and hence the effect of drugs and other compounds on these pathways can be determined. For example. cer-

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tain anaesthetics have been shown to increase 0,-consumption (Becker, 1988),decrease ureogenesis and increase the cytoplasmic redox state (Stacey et al., 1978). Urea synthesis is also inhibited by the antiinflammatory drug, butibufen (del Prado Miguez et al.. 1986) : i d acetylsalicylate (Kay, 1988). Paracetamol (Burcham and Harinan. 1989) and the sulphonylureas, tolbutamide and glipizide (Mojena ct al., 1989) increase glycogenolysis by activation of glycogen phosphorylase, while the Ca2+antagonist, verapamil, has been shown to inhibit ketogenesis in isolated hepatocytes (Olubadewo et al., 1988).

7.3.2. Indications of cell dumuge The gross signs of cellular injury brought about by drugs are similar to those associated with cell damage from other causes (such as anoxia). Hence, cellular injury caused by toxic substances is frequently accompanied by an increase in plasma membrane permeability (Smith and Orrenius, 1984) that can be estimated by uptake of trypan blue and the leakage of enzymes such as LDH as described in Chapter 4. Goethals et al. (1984) found that paracetamol and the psychotropic drug, chlorpromazine, inhibited [14C]leucineincorporation into total protein at concentrations which did not cause membrane damage as judged by LDH leakage. The antihistaminic drug, promethazine, inhibited protein synthesis at a concentration of 0.01 mM. and inhibited intracellular glycogen accumulation at 0.1 mM. but increased LDH leakage only at the higher level of 0.5 mM. From these results, Goethals and co-workers suggested that impairment of protein synthesis and glycogen metabolism are more sensitive indicators of cell damage caused by xenobiotics than are other measures such as LDH leakage (cf. Krebs et al., 1979a; Section 4.2). When assessing the toxic effects of a particular substance. alterations in the intracellular concentrations of cell components such as glutathione, Ca?+,K+ and ATP are frequently estimated using procedures described in Protocols 6.3 to 6.5. However, as pointed out

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by Fariss et al. (1989, in examining toxicity due to a particular compound, it is desirable to be able to distinguish between events which contribute to cell death and those which occur as a consequence of it. As an aid to achieving this goal, it would be advantageous to be able to assess changes in the intracellular concentrations of metabolites in intact cells exposed to a particular toxin before any such changes had resulted in loss of integrity of the plasma membrane and leakage of cellular contents. For some metabolites separation of intact from damaged cells by centrifugation through Percoll may suffce (Section 2.8). If the metabolites under study are labile, a separation step that rapidly brings about cessation of metabolism will be necessary. Fariss et al. (1985) measured the concentrations of intracellular metabolites following centrifugation of hepatocytes through an oil layer of dibutyl phthalate (p = 1.046),into PCA, in a method analogous to centrifugation through silicone oil (Protocol 6.5). More than 90% of hepatocytes centrifuged through dibutyl phthalate excluded trypan blue, independently of the percentage that had done so in the original suspension. Previous workers have determined the density of damaged hepatocytes (as judged by trypan blue exclusion) to be 1.02-1.03 g/ml, markedly lower than that of intact cells ( p = 1.08) (Pertoft et al., 1977; Dalet et al., 1982). Thus, on theoretical grounds, intact cells would be expected to pass through the dibutyl phthalate layer, while damaged cells would be retained by it. Intracellular metabolite concentrations in PCAextracts of freshly prepared hepatocytes were similar to those of cells extracted after centrifugation through dibutyl phthalate or silicone oil. Thus, centrifugation through dibutyl phthalate does not adversely affect the measurement of intracellular concentrations in intact cells. Investigations in our laboratory have suggested that centrifugation through silicone oil according to Protocol 6.5 is also effective in separating damaged and undamaged cells (as judged by trypan blue exclusion), provided that the density of the silicone oil is adjusted to approximately 1.07 g/ml to optimize separation. It should be noted that this technique can hardly be considered a

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sensitive one, since it is capable only of distinguishing intact cells from those that have suffered a loss of plasma membrane integrity sufficient to allow leakage of intracellular contents. Nevertheless, it does allow measurement of the intracellular content of intact hepatocytes, without contamination by grossly damaged cells and therefore enhances the possibility of detecting changes in cells that are still intact, as judged by trypan blue exclusion, but that may be exhibiting early signs of toxic damage. 7.3.3. Investigation of solvent damage Many organic solvents are capable of causing damage to isolated hepatocytes by their direct solvent effects on the plasma or other membranes, whereby the physico-chemical properties of these membranes are disturbed. Most substances of this nature are of no biological interest, but some are important as anaesthetic agents (chloroform and halothane), some like carbon tetrachloride are toxic to man, while others have been used experimentally (e.g. toluene) as ‘permeabilizing agents’ (Hilderman et al., 1975) (Section 12.7). While this solvent effect certainly plays a part in the observed hepatotoxicity of these compounds (Berger et a1 , 1986), its relative contribution depends on the concentration of the substance to which the cell is exposed (Ungemach, 1987). For example, cellular injury as a result of exposure to CCl, above 1.3 mM is thought to be due to a direct solvent effect (Pencil et al., 1982). At lower concentrations damage due to mechanisms such as lipid peroxidation predominates (Ungemach, 1987). Many of these substances are volatile and have low solubilities in water (e.g. carbon tetrachloride). Therefore there is no certainty that addition of the agent directly to the medium without a pre-incubation period, as is frequently practised (Stacey et al., 1978; Long and More, 1987), will generate the desired initial concentration. Insufficient attention to such incubation conditions may result in misinterpretation of results (Ungemach, 1987).

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To overcome this difficulty Glende and co-workers (Pencil et al., 1984; Glende and Pushpendran, 1986)placed tubes (6 x 50 mm), containing the substance under study, into 50-ml conical flasks containing incubation medium alone, stoppered these flasks and incubated them for at least 15 min to establish equilibration of the volatile compound between gas and liquid phases. Manometer vessels, with centre wells (Section 6.5.2) can also be used for this purpose. 7.3.4. Investigation of the metabolism of drugs associated with free-radical formation

The isolated hepatocyte preparation has proved a useful tool for demonstrating the metabolism of a wide variety of drugs to free-radical intermediates. It has proved possible to detect these intermediates by means of ‘spin traps’. For example, Tomasi and co-workers have detected free-radical intermediates of hydrazine derivatives (Albano et al., 1989),chloroform (Tomasi et al., 1985)and 1,2-dibromoethane (Tomasi et al., 1983). Care must be taken in using spin traps that the trap itself does not inhibit hepatocyte metabolism (Albano et al., 1986). An important class of drugs capable of being metabolized to freeradicals are quinones (Powis et al., 1984; Smith et al., 1985; Mirabelli et al., 1988; Thor et al., 1988). Their hepatic metabolism is of considerable interest because many important antitumour drugs contain the quinone nucleus. The quinones are substrates for flavoenzymes and can undergo either two-electron reduction to the hydroquinone or one-electron reduction to the semiquinone radical (Iyanagi and Yamazaki, 1970). The antitumour and cytotoxic effects of quinonerelated drugs are believed to be brought about through their oneelectron reduction to semiquinone radicals and subsequent autoxidation to generate superoxide (02-) and re-form the quinone. The possibility therefore exists for a cyclic process, known as redox cycling (Kappus and Sies, 1981; Smith et al., 1985; Kappus, 1986), whereby a small amount of quinone can serve as a catalyst for the oxidation of substantial quantities of NADH, with the concomitant formation

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of stoichiometric amounts of 0,-.As a result, the cell is placed under oxidative stress by depletion of NADH and exposed to oxygen radicals that cause intracellular glutathione depletion, damage to membranes through lipid peroxidation, and oxidation of thiol groups in proteins (Thor et al., 1982; Gutteridge and Halliwell, 1990). Furthermore, two 02-radicals dismutate to yield hydrogen peroxide (H,O,) - - and 0,, either spontaneously or as a result of catalysis by superoxide dismutase. - - thus produced can then react with further 0,- to produce The H,O, damaging species such as the hydroxyl radical and singlet oxygen in a reaction cdtalysed by metals such as Fez+or Cu' (Smith et al., 1985; Gutteridge and Halliwell, 1990). The detection of 0,- production by whole cells, as a measure of drug toxicity, is not easy. The usual test for intracellular 02-production is the reduction of cytochrome c or acetylated cytochrome c (Azzi et al., 1975; Thayer, 1977). These substances would not be expected to penetrate intact cells, so that any reduction of cytochrome would imply release of 0,-from the hepatocyte. Normal hepatocytes do not release OF, but reduction of acetylated cytochrome c has been reported in hepatocytes exposed to menadione (Thor et al., 1982). However, the toxicity of this quinone raises the possibility that the detectable 0,- is being generated by damaged cells whose contents have become accessible to cytochrome c. The reduction of cytochrome c is not in itself sufficient evidence for the production of 0,- since it can also be caused by other compounds such as menadiol, and it is necessary to show that the reduction is inhibited by superoxide dismutase. Hydrogen peroxide, which is frequently produced during the metabolism of xenobiotics, including quinones, can also be measured. However, the detection and quantitation of intracellular H,O, presents similar difficulties to those associated with superoxide measurement. Until recently only a semi-quantitative approach was available involving the spectrophotometry of the primary catalase H,O, complex (Compound I) (Margoliash et al., 1960). Boutin et a]. ( 1989) have now published a novel method for determination of H,Oz

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production within hepatocytes. It depends on the fact that the irreversible inhibition of catalase by 3 amino-l,2,4-triazole (ATz) is mediated by a complex formed by ATz with Compound I. Methanol has a protective effect on catalase activity during ATz inhibition that is reversible as a function of H 2 0 2production. It follows that the inhibition of catalase activity by ATz is dependent on the formation of H,02. In the assay, the ability of hepatocytes to metabolize H202via the catalase pathway is determined on a portion of the cell suspension after incubation of the cells for 30 min in the presence of 2 mM methanol and 20 mM ATz. The remainder of the hepatocytes are then incubated with the xenobiotic under study for 10 min and then gently pelleted. The supernatant is discarded and the cells resuspended in the same volume of Krebs-Henseleit medium for re-estimation of their capacity to metabolize H,02. There appears to be a direct relationship between the quantity of H 2 0 2produced during metabolism of the xenobiotic and the fall in the subsequent capacity of the hepatocyte to metabolize HzO,. The ability of externally generated free-radicals to irreversibly damage isolated hepatocytes has also been demonstrated. Free-radicals have been derived from the metabolism of cysteine (Vifia et al., 1983) or eugenol (Thompson et al., 1989). In both instances more than 80% of the cells were rendered permeable to trypan blue within 60 min. 7.3.5. Morphological indications of drug-induced damage

Potential cellular damage caused by the incubation of hepatocytes with a xenobiotic can be readily assessed by light microscopy (Section 5.1.2). If warranted, more detailed information can be obtained by transmission electron microscopy (Section 5.2.4). Blebbing of the plasma membrane (Section 5.1.2) is an early sign of damage readily visible under the light microscope. Such blebs are produced by a wide range of toxins including quinone-related compounds such as menadione and p-benzoquinone (Jewel1 et al., 1982; Thor et al., 1982; Mirabelli et al., 1988; Thor et al., 1988), CCl, (Yamamoto and Sugihara, 19881,

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phalloidin (Watanabe and Phillips, 1986) and microcystin-LR isolated from cyanobacteria (Eriksson et al., 1989). Each of the above bleb-inducing agents has been shown to produce cytoskeletal changes in the amount and structure of cytoskeletal protein and particularly actin filaments (Watanabe and Phillips, 1986; Mirabelli et al., 1988) although the actual mechanisms elucidated to date for the individual toxins appear to differ (Thor et al., 1988; Eriksson et al., 1989). For example, oxidation of protein thiols and disturbances in Ca2+homeostasis are intimately related to bleb formation caused by quinones such as menadione (Jewel1 et al., 1982; Kass et al., 1988; Mirabelli et al., 1988) whereas such changes are not observed under conditions in which blebbing is induced by microcystinLR (Eriksson et al., 1989). The relationship of Ca2+homeostasis, bleb formation and cell toxicity has recently been reviewed (Orrenius et al., 1987, 1989; Thomas and Reed, 1989). 7.3.6. Glutathione depletion

A key indicator of free-radical damage is depletion of cellular glutathione. Several recent reviews provide detailed discussion of the metabolism and functions of glutathione in isolated hepatocytes, agents that deplete intracellular glutathione and the mechanism by which such depletion causes cell death (Orrenius et al., 1983, 1987; Comporti, 1987). The measurement of intracellular GSH and GSSG is not easy because the reduced form is readily oxidized during chemical analysis. In isolated hepatocytes, greater than 99% of glutathione is normally present in the reduced form (Aw et al., 1986). In the presence of a free-radical generator such as menadione, much of this is oxidized (Di Monte et al., 1984). The oxidized form can be released from the cell. While glutathione depletion is commonly associated with the metabolism of drugs that generate free-radicals, this is not an invariable finding. Carbon tetrachloride toxicity, for example, is not associated with a fall in hepatic glutathione content (Orrenius et al., 1980). Conversely, glutathione depletion in some cases is due not to free-radical

196


generation but rather to its use as a conjugating agent (Moldtus, 1978; Comporti, 1987). The total concentration of unbound glutathione (GSH and GSSG) can be measured in isolated hepatocytes by colorimetric, spectrofluorometric and enzymatic methods (for a review, see Tyson and Green, 1987). However, perhaps the most specific and reliable method for the measurement of total glutathione, GSSG and hence by difference GSH is the enzymatic recycling method using glutathione reductase and 5 3 '-dithiobis-(2-nitrobenzoic acid) (Owens and Belcher, 1965; Tietze, 1969) as modified by Griffith (1980, 1985). Duplicate samples are assayed; to one of these is added 2-vinylpyridine which derivatizes GSH, preventing its participation in the reaction, and allows the measurement of GSSG alone. This assay is very sensitive, permitting the detection of pmolar levels of GSH. N-ethyl-maleimide has also been used as the masking agent (Tietze, 1969; Akerboom and Sies, 1981). Methods are also available to measure protein mixed disulphides, and protein sulphydryl group content (Di Monte, 1984). 7.3.7. Lipid peroxidation

Lipid peroxidation by oxygen free radicals affects unsaturated lipid components of cells, particularly membrane phospholipids. The mechanism by which this peroxidation occurs and its role in toxic liver damage has been recently discussed in detail (Ungemach, 1987; Gutteridge and Halliwell, 1990). A wide variety of products may result, and a range of analytical techniques are available for their measurement. These have been extensively reviewed (Recknagel et a]. 1982; Kappus, 1985; Tyson and Green, 1987; Gutteridge and Halliwell, 1990). The technique most commonly used in hepatocyte studies is measurement of malondialdehyde accumulation (for a detailed method see Tyson and Green, 1987), but frequently special non-physiological conditions are required to detect this accumulation. Moreover, artifacts are frequently encountered that give rise to an over-estimation of the degree of lipid peroxidation. These include interaction of the detec-

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ting agent, thiobarbituric acid, with aldehydes and other cell coniponents, autoxidation of lipids during the analytical procedures, and formation of malondialdehyde by oxidation of non-lipid components (Tyson and Green, 1987). On the other hand, malondialdehyde can be further metabolized by mitochondria in intact cells (Smith and Orrenius, 1984). There is some degree of consensus that malondialdehyde accumulation is not a particularly reliable, and certainly not a quantitative measure, of hepatocyte lipid peroxidation. An alternative technique, presently considered the most reliable, is the measurement by gas chromatography of the production of hydrocarbon gas, particularly ethane, generated by the breakdown of unsaturated fatty acid hydroperoxides (Smith et al., 1982; Kappus, 1985). Measurement 01’ pentane has also been employed, but there is evidence that this gas can be metabolized by the liver. Moreover, some pentane remain5 dissolved in the aqueous phase (Wendel and Dumelin, 1981; Smith et al., 1982). It should be noted, however, that the degree of ethane production is 100-500-fold lower than that of malondialdehyde. and malondialdehyde generation is considered to involve only lol’/~lo r the total lipids peroxidized (Kappus, 1985). There are also technical ditficulties in ethane detection, in that the gas phase above the incuhation mixture must be reduced to enhance the probability of detection by gas chromatography. Smith et al. (1982) incubated 50 ml of cell suspension in Erlenmeyer flasks with a total volume of 67 ml. This limits the amount of 0, available to the cells and shortens the length of the incubation. The substitution of carbogen for air may overcome this difficulty, but the use of a gas phase rich in 0, in studies on lipid peroxidation has been questioned (Tyson and Green, 1987). Indeed, it has been demonstrated that lipid peroxidation induced by haloalkanes (e.g. CCI, and halothane) is favoured under air and that peroxidation decreases with increasing PO,. being low under carbogen (Ungemach, 1987). Other methods that have been employed to monitor lipid peroxidation include the detection of chemiluminescence (Smith et al.. 1982;

198


Cadenas and Sies, 1984), fluorescent pigments (Recknagel et al., 1982; Smith et al., 1982) or conjugated dienes (Kappus, 1985). However, none have been shown to be more reliable than ethane evolution. Attempts to demonstrate the ability of toxic agents to cause lipid peroxidation when incubated with isolated hepatocytes have, on occasion, produced contradictory results (Ungemach, 1987). In addition to the effect of 0,-tension, lipid peroxidation may also be influenced by the composition of the incubation media. Iron contamination of added chemicals may be sufficient to induce intracellular propagation of lipid peroxidation. On the other hand, lipid peroxidation may be reduced in Tris buffer, probably due to its ability to scavenge free radicals (Weddle et al., 1976; Stacey et al., 1982). The presence or absence of albumin in the incubation medium is likely to have a similar effect. 7.3.8. Covalent binding

The covalent binding of foreign chemicals to cellular macromolecules has been proposed as an important mechanism of hepatotoxicity (Gillette, 1974; Mitchell and Jollows, 1975; Thorgeirsson and Wirth, 1977), although a role for such binding in causing cell death has not yet been established (Tyson and Green, 1987; Ungemach, 1987). The isolated hepatocyte preparation is well-suited for the measurement of adduct formation. Generally, a radiolabelled substrate is employed and measurement is made of the amount of label that is bound strongly to protein (Pohl and Branchflower, 1981). The macromolecules are usually precipitated with TCA and collected by filtration on glass fibre filters or by centrifugation. Samples are then repeatedly extracted with solvent to remove non-covalently bound material. Samples may also be washed with NaOH and then reprecipitated with acid, to demonstrate that the radiolabel remains associated with amino acids and small peptides, and therefore that the bond between the chemical agent and the protein is covalent (Thorgeirsson and Wirth, 1977). Remaining protein containing covalently-bound radioactivity is dissolv-

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ed in NaOH or a tissue solubilizer and the radioactivity counted by standard methods. Misleading results can occur if the test substance contains labelled impurities or if non-specific binding occurs. To correct for non-specific binding that is not removed by repeated precipitation and washing, the radioactivity associated with heat- or acid-denatured cells is subtracted from the radioactivity associated with the unprecipitated cells (Pohl and Branchflower, 1981). Some researchers have attempted to measure covalent binding in fractions of hepatocytes prepared after exposure of intact cells to the binding agent. For example, Loeper et al. (1989) incubated hepatocytes with radiolabelled isaxonine, homogenized them and then fractionated the suspension by differential centrifugation. A problem with this approach is that it is very difficult to distinguish between covalent and non-covalent binding. Methods are also available which measure the degree of binding to specific macromolecules such as DNA, RNA and protein (Shaw et al., 1975; Beland et al., 1979).

7.4. Metabolism and effects of ethanol A drug of great interest to biochemist, pharmacologist, pathologist and clinician alike is ethanol. The isolated hepatocyte preparation has been used extensively to investigate ethanol metabolism (e.g. Williamson and Tischler, 1979; Cronholm, 1987) and to study the effects of ethanol on hepatic intermediary metabolism (e.g. Christensen and Higgins, 1979). There are a number of technical points that need to be appreciated when studying ethanol metabolism. Because of the volatility of ethanol there can be a considerable loss from the incubation medium. To avoid this it is desirable to gas incubation vessels prior to addition of the ethanol and to seal each flask immediately after ethanol addition. Even so, there is a likelihood of some loss of ethanol during the unavoidable extraction and neutralization procedures that follow incubation. Since such a loss will inevitably cause an overestimation of the amount of

200


ethanol metabolized, an alternative approach to measuring ethanol metabolism is desirable. In liver, the major end-product of ethanol metabolism is acetate (Williamson et al., 1969). Although acetic acid is also volatile, the salts of the acid are not. The amount of acetate formed can be measured spectrophotometrically or alternatively [ ''C]ethanol can be used as the starting substrate. At the end of the incubation a neutralized extract is prepared in the usual manner. A measured sample (usually 0.5 ml) is transferred to a scintillation vial and brought to pH 9 with 50 pl 2 M KOH. The vial is then transferred to a hot-plate and the contents evaporated to dryness. The residue is then taken up in 0.5 ml H,O, a water-compatible scintillation fluid (e.g. ACSII, Amersham) added, and the counts in acetate determined. This procedure becomes even more necessary when employing the high ethanol concentrations (>40 mM) required for studying the properties of the ethanol oxidating system located in the endoplasmic reticulum. Isolated liver cells have been used to study the toxic effects of ethanol. Dianzani (1985) reports evidence for lipid peroxidation and glutathione depletion in hepatocytes incubated for 120 min in the presence of ethanol. Hepatocytes in monolayer culture have also been used to study the possible toxic aspects of ethanol metabolism such as the formation of acetaldehyde adducts with protein (Lin and Lumeng, 1990).

CHAPTER 8

Study of the effects of hormones

8.1. Hormonul responses exhibited by isolated hepatocytes Hepatocytes respond to a variety of different hormones when incubated in suspension. Rapid responses, involving changes in the concentration of intracellular messengers (e.g. cyclic AMP, inositol trisphosphate, Caz+and diacylglycerol), the phosphorylation of target proteins or the formation of an allosteric regulator, are induced by glucagon, a- and 6-adrenergic agonists, vasopressin and angiotensin I1 (Table 8. I ) . Longer term actions of these hormones or intracellular messengers, which can lead to an increase in synthesis of specific proteins within about 90 min (e.g. the induction of phosphoenolpyruvate carboxykinase activity by cyclic AMP), are also observed in suspensions of isolated hepatocytes. The available evidence suggests that glucagon, adrenaline, vasopressin and angiotension 11 each produce responses in isolated hepatocytes that are qualitatively and quantitatively similar to those induced in perfused liver. This comparison is difficult to make because the use of isolated hepatocytes has allowed the performance of a large number of experiments, directed towards investigation of the mechanism of action of hormones on liver cells, that cannot be carried out using the perfused liver.

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During the past 15 years many workers have studied the hormonal responses of hepatocytes to vasopressin and angiotensin 11. These hormones probably do not normally exert physiological effects on hepatocytes, although they may do so in certain pathological states. They have been particularly useful in studies of the role of inositol 1,4,5-trisphosphate and Ca2+ as intracellular messengers in hepatocytes because they do not induce changes in cyclic AMP. By contrast, adrenaline and other adrenergic agonists can bind to both a- and 0-adrenergic receptors and cause changes in Ca2+as well as cyclic AMP. Many of the actions of insulin and epidermal growth factor (EGF) that are rapid in onset are observed in isolated hepatocytes. These hormones interact with plasma membrane receptors possessing a proteintyrosine kinase catalytic site. The effects of insulin and EGF mainly involve changes in intracellular messengers such as cyclic AMP or Ca2+,and bring about immediate effects on metabolism. For example, the presence of insulin prevents the stimulation of hepatic glycogenolysis by glucagon or adrenaline (Table 8.1). Some slow-onset effects of insulin, such as the inhibition of protein degradation, are also observed in hepatocytes (Table 8.1). Many of the other slow-onset effects of insulin on the liver, observed in vivo, have not been clearly defined in any in vitro system including isolated hepatocytes. Responses of the liver to glucocorticoids are difficult to observe in isolated hepatocytes. This is principally because all these responses are slow in onset and therefore require incubation of the cells for at least 5 h. However, some effects of glucocorticoids on enzyme induction (see Chapter 10) have been observed (Table 8.1).

8.2. Optimal conditionsfor observing the effects of hormones 8.2.1. Isolation of hepatocytes

In early studies of hormone-responsive hepatocytes, some evidence for a loss of hormone receptors was reported (Garrison and Haynes,

TABLE 8. I Examples of responses in rat hepatocytes induced by hormones

Hormone

Example of response

References

Glucagon

Increased gluconeogenesis

Johnson et al. (1972) Claus et al. (1975) Pilkis et al. (1975) Garrison and Haynes (1975) Garrison and Haynes (1975) Pilkis et al. (1975)

Increased 0-utilization Increased pyruvate carboxylation Increased formation of cyclic AMP Increased glycogen synthesis Decreased glucagon- or a,-adrenergic agonist-stimulated glycogenolysis Decreased protein degradation

Johnson et al. (1972) Thomas and Williamson (1983)

Epidermal growth factor

Increased formation of inositol trisphosphate

Johnson and Garrison (1977)

Adrenaline (P-adrenergic receptors)

Increased glycogenolysis Increased formation of cyclic AMP

Cherrington et al. (1977)

Adrenaline (a-adrenergic receptors)

Increased glycogenolysis Decreased glycogen synthesis Increased intracellular free Ca”

Hutson et al. (1976) Hutson et al. (1976) Charest et al. (1985)

Vasopressin

Increased intracellular free Ca”

Increased formation of diacylglycerol

Thomas et al. (1984) Charest et al.. (1985) Thomas et al. (1984) Burgess et al. (1984) Hughes et al. (1984)

Induction of phosphoenolpyruvate carboxykinase

Salavert and lynedjian (1982) Watford et al. (1983)

Insulin

Increased formation of inositol trisphosphate

Blackmore et al. (1979) Hopgood et al. (1987)

Morgan et al. (1983)

0 5 Po

m

7 R 2 2

0 ;a

B2

2 !

h)

Glucocorticoids

8

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1973). This may have been due to the action of collagenase during perfusion of the liver and to proteases carried over into the final cell suspension (Pilkis et al., 1975). Some workers have used the stimulation by glucagon of gluconeogenesis as a monitor of hormoneresponsiveness and metabolic integrity (Pilkis et al., 1975). These and other studies established that in the preparation of hormone-sensitive cells, hyaluronidase should be omitted from the digestion medium and particular attention paid to: (1) the use of the shortest exposure of the liver to collagenase consistent with the separation of individual hepatocytes; (2) avoidance of hypoxia during perfusion of the liver; and (3) good temperature control of the perfused liver (Zahlten and Stratman, 1974; Pilkis et al., 1975; Hutson et al., 1976; Blackmore and Exton, 1985).These precautions appear to reduce damage to plasma membrane receptor proteins and other proteins involved in intracellular signalling, such as ion channels and transporters, adenylate cyclase and cyclic AMP phosphodiesterase. There is some evidence indicating that, in the preparation of hormone-sensitive hepatocytes from fed rats, the maintenance of high concentrations of glycogen during the isolation procedure improves the actions of glucagon and insulin (Wagle, 1974; 1975). Exton and his colleagues have routinely incorporated washed erythrocytes in the preparation of hormone-sensitive hepatocytes (Hutson et al., 1976; Blackmore and Exton, 1985) in the belief that this improves the quality of the hepatocytes. However, good hormonesensitive hepatocytes can be prepared without including red blood cells in the liver perfusion medium. For most experiments, the additional time and work involved in the use of erythrocytes is probably not warranted. Likewise, the inclusion of 1.5% gelatin in the medium used for incubation and washing of the cells during isolation (Blackmore and Exton, 1983) and the inclusion of fumarate, glutamate, glucose and pyruvate in the perfusion medium (Zahlten and Stratman, 1974) may enhance the quality of the cells, but their inclusion in the experimental protocol is generally considered not to be necessary.

Ch. 8.

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8.2.2. Incubation medium

An additional buffer is sometimes added to the Krebs-Henseleit medium in order to maintain pH at 7.4 duringthe incubation medium. 2 4 [ 2-hydroxy- 1,1 -bis(hydroxymethyl)-ethyl]amino)ethanesulfonic acid (TES) is most commonly used. For example, Haynes and his coworkers (Sistare and Haynes, 1985)have employed 100 mM TES buffer (pH 7.4), and have compensated for the increase in osmolality by decreasing the concentration of NaCI. For the incubation of hepatocytes in hormone studies, we recommend a medium containing 117 mM NaCl; 4.7 mM KCl; 1.2 mM KH,PO,; 1.3 mM Ca2+; 1.2 mM MgSO,; 24 mM NaHCO,; and 20 mM TES, equilibrated with carbogen a t 37°C and adjusted to pH 7.4 with KOH. This is a modification of Krebs-Henseleit bicarbonate buffer (KHBT). Some workers include gelatin or albumin in the incubation medium when studying the actions of hormones such as glucagon. Hutson et al. (1976) used 1.5% gelatin, while Thomas and Williamson (1983) employed 2% dialysed BSA. It has been suggested that gelatin or albumin improve the quality of hepatocytes in suspensions incubated for periods greater than an hour, although there is little definitive evidence about this. In the author’s experience, inclusion of gelatin or albumin for short incubation periods is not necessary, but the use of gelatin, or complex media (e.g. Waymouth medium) is desirable for prolonged incubations. Different approaches have been made to the question of the concentration of CaZ+to be employed in the extracellular incubation medium. There are a few studies which suggest that the presence of physiological Ca*+concentrations in the extracellular medium is actually detrimental to isolated hepatocytes. However, the great majority of workers believe that optimal incubation conditions include the presence of extracellular free Ca2+at a physiological concentration. As discussed earlier (Chapter 2), it is considered desirable to add the Ca2+back to hepatocytes as soon as possible after isolation of the

206


cells. It should be noted that the normal concentration of free Ca2+ in the blood of an animal is about 1.3 mM. Therefore, unless a Ca2+-bindingprotein such as albumin is present in the incubation medium, the concentration of total Ca2+in the incubation medium should be 1.3 mM. A pre-incubation at 37°C should be carried out for at least 15 min prior to hormone addition in order to allow re-establishment of basal metabolism and ion gradients. Some workers use an even longer preincubation period. When hormone additions are made to incubation mixtures, the hormones are generally dissolved in 150 mM NaCl to minimize effects on the osmolality of the suspension. In hepatocytes isolated from fed rats, glycogen is slowly broken down during subsequent incubation of the cells. Glycogen stores can be maintained by inclusion of glucose at a concentration of 15 mM in the incubation medium. Extracellular glucose both maintains adequate stores of glycogen, which may be required in studies of glycogen metabolism, and provides an exogenous source of energy for the cell. 8.2.3. Incubation vessel

In practice, the most convenient incubation vessel is a 20-ml plastic screw-capped scintillation vial. Cell incubations can also be successfully performed in 25-ml or 250-ml Erlenmyer flasks. In each case the volume of the incubation medium should be about 10% of the total volume of the incubation vessel. This ratio of incubation medium to gas space allows adequate oxygenation of cells. In all cases the containers should be gassed with carbogen and shaken at 90 oscillationdmin. The concentration of cells should be within the range 1-5 x lo6 cells/ml. There is a risk of hypoxia if a higher concentration of cells is used as the rate of diffusion of O2 into the cells may become limiting. A disadvantage of the incubation vessels described above is that they are not convenient for rapid removal of samples from the incubation medium. This difficulty can be overcome by the use of stationary

Ch. 8.

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207

cylindrical water-jacketed incubation chambers (diameter 2 cm, height 6 cm) in which the incubation medium is stirred. The incubation module designed by Yellow Springs Instruments for measurement of 0,-tension (Section 6.5.1) is highly satisfactory (Barritt et al., 1981). Four or eight of these cylindrical incubation chambers (one or two modules) can be employed at one time. The use of these chambers allows a large number of samples, ranging in volume from 0.1-2.0 ml, to be removed from the incubation mixture at relatively short time intervals. Thus it is possible to obtain time courses for the effects of hormones on enzyme activities such as glycogen phosphorylase and on the level of intracellular messengers (e.g. cyclic AMP). Hepatocytes can be incubated satisfactorily for periods up to 60 or 90 min using the stirred cylindrical chambers. However, the cells exhibit higher rates of leakage of LDH than cells incubated in shaken plastic vials.

Protocol 8.1 Effect of glucagon on cyclic A M P concentrations in hepatocytes (i) After their preparation, suspend the hepatocytes at a density of 6 x lo6 cells/ml in KHBT medium at a final pH of 7.4. (ii) Set up a number of incubation vials as follows. Add the cell suspension (0.5 ml) to 1.O ml of KHBT-medium in a 20-ml plastic vial and gas the vial for at least 10 s with carbogen at 1 I/min and seal the vial. Incubate the cells for 15 min. Add glucagon M will give maximum response) and continue incubation for the desired period of time. At the selected times (prior to and after the addition of glucagon) remove duplicate samples of the cell suspension (0.5 ml), and mix each sample with 0.6 M PCA (0.4 ml). After centrifugation at 1000 x g , in order to rerpove precipitated protein, and adjustment of the pH to 7.0 (Protocol 6.2), determine the amount of cyclic AMP present. Cyclic AMP is commonly measured by a protein binding assay (Gilman, 1970). The incubation conditions described in Protocol 8.1 can also be

208


used in studying the effects of glucagon on enzyme induction, for example the induction of phosphoenolpyruvate carboxykinase. However, the incubation period should be no longer than 90 min. For longer incubations, the conditions of Protocol 8.3 are used.

8.3. Special conditions for the study of specific hormonal effects Changes in the concentrations of phosphatidylinositol 4,5-bisphosphate, inositol 1,4,5-trisphosphate or diacylglycerol, as a consequence of hormone action, have been investigated by employing a variety of radiolabelled compounds. Workers have studied hormonal effects on the following reactions: phosphatidylinositol 4,5-bisphosphate hydrolysis (using ['Hlinositol and "Pi); inositol polyphosphate formation ([-'H]inositol); phosphatidyl choline hydrolysis and diacylglycerol formation (['Hlcholine, ['Hlglycerol and [3H]arachidonic acid). Cells have been successfully labelled with 32Pi(Billah and Michell, 1979; Kirk et al., 1979; Tolbert et al., 1980; Litosch et al., 1983; Seyfred and Wells, 1984), pH]inositol (Tolbert et al., 1980; Prpic et al., 1982; Litosch et al., 1983; Johnson and Garrison, 1987) or [-'H]arachidonic acid (Takenawa et al., 1982; Thomas et al., 1983; Pickford et al., 1987) by incubating hepatocytes in the presence of the labelled compound. An example of the pre-labelling technique is given below.

Protocol 8.2 The pre-labelling of hepatocytes with [JH]inositol (i) After isolation, suspend the hepatocytes at a density of 6 x lo6 cells/ml in KHBT medium containing 1.5% gelatin (KHBT/gelatin) and keep the cells at 37°C. (ii) Add the hepatocyte suspension (0.75 ml) to 1.25 ml of KHBTIgelatin in a 20-ml plastic vial. Add glucose to give a final con-

Ch. 8.

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centration of 15 mM, and my0-[2-~H]inositol (40 Ci/mol) to give a final concentration of 0.1 mM. After addition of the hepatocyte suspension, gas each vial for 10 s with carbogen delivered at a rate of 1 I/min and seal the vial. Incubate the cell suspensions at 37°C and shake at 90-120 oscillations per min. The total incubation time is 60 min. Gas each vial as described, every 30 min. After completion of the incubation, centrifuge the cell suspension at 50 x g for 75 s, remove the supernatant and suspend the cells in 2 ml of fresh KHBT/gelatin. Repeat this washing step once. The final cell suspension (about 2 x loh cells/ml) can then be incubated under the desired conditions in the presence or absence of the hormone under test. The cells respond quite well to hormones after the pre-labelling period. The labelling of isolated hepatocytes by the method described in Protocol 8.2 is much less expensive than a frequently employed alternative procedure that involves the administration of the radioactive label to rats by intraperitoneal injection (Kirk et al., 1979; Prpic et al., 1982; Polverino and Barritt, 1988). However, the labelling of hepatocytes with some compounds, such as [3H]choline o r ['Hlglycerol, requires incubation times longer than an hour, especially if an approach to steady-state, with respect to the distribution of the label, is required. When using these labelled metabolites it is therefore desirable to administer the radiolabelled compound to rats by intraperitoneal injection.

8.4. Actions of insulin Although hepatocytes respond to insulin (Table 8.1) the concentrations of the hormone required to induce changes in hepatocyte metabolism are often higher than those required to induce effects in the perfused liver (Pilkis et al., 1975). Moreover, the effects of insulin on hepatocytes are somewhat variable. It seems that insulin receptors, and receptors for other hormones and growth factors in which the

210


receptor protein includes a tyrosine kinase catalytic site, are more susceptible to damage during isolation of the hepatocytes. However, this point is not well documented. Hopgood et al. (1977) described specific conditions for the study of the inhibition by insulin of protein degradation in rat hepatocytes over a period of 6 h. Maximal proteolysis and response to insulin were obtained in an incubation mixture consisting of Ca*+-free KHBTlgelatin that also contained 2 mM leucine, 16.5 mM glucose, 20 mM TES, 100 pM EDTA and 10 pglml phenol red, 60 pglml of penicillin and 10 pg/ml of streptomycin sulphate. The incubation vessels were polyethylene scintillation vials with a volume of 20 ml. Each vial contained 2-3 x lo6 cells in a total volume of 1.5 ml. The authors suggested that the function of EDTA was to chelate heavy metal ions; EDTA appeared to protect cells from damage, while the omission of Ca2+reduced clumping.

8.5. Action of steroid hormones The induction of enzymes such as phosphoenolpyruvate carboxykinase and tyrosine aminotransferase by glucocorticoids is slow in onset. This presents a problem in studies with isolated hepatocytes since relatively long periods of incubation are required. A successful protocol for investigation of the effects of glucocorticoids on the induction of tyrosine aminotransferase activity that can also be used for studies of enzyme induction by non-steroid hormones, such as glucagon, is given in Protocol 8.3.

Protocol 8.3 Induction by dexamethasone of tyrosine aminotransferase activity (i) At the final stage of preparation suspend the hepatocytes, at a density of between 1 x 106 and 3 x lo6 cells/ml, in Waymouth medium modified to contain 0.2% BSA, 100 U/ml of penicillin and 100 pglml streptomycin.

Ch. 8.

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21 1

(ii) Incubations are performed in 20-ml plastic vials. To each vial, add 1.7 ml of the cell suspension and 0.3 ml of 5 pM dexamethasone in 0.9% NaC1. Gas the cell suspension with carbogen for 10 s and seal the tubes at the beginning of the incubation. Repeat gassing procedure at 30-min intervals during the incubation. The vials are incubated at 37°C with reciprocal shaking at 90 cycles per min. On completion of the incubation period of 5 h, remove a sample of the cells (1.0 ml) for assay of tyrosine aminotransferase activity. Since the incubation period is relatively long there is some degree of cell damage. Under optimal conditions the number of cells which exclude trypan blue decreases from greater than 90% at the beginning of the incubation to about 75% or 80% after 5 h.

8.6. Hormone effects on mitochondriu isolated from hepatocy tes It is also possible to study changes in mitochondria isolated from hepatocytes that have previously been exposed to a hormone. In the protocol below a Dounce homogenizer is used to rupture the cells. Care must be taken that this procedure does not also damage the mitochondria.

Protocol 8.4 Isolation of intact mitochondria from hepatocytes (i) Incubate hepatocytes (final density 1 x lo6 cells/ml) in 250-ml glass Erlenmyer flasks, shaken at 90 oscillations per min, at 37"C, in a medium comprising KHBT, 100 pM EDTA, 1.5% gelatin and 15-20 mM glucose (for studies on cells from fed rats) and with an atmosphere of carbogen. The total volume of the cell incubation mixture in each flask is 60 ml. In studies of hormone effects on mitochondria, an incubation time of between 0 and 90 min for pre-treatment of the intact hepatocytes with hormone can be used without undue damage to the cells. Add the appropriate quantity of the hormone to each flask to obtain the hormone concentration range under study.

212


(ii) Centrifuge the contents of each flask at 50 x g for 45 s at room temperature. Wash each pellet twice with 40 ml of ice-cold 250 mM sucrose, 5 mM HEPES, 0.5 mM EGTA, 0.05‘%,defatted BSA, adjusted to pH 7.4 at M ” C with KOH (Isolation Medium plus EGTA). Suspend each cell pellet (about 1 g wet weight of loosely packed cells) in 6 ml of Isolation Medium plus EGTA. (iii) Carry out the following steps at 0 4 ° C . Homogenize each cell suspension in a glass Dounce homogenizer (7 ml capacity) with 30 upand-down strokes of a tight-fitting glass pestle (Wheaton “A” or Kontes “B”). Centrifuge the homogenate at 300 x g for 10 min and remove the supernatant. Centrifuge the supernatant at 4500 x g for 5 min. The resulting pellet is composed mainly of mitochondria. Resuspend the mitochondrial pellet in 5 ml of Isolation Medium, free of EGTA, in the homogenizer by means of two up-and-down strokes of a loose-fitting pestle (Wheaton “B” or Kontes “A”). Pour into a centrifuge tube, add a further 4 ml of EGTA-free Isolation Medium, mix and centrifuge at 4500 x g for 5 min. Suspend the resulting mitochondrial pellet in 0.45 ml of Isolation Medium, without EGTA, and keep at U ” C . The expected yield of mitochondria is about 10 mg mitochondrial protein from 1.0 g wet weight of cells. The respiratory control ratios (the ratio of the rate of 0, consumption in the presence of ADP compared with that after conversion of all ADP to ATP), measured in the presence of succinate, should be about 4.5 for mitochondria from freshly-isolated hepatocytes, and 3.5 for mitochondria from hepatocytes previously incubated at 37°C for 60 min, compared with a value of about 6.0 for mitochondria prepared from fresh liver tissue. The presence of defatted BSA in the media used for the isolation and washing of the mitochondria is important as this improves their quality. This is reflected in a significant increase in ADP-stimulated respiration and in the respiratory control ratio. There is some variation in the tightness of fit of different Dounce homogenizers. Therefore a new homogenizer should be tested to deter-

Ch. 8.

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mine the number of passes required to optimize the yield and quality of mitochondria. Unless considerable cure is exercisrd thr Doirricc homogenizer can damage the mitochondria. An important factor in the isolation of well-coupled mitochondria from hepatocytes is the integrity of the hepatocytes during incubation at 37°C. Good mitochondria are obtained from cells which have suffered least damage. I t is considered that the inclusion of a protein such as albumin or gelatin (see Chapter 6 ) and maintenance of the pH of the incubation medium at 7.4 play an important par1 i n improving the quality of the cells at the end of the incubation period. Gellerfors and Nelson ( 1979) have used sonication to break hepatocytes, and have obtained well-coupled mitochondria. However, in our laboratory it has proven difficult to find conditions of sonication which give a high yield of well-coupled mitochondria (Hughes and Barritt, unpublished results). The time of sonication (about 15 s) recommended by Gellefors and Nelson is very short, making it difficult to avoid under- or over-sonication of the cells. Shears and Kirk (1984a) have developed a method for the preparation of small quantities of a mitochondria-enriched fraction from isolated hepatocytes. The method involves lysis of the cells with digitonin, mechanical disruption of the cells by forcing the cell suspension through a 23G syringe needle at high pressure, followed by rapid centrifugation of the broken cells through a layer of silicone oil. This results in a partially-purified mitochondria1 fraction which is contaminated with cell nuclei. This method has been used to measure mitochondrial membrane potential (Shears and Kirk, 1984a) and the amount of 45Ca2+in mitochondria from hepatocytes treated with hormones (Shears and Kirk, 1984b). Measurements of rates of respiration, Ca" movement and other metabolic parameters have shown that the properties of mitochondria isolated from hepatocytes are similar to those isolated from intact tissue, although, as noted above, the respiratory control ratios of mitochondria prepared from cells are lower than those prepared

214


from intact liver tissue (Garrison and Haynes, 1975; Hughes and Barritt, 1984). However, there have been few rigourous assessments of the degree of contamination of the mitochondria1 fraction by constituents of other organelles by means of a complete analysis of distribution of marker enzymes.

CHAPTER 9

Ca2+ion transport and compartmentation

9.1. Measurement of intracellular Ca2+ The main aims of experiments involving measurement of Ca2+in hepatocytes have been to elucidate the effects of hormones on Ca2+ pools and fluxes. Some experimenters have attempted to measure the amount of Ca2+present in intracellular pools of isolated hepatocytes and to compare these values with those obtained by other techniques, such as determination of the Ca2+content of isolated mitochondria. Methods have been developed for the measurement of changes in total cellular Ca2+,organelle Ca2+, Ca2+ fluxes and the concentration of free Ca2+in the cytoplasmic space. Hormones such as vasopressin, a,-adrenergic agonists and angiotensin I1 increase the hepatocyte cytoplasmic free Ca2+concentration by inducing the release of Ca2+from the endoplasmic reticulum and an increase in Ca2+inflow across the plasma membrane (Barritt et al., 1981; Charest et al., 1985). A considerable amount of the CaZ+released from the endoplasmic reticulum is subsequently pumped out of the cell by the plasma membrane (Ca2++ Mg2+)ATPase, even in the presence of a physiological extracellular Ca2+concentration. This can result in a transient decrease in total cell Ca2+soon after addition of the hormone (Blackmore et al., 1978).

216


The recommended incubation medium for most studies of intracellular Ca2+is Ca2+-freeKHBT (see Section 8.2.2). Where longer incubations are required, as when loading cells with quin2, it is our practice to supplement this medium with glucose (5 mM), EDTA (50 pM) and gelatin (1.5% w/v). 9.1.1. Determination of total hepatocyte Ca2+

Measurement of total hepatocyte Ca2+content has demonstrated that hormones, such as vasopressin and phenylephrine, can decrease total cellular Ca2+content (Blackmore et al., 1978; Blackmore and Exton, 1985), and has also revealed the ability of a variety of hormones to stimulate Ca2+movement into hepatocytes (Blackmore et al., 1984). The use of atomic absorption spectroscopy for the measurement of CaZ+is limited to the provision of information on changes in total cell Ca2+content and does not provide information on intracellular CaZ+ distribution.

Protocol 9.1 Measurement of total Ca2+by atomic absorption spectroscopy (i) Prior to measurement, cells will normally be incubated with various concentrations of Ca2+.A suitable apparatus is the incubation chamber of the 0,-electrode system supplied by Yellow Springs Instruments (Section 8.2.3). The initial volume of the incubation mixture is 12 ml.Cells are kept in suspension by means of a magnetic stirrer and bar which is rotated at the slowest speed that will keep all the cells in suspension. Carbogen is gently and continuously introduced to the top of the chamber at a rate of 1 I/min, though a thin polythene tube. The gas should pass through 150 mM NaCl before reaching the incubation chamber in order to increase its humidity and prevent evaporation of water during the incubation. (ii) After preparation of the isolated hepatocytes, suspend the cells at a density of 6 x lo6 cells/ml in Ca*+-freeKHBT medium, and keep

Ch. 9.

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the suspension at 0-4"C. Commence incubation of the cells by addition of 4 ml of this hepatocyte suspension to 8 ml of KHBT medium to which CaCI, has been added to give the desired final Ca2+concentration in the total volume of 12 ml. The final concentration of cells is 2 x lo6 cells/ml. (iii) To examine the effects of a hormone as a function of time, incubate the cells at 37°C for 15 min before addition of the hormone under test. The hormone is dissolved in 150 mM NaCl. An equal volume of 150 mM NaCl is added to control incubations. At the desired times, remove two 2-mI samples of the suspension and mix each sample with 1 1 ml of ice-cold wash medium comprising 150 mM NaCl containing 10Y0sucrose and 0.5 mM EGTA (pH 7.4) at 0 4 ° C in a 15-ml plastic conical centrifuge tube. Immediately centrifuge the tubes at 500 x g for 1 min. A centrifuge with rapid acceleration and deceleration times such as an MSE bench centrifuge, is desirable. (iv) Remove the supernatants carefully by aspiration, invert the tubes and allow them to drain thoroughly at room temperature. Remove any remaining drops of liquid in the tube using tissue paper wrapped around a glass rod. Take care not to touch the pellet. (v) Extract the Ca2+from the cell pellet by addition of 1 ml of 0.3 M PCA, containing 1 mM LaCI,. Mix the contents of the tubes on a vortex mixer in order to fully suspend the precipitate, and allow them to stand for 15 min at U 0 C before centrifugation for 10 min at 1500 x g. Remove and retain the supernatant and determine the amount of Ca2+present by atomic absorption spectroscopy, using standard procedures. Plot the resulting data as the amount of Ca2+associated with the cells (nmol per mg wet weight) as a function of time. One of the main problems in the measurement of total Ca2+is removal of the extracellular Ca2+.This can be facilitated by making a larger dilution of the cells into the wash medium, which will necessitate the use of a larger tube for centrifugation of the cells. Correction for the amount of Ca2+in the extracellular medium can be made using (hydr~xy[~~C]methyl)inulin (0.2 pCi/ml) or a similar marker of extracellular volume (Protocol 6.3) in a separate incuba-

218


tion. The cells are incubated, centrifuged and the pellets obtained and extracted, as for the atomic absorption measurements, after which the amount of (hydr~xy['~C]methyl)inulinin the PCA-extract is measured. 9.1.2. Determination of Ca2+content of the mitochondria and endoplasmic reticulum

An indirect method for the measurement of the amount of Ca2+in mitochondria and the endoplasmic reticulum in intact hepatocytes depends on the ability of an uncoupler of mitochondria1 oxidative phosphorylation, such as FCCP, to bring about the release of Ca2+ specifically from mitochondria, and of a Ca2+selective ionophore, for example A23187, to cause the release of Ca2+from the endoplasmic reticulum (Joseph et al., 1983). In cells incubated in the absence of added extracellular Ca2+,the Ca2+that is released from intracellular stores is extruded from the cell by the (Ca2++ Mg2+)ATP-ase in the plasma membrane. Cells are incubated for a short period of time in the absence of added extracellular Ca2+and the concentration of Ca2+in the extracellular medium is monitored using arsenazo I11 and dual wavelength spectroscopy (Joseph et al., 1983). When a baseline for extracellular Ca2+ has been established, FCCP (about 10 nmol/2 x lo6 cells) is added. The increase in extracellular Ca2+concentration is followed until a plateau is established, at which time the ionophore A23187 (about 2 nmol/2 x lo6 cells) is added and a second plateau of extracellular Ca2+established. The increase in extracellular Ca2+induced by FCCP is due chiefly to the release of Ca2+from the mitochondria. There is reasonable evidence to indicate that the second increase, induced by A23187, is due to Ca2+outflow from the endoplasmic reticulum. There are some difficulties in interpretation of results obtained using this technique. FCCP and A23187 may have effects on Ca2+ movement through membranes other than those of the mitochondria and endoplasmic reticulum. Moreover, detection of the released Ca2+

Ch. 9.

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219

depends on the ability of the plasma membrane (Ca2++ Mg2+)ATPase to transfer Ca2+from the cytoplasmic space to the medium. Furthermore, the plateaus reached after addition of FCCP or A23187 are not always clearly defined. Nevertheless, the method has been used successfully to estimate the amounts of Ca2+stored in the mitochondria and endoplasmic reticulum in the absence of hormones, to obtain information on the ability of hormones to release Ca2+from the endoplasmic reticulum (Joseph et al., 1983; Joseph and Williamson, 1983), and to study the action of toxic agents on these intracellular Ca2+stores (Bellomo et al. 1982).

9.2. Changes in Ca2+ movement monitored using 45Ca*+ 9.2.1. Available methods

The use of 45CaZ+ as a tracer for Ca2+movement and distribution in hepatocytes has provided information on the amounts of Ca2+in intracellular pools, rates of movement of Ca2+across the plasma membrane and the membranes of intracellular organelles, and on the effects of hormones and toxic agents on these processes. In principle, cells are incubated in the presence of 45Ca2+for a given period of time; the cells are then separated from the incubation medium and the amount of 45Ca2+ associated with the cells measured. If the time of incubation of cells with 45Ca2+is varied, plots of the amount of 45Ca2+associated with the hepatocytes as a function of time can be obtained. Experiments can be performed under steady-state conditions with respect to unlabelled Ca2+by adding 45Ca2+to the cells following prior incubation with unlabelled Ca2+for a time sufficient to allow the cells to achieve a constant concentration of Ca2+within intracellular compartments. Alternatively 45Ca2+can be added at an earlier stage, before such a steady-state has been reached (non-steady-state conditions). The majority of experiments have measured the uptake by cells of 45CaZ+ from the extracellular medium. However, Borle and his col-

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leagues (Studer and Borle, 1983; Studer et al. 1984) have measured rates of 45Ca2+efflux from cells prelabelled with 45Ca2+. While the steady-state approach has provided valuable information on pools and fluxes of Ca2 in hepatocytes, and on the effects of hormones on these processes, it has the disadvantage of being somewhat ambiguous. Thus, a number of alternative compartmental models are often consistent with the same set of experimental data. Distinction between some of these models requires each data point measurement to have a low standard error, and may need corroboration by the use of other experimental techniques. Another problem is that it is not always easy to show that hepatocytes are in a steady-state with respect to Ca2+.Furthermore, establishment of the steady-state may require relatively long incubation periods. The use of 4sCa2+under non-steady-state conditions has been useful in assessing the effects of hormones on intracellular Ca2+stores (Chen et al., 1978; Whiting and Barritt, 1982) and in evaluating the effects of energy status of hepatocytes on intracellular Ca2+disposition (Krell et al., 1979; Krell et al., 1983). In these experiments, hepatocytes labelled with 45Ca2+are incubated in the absence of added extracellular Ca2+or at low concentrations of extracellular Ca2+ and the hormone of interest is added during the incubation. Plots of the amount of 45Ca2+associated with the cells as a function of time reveal a sharp hormone-induced decrease in cellular 45Ca2+,which represents the outflow of Ca2+from the endoplasmic reticulum and the subsequent removal of this Ca2+from the cytoplasmic space by the plasma membrane (Ca2++ Mg2+) ATP-ase. Attempts have also been made to use non-steady-state approaches that employ 45Ca2+to measure the rates of Ca2+inflow across the plasma membrane (Assimacopoulos-Jeannet et al., 1977; Keppens et al., 1977; Foden and Randle, 1978). Although these experiments show stimulation of inflow of 45Ca2+by hormones such as vasopressin, adrenaline and angiotensin 11, consistent with the results obtained under steady-state conditions, the results of such experiments con-

Ch. 9.

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ducted under non-steady-state conditions are more difficult to interpret. A more detailed discussion of assumptions involved is given by Barritt et al. (198 1) and the references within that paper.

Protocol 9.2 Measurement of rates of 4-yCa2+ exchange under s t t w d j w u t P conditions (i) The incubation chambers employed are the same as those described for Protocol9.1.Theprocedurefortheseparationofhepatocytes from the incubation medium requires the preparation of 1.5-ml plastic centrifuge tubes (e.g. Eppendorf) containing three distinct layers of media as follows. The bottom layer is composed of 0.2 ml of 1.5 M PCA, the middle layer of 0.5 ml of silicone oil, and the top layer of 0.5 ml 150 mM NaCl containing 7% BSA and 2.3 mM LaCl,. The silicone oil is a mixture of 7 parts of Dow Corning 550 oil mixed with 1 part of Dow Corning 200 (1.5 centistokes) oil. Add the silicone oil and the top layer of NaCl using a smooth-working air displacement pipette (e.g. Eppendorf). Both the middle and upper layers must be added very carefully in order to prevent mixing of the layers. Some practice may be required in order to achieve this. After preparation of the three layers, centrifuge the tubes at 12,000 x g for 15 s (Eppendorf Microfuge 5415) in order to define the layers clearly. (ii) After preparation of the isolated hepatocytes, suspend the cells at a density of 6 x lo6 cells per ml in KHBT medium containing unlabelled CaCl, at the required final concentration (e.g. 1.3 mM). The cell suspension is incubated for 15 min at which time trace amounts of 4sCa2+are added (5 pCi, 16 nmol, 4sCaC1,) in a volume of approximately 5 pI. (iii) At the desired times, remove duplicate 100-p1 samples of incubation medium using an air displacement pipette and carefully add each sample to the upper layer of two silicone oil separation tubes. Immediately centrifuge the two tubes at room temperature at 12,000 x g for 10 s. Replace the tubes in the rack on the bench and allow

222


them to stand at room temperature until completion of all incubations (approx. 30-60 min). (iv) With a 25O-pl microsyringe, remove 100 pl of the PCA extract at the bottom of each plastic centrifuge tube. The amount of radioactivity present in the PCA-extract is determined by liquid scintillation counting. On beta counters that are not equipped with flexible programming windows, 45Ca2+can be satisfactorily measured using the setting for I4C-labelled compounds. (v) Removal of the sample from beneath the silicone oil layer must be performed carefully in order to prevent contamination of the PCA by fluid from the upper layer, since this contains a high concentration of 45Ca2+(derived from the extracellular medium). Place the needle of the microsyringe down the side of the plastic tube, being careful not to disturb the silicone oil layer or the precipitated protein at the bottom of the tube. After removing the needle, wipe it carefully to remove adhering upper layer and silicone oil. When the sampling of a series of plastic silicone oil tubes (for example, those obtained from one time course) is complete, rinse the syringe ten times with 1 mM unlabelled CaCl, and ten times with water before beginning the next series of sampling procedures. (vi) Some extracellular medium remains associated with hepatocytes and is carried through the silicone oil layer (see Protocol 6.5). Therefore, it is necessary to estimate the amount of 45Ca2+ present in this fluid. This is done by including [3H]inulin (15 p W 6 ml), added at the same time as 45Ca2+,in the incubation medium. Radioactivity in 45CaZ+ and [3H]inulin is determined using a dual isotope counting procedure. The time over which [3H]inulin measurements are made should be short, e.g. about 5 min, in order to reduce the uptake of [3H]inulin by pinocytosis. The observed volume of the extracellular fluid is about 0.25-0.3 pl/mg wet weight cells. After subtraction of the amount of 45Ca2+in the extracellular space, the amount of 45Ca2+ associated with the cells is expressed as nmol 45Ca2+per lo6 cells (see Section 2.6.1).

Ch. 9.

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Upon storage, [3H]inulin appears to be degraded to smaller [3H]labelled compounds which are able to enter hepatocytes and hence give an erroneous measure of the volume of the extracellular space. The decomposition of [3H]inulincan be minimized by storage of the [3H]inulin at -70°C. Another way to overcome this difficulty is to use the chemically stable [WrIEDTA (Stacy and Thorburn, 1966) in place of [3H]inulin. An alternative method for the separation of hepatocytes labelled with 4sCa2+from the incubation medium is direct centrifugation of the cell suspension (Assimacopoulos-Jeannet et al., 1977). This procedure involves a five- or ten-fold dilution of the cell incubation medium in a solution which contains 150 mM NaCl, 7% BSA and 2.5 mM LaCI,. The cells are centrifuged and the supernatant removed by aspiration. La3+ is added in order to displace 45Ca2+from the extracellular surface of the cells. Claret and his colleagues (Mauger et al., 1984) have employed a similar technique in which 4 mM CaCl, instead of LaCI, was used to displace extracellular Ca2+.In their experiments, the cells were separated from the incubation medium by filtration. However, this procedure has the potential to damage hepatocytes. An advantage of the silicone oil technique for the separation of hepatocytes from the incubation medium is that it allows a relatively large number of samples to be processed quickly in a short period of time. As indicated above, most experimenters have included La3+ or unlabelled Ca2+in the medium into which 45Ca2+-labelledcells are placed before centrifugation. It is assumed that the presence of La3+ or unlabelled Ca2+displaces 45Ca2+ from extracellular sites. These probably include binding sites for Ca2+on the extracellular domains of plasma membrane proteins and on phospholipids which compose the lipid bilayer of the membrane. However, the number of extracellular Ca2+binding sites on isolated hepatocytes appears to be small compared with the number present in the extracellular matrix of the intact liver (Barritt et al., 1981).

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9.3. Measurement of intracellular free Ca2+ using jluorescen t indicators 9.3.1. Available methods

The concentration of intracellular free Ca2+in hepatocytes has been measured using the fluorescent intracellular Ca2+ chelating agents quin2 (Charest et al., 1983; Thomas et al., 1984; Berthon et al., 1984) and fura2 (Monck et al., 1988; Kass et al., 1989; Kawanishi et al., 1989), the luminescent protein aequorin (Freudenrich and Borle, 1988), and by a null point titration method (Murphy et al., 1980). The method which has proved to be most useful and generally applicable involves the use of quin2. 9.3.2. Principle of the use of quin2for measurement of intracellularfree Ca2+

Quin2 is a fluorescent Ca2+chelator (Tsien, 1980) which has been successfully used to measure values of intracellular free CaZ+in hepatocytes in the absence of hormones, and changes in this parameter induced by hormones (Charest et al., 1983; Berthon et al., 1984; Thomas et al., 1984)or toxic agents (Moore et al., 1985). Quin2, which has four free carboxyl groups, can chelate one Ca2+.Upon binding of Ca2+at a pH of 7, the Caz+-quin2complex exhibits an approximately five-fold increase in fluorescence compared to the uncomplexed quin2 molecule. The free-acid of quin2 does not readily pass through the plasma membrane of cells. In order to introduce quin2 to the cytoplasmic space, cells are incubated for a period of time, usually 1-30 min, with the acetoxymethylester of quin2 (quinZAM), a lipidsoluble form of the chelator. Quin2-AM diffuses across the plasma membrane and accumulates in the cytoplasmic space, where it is hydrolysed by endogenous esterases to yield quin2 free acid. The liberated quin2 can then bind Ca2+present in the cytoplasmic space.

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Protocol 9.3 Measurement of intraceilulur free Cu2+ concentrations using quin2 (i) Loading of cells with quin2. After isolation, centrifuge the hepatocytes at 50 x g for 1 min and resuspend them in a total volume of 4 ml at a density of 2 x lo6 cells/ml in KHBT medium containing 0.5 mM CaCI,, 5 mM glucose, 50 pM EDTA and 1.5% gelatin. Incubate the cells at 37°C in 25-ml plastic vials in a shaking water bath (90 oscillations/min). The vials are gassed with carbogen. After a 10-min preincubation period that allows the cells to adjust their ion gradients, add 16 pl of quin2-AM to each vial from a stock solution of 25 mM quin2-AM in DMSO. The final concentration of quin2-AM is 100 pM. Incubate the cells for a period of 15 rnin in order to allow quin2-AM to enter the cytoplasmic space. (ii) Centrifuge the cells at room temperature at 50 x g for 1 rnin and resuspend them in 5 ml of the medium described above but containing no added Ca2f. This removes all extracellular quin2-AM which has not been taken up by the cells. Incubate the cells for a further 10 min at 37°C in order to allow endogenous esterases to complete the hydrolysis of quin2-AM. Centrifuge and resuspend the cells at a final density of 1-2 x lo6 cells/ml, in 3 ml of KHBT medium containing 1.3 mM Ca2+.This step removes extracellular quin2 and quin2-AM that have leaked out during the previous incubation. Incubate the cells for a further 5 min at 37°C in order to allow the ion gradients of the quin2-loaded cells to reach a steady state at the temperature that is to be employed for the measurement of q u i d fluorescence. In all incubations the cells are gassed with carbogen. Control cells which are not loaded with quin2 are prepared in the manner described above except that DMSO alone is added in place of quin2-AM. (iii) The quin24oaded cells (3 ml) are transferred to the cuvette of a spectrofluorometer (for example, a Perkin-Elmer LS-50 instrument) equipped with a temperature-controlled cuvette housing and a

226


magnetic stirrer which is used to maintain the cells in suspension. The cell suspension should be gassed with carbogen at a rate of 1 Vmin by a continuous flow of gas directed across the top of the cuvette from a thin polythene tube, Set the excitation and emission wavelengths at 340 nm and 490 nm, respectively. (iv) The effect of vasopressin on intracellular free CaZ+([Ca2+],)is investigated in the following way. Record fluorescence as a function of time for a period of 2 min in order to obtain a value for basal [Ca2+Ii.Add vasopressin (10 nM) and continue to record the fluorescence (F) which should show an increase as Ca2+enters the cells and is released from intracellular stores. At the end of the period of measurement (for example after 5 min) determine the amount of extracellular quin2 present by adding 50 pM MnZ+.This rapidly quenches the fluorescence of any extracellular quin2. The resulting decrease in fluorescence equals the fluorescence due to extracellular quin2. (v) At about 1 min after the addition of Mn2+, add 100 pM diethylene-triaminepenta-aceticacid which chelates Mn2+and allows the next step of the procedure, calibration of the fluorescence readings, to be undertaken. (vi) Add Triton X-100 (0.1% (v/v) final concentration) followed by CaC12 (2 mM final concentration) and continue to record the fluorescence signal. The detergent lyses the cells and allows all of the quin2 to be exposed to a saturating Ca2+concentration, so that a In order to stable maximal value for fluorescence is obtained (FmdX). calculate F,,,, two approaches have been used (Rink and Pozzan, 1985). Briefly, one can determine F,,, by quenching the fluorescence of all quin2 either by the addition of excess EGTA and alkalization of the medium, or by the addition of MnCl, (1 mM final concentration). The relative merits of the two approaches are discussed by Rink and Pozzan (1985). (vii) Repeat the entire procedure using control unloaded cells which have been incubated with DMSO in place of quin2, as described above, in order to obtain values for the autofluorescence of the cells-the

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fluorescence which is not due to quin2. The times of additions of the vasopressin and agents used for calibration should be exactly the same as those for the experiment with quin2-loaded cells. (viii) In order to calculate values of free Ca2+from the fluorescence data, the autofluorescence signal at each time point is subtracted from the fluorescence of quin2-loaded cells. The fluorescence due to extracellular quin2 is also subtracted. For the Perkin Elmer LS-50 spectrofluorometer this subtraction can be performed using an appropriate computer programme. The values of Fmaxand Fmin must also be corrected for autofluorescence. The value of [Ca2+Iiwhich corresponds to the fluorescence (F) at any given time is calculated using the values of F,,,, Fminand the following equation (Tsien et al., 1982).

in which K, is the dissociation constant for the binding of Ca2+to quin2 (Charest et al., 1983). The value of K, depends on the ionic environment of the quin2. We have used a value of 115 nM (Charest et al., 1983). 9.3.3. Technical considerations

Crofts and Barritt (1989) have reported that the amount of extracellular quin2 in suspensions of quin2-loaded hepatocytes is 15-20% of the total amount of quin2 present. This is high in relation to the values of extracellular quin2 cited for other cell types. The relatively high concentration of extracellular quin2 in hepatocytes is probably due to rapid leakage of quin2 out of the cells during their incubation (Crofts and Barritt, unpublished results). The leakage of quin2 from hepatocytes may be due to active secretion of the molecule by exocytosis or possibly to leakage through damaged plasma membranes. The high concentration of extracellular quin2 places some restriction on the use of this Ca2+indicator in the measurement of intracellular

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free Ca2+concentrations in hepatocytes. In particular, the number of experiments which can be performed with one batch of quin2-loaded cells is in practice limited because of quin2 leakage. Claret and his colleagues have successfully employed somewhat different conditions for the loading of cells with quin2 (Berthon et al., 1984). The differences include the use of a more complex incubation medium, as defined by Eagle (1 959), a much shorter incubation time (150 s) with quin2-AM and no further incubation (except incubation in the spectrofluorimeter cuvette) after washing of the cells. The ability to load cells with quin2 in the short time period may reflect the uptake of quin2-AM by pinocytosis and its rapid hydrolysis by esterases present in the hepatocyte. 9.3.4. Use of fura2

Although there are a number of advantages in the use of fura2 for the measurement of intracellular free Ca2+concentrations in cells (Cobbold and Rink, 1987) there are only a small number of reports of the successful use of fura2 in hepatocytes, compared with the number of reports in which quin2 has been used. Moreover, there is some indication that it is difficult to load fresh suspensions of hepatocytes with fura2. The groups of Orrenius, Monck and Tsien (Monck et al., 1988; Kass et al., 1989; Kawanishi et al., 1989) have measured intracellular free Ca2+concentrations in freshly-isolated hepatocytes loaded with fura2. The success experienced by these groups seems to be due, in part, to the use of much longer periods of incubation of the cells with fura2-AM than those employed for loading hepatocytes with quin2, the use of a substantial further incubation period to ensure complete hydrolysis of fura2-AM, and possibly to preincubation of hepatocytes before the addition of fura2-AM. Monck and Tsien measured intracellular free Ca2+concentrations in single cells rather than in cell suspensions. Tsien and his colleagues employed pluronic acid to enhance the loading of cells with fura2-AM and used a 40-min incubation of the

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cells with this ester. These workers also reported variations between different batches of furaZAM due to the presence of different amounts of unidentified contaminants. They comment that liver cells are more susceptible to the presence of these contaminants than at least one other cell type, the rat embryo fibroblast cell line REF52. After taking precautions to ensure the absence of contaminants and the complete hydrolysis of fura2-AM, Tsien and his colleagues were able to observe oscillations in intracellular free Ca2+concentrations induced by vasopressin and phenylephrine (Kawanishi et al., 1989).

9.4. Determination of rates of Ca'+inflow using glycogen

phosphorylase or fluorescent Ca2+indicators 9.4.1. Available methods

The measurement of rates of Ca2+inflow in isolated hepatocytes is important because a stimulation of the rate of Ca2+movement into cells is one of the two processes by which hormones such as phenylephrine, vasopressin and angiotensin I1 induce increases in the concentration of cytoplasmic free Ca2+(Barritt et al., 1981; Parker et al., 1983; Mauger et al., 1984). The other process is the release of Ca2+ from the endoplasmic reticulum, mediated by inositol 1,4,5-trisphosphate (Burgess et al., 1984; Joseph et al., 1984; Charest et al., 1985). Accurate measurement of rates of hormonal-stimulated Ca2+inflow in intact hepatocytes has proven difficult. This is because, in hepatocytes, there is a large basal inflow of Ca2+,independent of hormonal activation, and also because it has been difficult to develop a method which distinguishes inflow of Ca2+across the plasma membrane from other intracellular movements of Ca?+. Four main methods have been employed to measure rates of Ca2+ inflow across the plasma membrane. These are: (a) the use of 45Ca2+ under steady-state conditions, described earlier; (b) measurement of the increase in intracellular free CaZ+concentration as determined by


230

intracellular quin2 following the addition of Ca2+to cells previously incubated in the absence of added extracellular Ca2+;(c) measurement of the rate of quenching of the fluorescence of intracellular quin2 following the addition of Mn2+;and (d) measurement of the initial rate of activation of glycogen phosphorylase following the addition of extracellular Ca2+to cells previously incubated in the absence of added Ca2+. 9.4.2. Measurement of the stimulation by vasopressin of Ca2+inflow using glycogen phosphorylase

The measurement of glycogen phosphorylase activity as an estimate of Ca2+ inflow is one of the most convenient assays for this parameter. It has the advantage that a large number of experimental conditions can be tested relatively easily. The method depends on the ability of Ca2+to activate glycogen phosphorylase kinase which, in turn, phosphorylates and activates glycogen phosphorylase (Assimacopoulos-Jeannet et al., 1977; Blackmore et al., 1978). The covalent modification of glycogen phosphorylase is stable in the presence of inhibitors of glycogen phosphorylase phosphate phosphatase, such as fluoride. The principle of the method is as follows (Binet et al., 1985; Cooper et al., 1985; Hughes et al., 1986). Hepatocytes are incubated in the absence of extracellular Ca2+.Ca2+are added and samples of the cell suspension removed for the assay of glycogen phosphorylase activity. Enzyme activity is measured by the procedure of Hutson et al. (1976) or Blackmore and Exton (1985). 9.4.3. Major reagents 1.

Cell disruption medium. This contains 500 mM sucrose, 140 mM NaF, 30 mM Na2EDTA, 140 mM Na,-&glycerophosphate and 5 mg/ml cysteine-HCI. (NaF and EDTA are added to inhibit phosphoprotein phosphatase activity.) The pH is adjusted to 6.2

Ch. 9.

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23 I

using KOH or HCI, as necessary. The cysteine is added just before use. The other components of the buffer can be made up and stored at M ” C . In our experience the disruption buffer (without cysteine) is stable for several months. 2. Glycogen phosphorylase assay mixture. This contains 1 mM caffeine, 2% glycogen, 100 mM glucose-1-phosphate, and 75 pl (7.5 pCi, 50 nmol) of [ U-14C]glucose-l-phosphate (e.g. Amersham) per 30 ml. The pH is adjusted to 6.1 with KOH or HCI. The assay mixture can be stored at -20°C for several months. This is best done in small aliquots, each of which is sufficient for a single experiment. Some investigators believe that only re-crystallized glycogen should be used for this assay. However, in our experience Sigma Type VII glycogen from oyster is satisfactory without re-crystallization.

Protocol 9.4 Technique for assay of cellular glycogen phosphorylase (i) After isolation, hepatocytes are suspended in incubation buffer at a density of 6 x lo6 cells/ml and are kept on ice. Incubations are conducted in cylindrical glass incubation chambers in which the cells are maintained in suspension by a magnetic stirrer and gassed, as described for Protocol 9.1. (ii) Begin the incubation by adding 2 ml of cell suspension to 4 ml of Ca2+-freeKHBT medium. After an equilibration time of 15 min, add vasopressin (10 nM final concentration) and continue the incubation for a further 10 min to allow the release of Ca” from intracellular stores (thought to be located in the endoplasmic reticulum) into the cytoplasmic space. At the end of this period remove a sample of the suspension (100 pl) and mix it with homogenization buffer as described below. Then add CaCl, (1.3 mM final concentration) and remove further samples of the incubation medium (100 pl) at 15-s intervals for a period of 90 s. A similar incubation, in which an equal

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volume of 150 mM NaCl is added in place of vasopressin, is conducted in order to gain an estimate of the rate of activation by Ca2+ of phosphorylase in the absence of vasopressin. (iii) Place each sample of incubation medium removed from the incubation vessel into 260 pl of ice-cold disruption buffer contained in a plastic tube and immediately immerse the contents of this tube in liquid N,. The sample should remain frozen until the assay of glycogen phosphorylase activity is performed. If this is not done immediately, the tubes can be stored frozen at -70°C. It is advisable to perform the assay of glycogen phosphorylase activity within 24 h as there is some inactivation of the enzyme, even at low temperatures. (iv) Assay of glycogen phosphorylase activity. Glycogen phosphorylase activity is assayed in the direction of glycogen synthesis by measuring the incorporation of radioactivity from [U-'4C]glucose-l-phosphateinto glycogen. Each cell extract is assayed in duplicate. The frozen cell extract is allowed to thaw and is then mixed. Add 50 pl of the cell extract to 50 pl of assay mixture in a glass tube, mix, and incubate at 30°C for 15 min. Stop the reaction by removing 50 pl of this mixture and spotting it onto a 20-mm square of Whatman No. 3 filter paper (marked with a pencil) that is then dropped into a beaker that contains 60% (v/v) ethanol in water to precipitate the radioactively-labelled glycogen. Excess [U-'4C]glucose-l-phosphate will disperse in the ethanol/water. Prepare blank samples by mixing 50 pl of disruption buffer with 50 pI of assay mix, spot 50 pl of this mixture onto filter papers and place in ethano1:water. Total radioactivity is measured by spotting 50 pl of the assay mixture onto filter papers. These are not placed in ethanol but are allowed to dry thoroughly before measurement of radioactivity. (v) When all the reactions have been stopped, wash the filter paper squares (which contain [I4C]glycogen and residual [i4C]glucose-1phosphate) in the 60'%,ethanol for 40 min while stirring with a magnetic stirrer and bar. After this time, replace the 60% ethanol with a fresh

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mixture of 60% ethanol and continue washing for a further 40 min. Repeat this step once more. Remove the ethanol and wash the filter paper squares for about 1 min in acetone. The papers are then allowed to dry in air. The radioactivity on each filter paper is measured by placing the filter paper into a 5-ml plastic scintillation vial with 4 ml of scintillation fluid. The activity of glycogen phosphorylase in a given sample is calculated from the amount of radioactivity present in [14C]glycogen, and the specific activity of the ['4C]glucose-l-phosphate after subtraction of blank values. Enzyme activity is expressed as pmol glucose- 1-phosphate incorporated into glycogenlmin per g wet weight.

9.4.4. Comments on the method There are some reservations about use of glycogen phosphorylase for the measurement of CaZ+inflow. Firstly, Ca2+ that enters the cytoplasmic space may be taken up by intracellular organelles, such as the endoplasmic reticulum and mitochondria. For this reason, rates of CaZ+ inflow obtained using this method are likely to be an underestimate of the true rates of Ca2+ inflow. Secondly, the experiments are performed with hepatocytes initially incubated in the absence of extracellular CaZ+so that the observed rates of CaZinflow may not reflect those present at normal extracellular Ca2+concentrations. Thirdly, other factors such as cyclic AMP can modify the activity of glycogen phosphorylase. Notwithstanding these reservations, the results obtained with glycogen phosphorylase for the estimation of CaZ+inflow are similar to those obtained using other methods.

9.4.5. Measurement of Caz+ inflow with Mnz+ Measurement of the uptake of MnZ+by cells loaded with quin2 or fura2 is potentially one of the best methods currently available for

234


the measurement of Ca2+inflow to hepatocytes. The basic assumption of the method is that Mn2+moves through the same channel as Ca2+ (Crofts and Barritt, 1990). The method depends on the observation that, whereas Ca2+ions increase the fluorescence of these dyes, Mn2+quenches the fluorescence (Hesketh et al., 1983; Arslan et al., 1985; Cobbold and Rink, 1987). One advantage of the fura2 assay is that by using an excitation wavelength of 360 nm, the quenching of fura2 by Mn2+can be followed without the complication that may be caused by changes in fluorescence due to variation in Ca2+concentration (Sage et al., 1989). This is because at the isosbestic wavelength of 360 nm, the fluorescence of fura2 is independent of Ca2+concentration (Grynkiewicz et al., 1985). Because it is difficult to load isolated hepatocytes with fura2, the following Protocol describes the measurements of Mn2+inflow, using cells loaded with quin2.

Protocol 9.5 Measurement of the vasopressin-stimulated rate of Mn2+inflow (i) Hepatocytes are loaded with quin2 (3 ml total volume), and incubated in Ca2+-free KHBT medium in the cuvette of a spectrofluorometer, as described in Protocol 9.3. The excitation wavelength is 340 nm and the emission wavelength 490 nm. (ii) After an incubation time of 2 min to allow establishment of a baseline fluorescence value, add vasopressin at a final concentration of 10 nM. Fluoresence is recorded for a further 4 rnin to allow completion of the release of Ca2+from intracellular stores. This is observed as a transient increase in fluorescence. (iii) Now add MnCl, (100 pM final concentration) and measure the fluorescence for a period of 2 min. Addition of MnCl, results in an initial rapid decrease in fluorescence followed by a slower decrease which is maintained for at least 2 min. The first phase is due to the quenching of extracellular quin2 by Mn2+.The second phase is due to the quenching of intracellular quin2 by Mn2+.The slope of the second phase, measured as the change in fluorescence as a function of time, represents the rate of entry of Mn2+into the hepatocytes.

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(iv) The experiment is repeated in the same manner but without the addition of vasopressin. Subtract the rate of Mn2+-inducedquenching of fluorescence observed in the absence of vasopressin from that observed in the presence of the hormone. This difference gives the rate of vasopressin-stimulated entry of MnZ+into the cytoplasmic space of the liver cell. 9.4.6. Comments on the technique As stated above, quin2 leaks out of hepatocytes rapidly compared with other types of cells so that the extracellular component of Mn2+ quenching of quin2 fluorescence is relatively large. One of the disadvantages of the use of the Mn2+quenching technique is that it is not easy to obtain a quantitative measure of the rate of Mnz+ inflow in cells loaded with quin2. This difficulty may be overcome in the future by the use of fura2 in place of quin2. An assumption of the Mn2+quench method is that most of the MnZ+that enters the cell binds to quin2. While this may be valid at low MnZ+ concentrations it is probably not correct at high extracellular MnZ+ concentrations (Crofts and Barritt, 1990).

9.5. Assessment of available techniques for the measurement of Ca*+in hepatocytes The methods described are those that are most reliable for the measurement of pools and fluxes of Ca2+in hepatocytes. However, as indicated in discussion of these methods, there are reservations about interpretation of the data obtained using some of the procedures. The measurement of total cell Ca2+by atomic absorption spectroscopy, the concentration of free Ca2+in the cytoplasmic space using quin2, and Caz+inflow across the plasma membrane using the quenching of intracellular quin2 by MnZ+are reliable. Difficulties of interpretation are present in the measurement of the amount of CaZ+inintracellular organelles and in fluxes of Caz+between the extracellular fluid, the

236


cytoplasmic space and intracellular organelles. These difficulties are chiefly due to the complexity of the structure of the cell and the lack of methods for the direct measurement of CaZ+in intracellular organelles and fluxes across specific membranes. Such problems are not restricted to the study of hepatocytes. In future they may be solved by the use of a more complex array of fluorescent intracellular Ca2+chelators and image analysis techniques.

CHAPTER 10

Hepatocyte isolation for primary culture and methods for non-adherent culture

10.1. Introduction For many investigations of hepatocyte function, the maintenance of freshly-prepared cell suspensions for 1-2 h is adequate. On the other hand for investigations of hepatocyte gene expression and its control, for studies on growth control and for areas of investigation such as toxicology and carcinogenesis, the ability to maintain functional hepatocytes for many hours, days or even months is important. Attempts to culture liver tissue date from early this century (Carrel, 1911). Liver cell-lines or strains, derived from normal liver or hepatomas, have been used for many years to provide cells that possess some of the properties of adult hepatocytes (Watanabe, 1970; Alexander and Grisham, 1970; Grisham 1979; Furukawa et al., 1987). However, in general, these propagatable cell-lines exhibit few of the specialized metabolic functions characteristic of normal adult mammalian liver in vivo. One aim of primary hepatocyte culture (in which the cells in culture are derived directly from disaggregated liver) is to provide, for study under defined conditions in vitro, cells that do exhibit a wider range of functions characteristic of hepatocytes in intact liver. Prior to 1972, primary cultures from embryonic chick liver had been used

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extensively by Granick (1966) and others to study the haem biosynthetic pathway; and human or rat foetal liver cultures exhibiting some differentiated functions had been described (Bissell and Tilles, 1971; Leffert and Paul, 1972). However at that time there was no culture system suitable for broad studies on the differentiated functions of adult mammalian hepatocytes. The uncertainties and frustrations in the art of liver cell culture in 1972 are aptly summarized by Potter (1972). About that time a number of laboratories began to exploit the highyield preparations of hepatocytes, obtained by the collagenase perfusion procedure of Berry and Friend (1969), for studies on gene expression extending over several hours, with cells maintained in suspension culture (e.g. Berg et al., 1972; Edwards and Elliott, 1974; Jeejeebhoy et al., 1975). Methods for suspension culture, with more recent refinements, continue to have limited application. However the major development of hepatocyte primary culture began with the report by Bissell et al. (1973) of methods for monolayer culture of hepatocytes prepared by collagenase perfusion. In the &4i days after plating, the cells exhibited metabolic activities such as albumin synthesis and secretion, gluconeogenesis from 3-carbon precursors, glycogen synthesis and responses to insulin and glucagon that resembled liver in vivo at least qualitatively. Independently, Bonney et al. (1974) developed a similar primary culture system and demonstrated tyrosine aminotransferase inducibility by glucocorticoids for 4 or more days in culture. In summarizing a 1974 conference on liver cell culture, Farber (1975) concluded that a good beginning had been made on maintaining differentiated non-proliferating liver parenchymal cells under in vitro conditions, while noting that attempts to grow cells from liver apparently resulted in breakdown of the differentiated state. By the time of the next major conference on liver cell culture in 1979 there were numerous groups around the world utilizing hepatocyte primary culture and many of the limitations of early culture systems for maintaining differentiated functions were recognized (Borek and Williams, 1980). In the subsequent decade enormous

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effort has been devoted to refining culture systems to achieve two major aims. One aim has been longer-term survival of cells with the retention of various differentiated functions, while a second objective has been to achieve proliferation of hepatocytes in vitro, at least comparable with that observed in vivo after partial hepatectomy. While efforts have been made to preserve both normal differentiation and the potential for growth in a common culture system, an emerging view is that the in vitro conditions required for optimal preservation of differentiated function are distinctly different from those required to promote cell proliferation, at least for adult hepatocytes. This Chapter, the first of two on primary culture, begins with discussion of those aspects of liver cell isolation especially relevant to maintaining well-defined populations in culture for periods of several hours, days or weeks, with emphasis on isolation of adult rat hepatocytes. Methods for maintaining hepatocytes in single-cell suspension for periods of several hours, together with advantages and limitations of such systems, are discussed. Longer-term cultures of non-adherent spheroids are also briefly described. The following Chapter considers monolayer culture methods.

10.2. Alternatives to perfusion with collagenase in isolating cells for culture The factors influencing successful isolation of hepatocytes have been considered in detail in Chapters 2 and 3. In general, the procedures optimal for preparing hepatocytes for short-term studies in suspension are also appropriate for longer-term studies in primary culture. The great majority of laboratories isolating hepatocytes for primary culture have used variations of the collagenase perfusion procedures discussed in Chapter 2. This Section briefly considers alternatives. In general, methods which involve essentially non-perfused preparations (Leffert et al., 1977) or perfusion with enzymes other than collagenase such as dispase (Wahid et al., 1984) or trypsin (Miyazaki et

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al., 1984) result in lower yields of hepatocytes and greater contamination with non-hepatocytes, with subsequent outgrowth of clear epithelial cells in culture. One interesting variation on collagenase perfusion, recently applied for cell culture by Wang et al. (1985) and Meredith (1988) is based on the EDTA perfusion procedure of Berry et al. (1983), described in Section 3.3. Although the cell yields from EDTA procedures may be lower than from collagenase perfusion and the initial cell isolate of lower quality, substantial yields of intact hepatocytes, apparently free of other cell types, can be obtained after centrifugation through Percoll to remove damaged cells. When plated at high density on plastic (Wang et al., 1985)or collagen-coated dishes (Meredith, 1988), confluent monolayers form within a few hours. In short-term studies, Wang et al. (1985) demonstrated substantial rates of albumin synthesis. Meredith (1988) reported that monolayers were preserved with little change in morphology for up to 3 weeks. Isolated cells prepared by EDTA perfusion preserved constant total cytochrome P-450 and glutathione levels for 5 days and there was no induction of y-glutamyl transpeptidase, in contrast to findings for cells prepared with collagenase and maintained under identical conditions (Meredith, 1988). Appearance of y-glutamyl transpeptidase apparently reflects progressive change in the differentiated state of adult rat hepatocytes in culture (Sirica et a]., 1979; Edwards et al., 1987b). Although ‘selection’ of undamaged cells by Percoll centrifugation may have contributed to retention of normal function (see below), on the basis of the parameters studied by Meredith, it appears that cell isolation with EDTA (that is, in the absence of collagenase) may have advantages for the subsequent preservation of differentiated function under relatively simple culture conditions. It would be most interesting if this is confirmed by more extensive studies. Cells prepared with collagenase are known to internalize measurable levels of enzyme (Guzelian and Diegelmann, 1979). If collagenase or other contaminating proteases are exerting some intracellular effect, or alternatively removing or damaging cell-surface components relevant to long-term function, it may at least be desirable to minimize the period of exposure to collagenase where this is used for cell isolation.

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10.3. Choice of collagenase perfusion isolution procedure The cell preparation methods used by most laboratories concerned with hepatocyte culture have been based on the two-step collagenase perfusion procedure of Seglen (1976) (Section 2.3). although variations of the one-step procedure in Protocol 2.2 have also been used successfully (e.g. Flaim et al., 1985). Almost every laboratory has ils own modification of these methods and undoubtedly most of these give satisfactory preparations. I n general, the aims should be to obtain the maximum yield of hepatocytes while minimizing biochemical changes in the cells during isolation, minimizing contamination with non-hepatocytes and maximizing the proportion of cells capable of attaching and surviving in culture in the final preparation - all under sterile conditions, There are only a few published comparisons of different collagenase perfusion isolation procedures in relation to subsequent behaviour of cells in culture (Williams et al., 1977; Kreamer el al., 1986). The detailed choice of isolation method (e.g. age or sex of rats, extent of measures to eliminate non-hepatocytes) will depend to some extent on the phenomena under investigation. The following Sections list some factors which should be considered. 10.3.1. Age of rats used us donors

The age of rats used for cell isolation may be important for some applications. Inducibility of tyrosine aminotransferase in hepatocyte suspensions (Britton et al., 1976) or levels of DNA repair in monolayers (Sawada and Ishikawa, 1988) were independent of the age of the donor rat. However, for studies on growth and events related to the cell cycle it appears to be important to use rats less than 3 months old (Sawada and Ishikawa, 1988; Sawada. 1989). In our hands, young rats (150-200 g) give better yields of trypan blue-excluding cells per g liver and these attach better to collagen-coated dishes than do cells from rats weighing more than 250 g.

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10.3.2. Isolation media

For culture experiments, a modification introduced by Bonney et al. (1974) and subsequently used by some laboratories including our own (Whiting and Edwards, 1979; Buttenvorth et al., 1987), is to employ complex media rather than buffered salts solutions during isolation of hepatocytes, at least during the second perfusion step with collagenase and for washing the cells. Since cells after isolation survive better in complex media, with both amino acids and insulin contributing to preservation of nitrogen balance (e.g. Dickson and Pogson, 1977; Schwarze et al., 1982), it appears reasonable that the presence of amino acids and other components during isolation may help preserve normal biochemical properties. However, there does not appear to be any direct evidence that complex media are important for cell isolation in relation to subsequent culture, and simpler bufferedsalts solutions may well be adequate. 10.3.3. Potential contamination by non-hepatocytes

The Berry and Friend (1969) isolation procedure and most of its variations give high yields of hepatocytes with relatively complete removal of other cell types. For short-term metabolic studies, contamination with 2-5% non-hepatocytes is not a serious problem. However, in culture experiments, particularly where cells are maintained for longer periods or proliferation is under investigation, potential outgrowth of a small number of contaminating cells becomes an important issue. Collagenase disaggregates the lobular parenchyma of the liver, releasing hepatocytes and up to 80%of the non-hepatocytes. The nonhepatocyte fraction in the initial cell isolate contains about 70%)endothelial cells, 20% Kupffer cells and 10% lipocytes (fat storing, Ito or stellate cells) (Friedman and Roll, 1987; Smedsrod et al. 1990). All these cell types will attach to plastic dishes or other substrata and survive in culture for several days at least. Kupffer cells apparently do not proliferate (Friedman and Roll, 1987) but in media containing

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serum and/or growth factors a subpopulation of endothelial cells grow in culture (Irving et al., 1984; Morin and Normand, 1986). Similarly, in the presence of serum or growth factors, and on plastic or collagencoated dishes (but not on basement membrane matrix), lipocytes are ‘activated’ to myofibroblast-like cells which proliferate (Friedman et al., 1989; Bachem et al., 1989). Although less well-characterized, the initial cell isolate also contains substantial numbers of fibroblasts and cells that give rise to colonies of clear epithelial cells in culture (e.g. Furukawa et al., 1987). Both of these cell types proliferate rapidly in serum-containing media. Even in serum-free medium we have observed fibroblast-like cells growing in hepatocyte primary cultures (Edwards and Lucas, 1982). The portal tracts in liver are poorly disaggregated by perfusion with collagenase. Non-hepatocytes in the biliary tree and connective tissue associated with the portal tracts thus remain in the undissociated tissue. In this fraction, bile duct epithelial cells are the most numerous. These cells are released only by vigorous shaking with collagenase medium (Mathis et al., 1989) or further digestion with trypsin or pronase (Kumar and Jordan, 1986; Parola et al., 1988). Isolated bile ductular epithelial cells will attach to culture dishes but apparently most cells do not proliferate (Mathis et al., 1989). The biliary and connective tissue, left relatively intact by collagenase, presumably also contains the majority of fibroblasts and cells capable of forming clear epithelial cell colonies, since the numbers of these growing out in cultures is greatly increased by perfusion with other proteases such as dispase (Furukawa et al., 1987; Wahid et al., 1984) or trypsin (Miyazaki et al., 1984) or both (Marceau et al., 1986b), used in addition to, or in place of, collagenase. 10.3.4. Elimination of non-hepatocytes All of these non-hepatocyte cell types are potential contaminants in primary cultures, although most are eliminated in standard methods for preparing hepatocytes. The use of the relatively gentle collagenase

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perfusion procedure avoids contamination with bile duct epithelial cells and apparently minimizes release of fibroblasts and clonigenic epithelial cells. All of the non-parenchymal cells, with the possible exception of some fibroblasts, are smaller than mature hepatocytes. The smallest diploid hepatocytes from fed rats have a mean diameter of 17-18 pm, whereas Kupffer, endothelial and bile duct cells have diameters of 11, 8 and 6 pm, respectively (Klose et al., 1989; Parola et al., 1988). Findings of Marcedu et al. (1980) suggest clonigenic epithelial cells and fibroblasts have diameters of about 14 pm and 20 pm. Because of these size differences, the use of low centrifugal speeds (50 x g or less) and short periods of centrifugation in the final washing of hepatocyte preparations results in separation of the majority of hepatocytes in the pellet from most of the non-hepatocytes (as well as lighter cell debris and damaged hepatocytes) in the supernatant. The comparison of four isolation methods by Kreamer et al. (1986) suggests that contamination with non-parenchymal cells is less with methods involving shorter exposures to collagenase and extensive washing with low-speed centrifugation at 50 x g or less. Some laboratories allow hepatocytes to settle for 5-10 min at 0 4 ° C under gravity (Butterworth et al., 1987).Our experience (Edwards and Lucas, 1982), in line with other laboratories (e.g. Seglen, 1976), is that contamination with non-hepatocytes after washing as in Protocol 10.1 is between 2 and 5%. After a standard hepatocyte isolation the major contaminants are likely to be endothelial and Kupffer cells with a few lipocytes, fibroblasts and clonigenic epithelial cells. Because of their potential for relatively rapid growth, the cells most likely to complicate studies in culture are fibroblasts and the clonigenic epithelial cells. The nature of the latter cells has been the subject of lengthy debate. These cells may be immature cells derived from bile duct epithelium (Grisham, 1983).They may be related to epithelial progenitor cells found in foetal liver, which are capable of differentiation along hepatocytic or bile ductular lineages (e.g. Germain et al., 1988; Sell, 1990), and to putative stem cells in adult liver, thought to be associated with the canals of

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Hering, particularly around the portal spaces (Fausto and Mead, 1989). Alternatively, Marceau et al. (1986b) suggested some may derive from Glisson's capsula. There are a variety of possible approaches to reducing nonhepatocyte contamination further. The findings that Kupffer cells attach rapidly to plastic and that endothelial cells attach only slowly to protein substrata (Smedsrod and Pertoft, 1985) in contrast to intermediate rates of hepatocyte attachment suggest that differential attachment could be used to reduce Kupffer and endothelial cell contamination. As discussed in Section 1 1.4.1 1, appropriate serumfree media can minimize growth of fibroblasts and clear epithelial cells. The most comprehensive approaches for purification utilize the different size and/or density of hepatocytes, compared with most nonhepatocytes, to separate hepatocytes by centrifugal elutriation (e.g. Irving et al., 1984) or by centrifugation in Percoll. The Percoll procedure of Kreamer et al. (1986) is relatively rapid and simple (Protocol 2.5), depending on the use of a low centrifugal speed to sediment hepatocytes ( p 1.065-1.085 g/ml according to Singh et al., 1983) through Percoll, p 1.06 g/ml. The less dense endothelial cells ( p < 1.018 g/ml) and Kupffer cells ( p 1.018-1.033 g/ml) and probably other cell types are retained, along with damaged hepatocytes and debris, as an upper flocculent layer in the Percoll. After this procedure nonhepatocyte contamination is 1%) or less. 10.3.5. Eliminution of damuged hepatocytes

AS discussed previously, a standard procedure for hepatocyte isolation removes most of the damaged hepatocytes. A short period of shaking with collagenase after liver disaggregation helps break up damaged cells (Protocol 2.2) but was found by Williams et al. (1977) to reduce subsequent rates of attachment of the remaining intact cells to plastic dishes. After washing steps, preparations with more than 95% of cells excluding trypan blue may be obtained if care is taken to minimize the time and speed of centrifugations. However, the percentage of in-

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tact cells obtained in routine laboratory preparations may often be somewhat lower (e.g. Kreamer et al., 1986). With relatively short attachment times in monolayer culture experiments, most of the residual damaged cells fail to attach and are removed with a change of medium. However, even the short-term presence of these damaged cells may have adverse effects. In our experience the presence of such cells reduces attachment efficiency of the intact cells in the preparation. This can be a particular problem where a high plating density is used with the aim of forming a denselypacked monolayer. Studies in which long-term behaviour of cultures was compared after isolation with or without a Percoll purification step suggest that overall preservation and function of the initial intact cells is significantly improved by relatively complete removal of damaged hepatocytes prior to establishing suspension or monolayer cultures (Dalet et al., 1982; Kreamer et al., 1986). Thus, for at least some applications, centrifugation of cells through Percoll, as in Protocol 2.5, may be valuable both for minimizing non-hepatocyte contamination and maximizing the proportion of intact hepatocytes. As a general rule, the isolation procedure chosen should yield >90% intact cells as assessed by trypan blue staining.

10.4. Preserving sterility of cell preparations For maintenance of hepatocytes in complex media for more than 1-2 h, at least some measures to minimize growth of bacteria, yeast and other fungi are necessary. For monolayer or suspension experiments lasting up to about 24 h, it is probably sufficient to carry out the final washes during hepatocyte isolation with sterile medium, then incubate the cells in sterile vessels with sterile medium containing antibiotics. Where cultures are maintained for longer periods, sterile precautions are necessary at all stages of cell isolation and subsequent maintenance. Although antibiotics delay the appearance of obvious contamination, failure of sterile precautions during cell isolation usually leads to frustrating loss of cultures after 2-3 days.

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10.4.1. Handling of non-sterile animals

Cell isolation from non-sterile animals obviously involves some compromises in sterile technique. Experience in many laboratories is that perfusions can be conducted on the bench in a clean, draught-free area without using sterile rooms or cabinets; and that in situ perfusion, despite somewhat greater risks of contamination than with isolated perfused livers, can routinely yield sterile preparations. Commonsense precautions can minimize contamination during the perfusion stage. The abdomen of the animal is swabbed with 70% (v/v) ethanol and a patch of skin removed above the abdomen and rib-cage to avoid any contact between skin and liver and to minimize the danger of loose hairs adhering to the liver. Sterile gauze can be placed over the intestines and when positioning liver lobes during cannulation. The liver should be handled with sterile forceps. Care should be taken to avoid any damage to the stomach or intestines during removal of the liver after the perfusion. Finally, washing the intact liver with sterile medium prior to disaggregation helps minimize any transfer of contaminating organisms into the sterile media used in the final steps of cell isolation. 10.4.2. Sterilization of equipment and media

In general all instruments used in surgery must be sterile: immersion in 70% ethanol is the most convenient method of sterilization. Glassor plastic-ware, tubing (autoclavable-latex or silastic) and gauze used during cell isolation are in general sterilized by autoclaving. Short stretches of tubing within a peristaltic pump may be left in place and sterilized before and after each perfusion by passage of 70%)ethanol. Some thought in designing perfusion apparatus can help minimize contamination problems. Usually arrangements for oxygenating media and for equilibrating bicarbonate-buffered media with CO, are required. Our experience is that gas direct from cylinders can be passed through tubing that has been autoclaved prior to setting up the perfusion system and subsequently left in place over several months. However, tubing in contact with perfusion solutions must be sterile.

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To equilibrate solutions with carbogen, we use a set of rubber stoppers fitted with glass inlet and outlet tubes that are kept sterile prior to use, in autoclave bags. These provide a sterile connection through which gas can be bubbled into medium in a matching sterile flask. During the post-perfusion steps of cell isolation, the procedure is simplified if HEPES-buffered solutions which do not require gassing are used. The step at which disaggregated liver is filtered through gauze also requires some self-contained (autoclavable) arrangement to hold the gauze in place, and allow washing of the residue and recovery of the filtrate with a minimum of handling. All the solutions used during cell isolation must be sterilized, usually by passage through 0.2 pm-pore filters. Collagenase solutions usually contain undissolved material that rapidly blocks filters. To provide a convenient source of sterile collagenase for frequent cell preparations, a ten-fold concentrated stock solution of collagenase can be prepared in Ca2+-freeperfusion medium, centrifuged at 14,000 x R to remove particulate material and then filtered. We prepare 1-1 batches and use pressure filtration through a 0.2 pm pore size, 142 mmdiameter filter. The stock may be stored at -20°C for up to 3 months in aliquots sufficient for one or two perfusions. Once thawed, the collagenase should be used within 24 h.

10.5. A procedure for heputocyte isolution Many of the detailed procedures relating to hepatocyte isolation are given in Protocol 2.2. In the protocol below, based on that used in our laboratory for several years, only those points which differ from Protocol 2.2 are emphasized. The perfusion and wash solutions described are based on the commercially-availablecomplex culture media S-77 (which is Ca'+-free, available from Sigma) and Williams' Medium E (ICN Biomedicals).Alternative (and possibly less complex) formulations may give satisfactory results. The method below reflects the historical development of isolation procedures used in laboratories

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concerned with hepatocyte primary culture and resembles a protocol recently suggested by a panel from several such laboratories (Butterworth et al., 1987). There is no definitive evidence, however, that the variations from Protocol 2.2 (other than the need for sterility) are crucial in preparing cells for culture. 10.5.1. Solutions

Perfusion medium. Prepare Swim's S-77 medium (with 2.24 g NaHCO,/I) supplemented with O.I'% BSA, fraction V, and phenol red (10 mg/l), filter through a 0.2 pm filter and store at 0 4 ° C . On the day of the experiment add antibiotics and insulin in the following final concentrations: penicillin (60 pglml), streptomycin sulphate ( 100 pg/ml) and insulin, 0.3 pM.The antibiotics and insulin are added from sterile 200-fold concentrated stock solutions in saline which are stored frozen in small volumes. During the perfusion. a pH of 7.4 and adequate oxygenation are maintained by bubbling carbogen through the medium. First-stage medium. To 180 ml perfusion medium add 1.8 ml of a sterile 0.05 M stock solution of EGTA, adjusted to pH 7.4 with NaOH and stored at 0 4 ° C . Equilibrate at 37°C in the perfusion reservoir. Bubble with carbogen 10 min prior to, and during, perfusion. Second-stuge medium. To 135 ml perfusion medium add I ml of a sterile 0.6 M stock solution of CaCI, stored at 0 4 ° C plus 15 ml of sterile ten-fold concentrated collagenase stock solution (4 mg per ml in S-77/albumin/phenol red, stored at -20°C in small aliquots). Equilibrate at 37°C and bubble with carbogen for 10 min prior to use and during perfusion. Wush medium. Prepare Williams' Medium E without bicarbonate, supplemented with 15 mM HEPES and adjusted to pH 7.35 at 37°C. filter through a 0.2-pm membrane and store at 0 4 ° C . On the day of the experiment, supplement with antibiotics and insulin as for the perfusion medium. Keep at 0 4 ° C . Disuggregurion medium. To 36 ml wash medium, add 4 ml of tenfold concentrated collagenase. Equilibrate at 37°C.

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Protocol 10.1 Method for isolation of hepatocytes for primary culture (i) To prepare the anaesthetized rat for surgery, swab the abdomen liberally with 70"hethanol. Before opening the abdominal cavity, cut away a central strip of skin about 3 cm wide, from the pubis to about 3 cm above the sternum. (ii) After cannulation of the portal vein, perfuse the liver with the first-stage medium at 20 ml/min. The liver should clear immediately to an even tan colour. Increasing the flow rate briefly may help to achieve this. Perfusate is allowed to escape via the inferior vena cava, cut below the kidney. When the outflow cannula has been inserted and connected to outflow tubing, tie off the lower end of the vena cava near the kidney and allow perfusate from the outflow tubing to run to waste into a sterile, foil-covered beaker. Cover the liver with sterile plastic film or cotton gauze to minimize cooling due to evaporation. (iii) When 180 ml of first-stage medium has been drawn from the reservoir, allow a small amount of air into the tubing (subsequently caught in the bubble trap) then add the second-stage medium, preequilibriated with carbogen at 37"C, to the reservoir. Manipulating the end of the outflow tubing so that it remains sterile, transfer the outflow tube to the reservoir so that the second-stage perfusate (containing 0.04% collagenase plus 4 mM CaCI,) recirculates. Turn up the flow rate to 30 ml/min. (iv) Continue the second-stage, recirculated perfusion for about 15 min. The exact period required varies by a few minutes with different batches of collagenase. When the liver is adequately perfused, gentle pressure on the underside of one of the large lobes with blunt forceps should leave a visible depression that fills with perfusate beneath the capsule. (v) At this point, turn off the pump, remove the portal cannula and carefully cut the liver away from diaphragm, veins and other tissue

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and place in about 20 ml warm wash medium in a sterile beaker. Transfer the beaker to a laminar flow cabinet for further manipulations. If reasonably intact, the liver can be held above the beaker and washed with additional wash solution, then transferred to a second sterile beaker containing the 40 ml warm ‘disaggregation medium’. The liver is broken up with scissors and cells released with a blunt spatula or dog comb. Ideally, only pale fibrous connective and ductular tissue should remain after this treatment. If necessary to complete disaggregation, the complete mixture can be shaken for 5-10 min at 37°C in a sterile 250-mI flask at 8&90 cycles/min. Where good dissociation has been achieved by the perfusion, the exposure of cells to collagenase can be minimized by proceeding without this further incubation. (vi) Pass the disaggregated liver cells through nylon mesh ( = 200 pm pore size) and wash the residue by pouring a further 100 ml cold wash through the filter. (vii) Transfer the filtrate to 3 or 4 sterile 50-ml plastic screw-cap centrifuge tubes and centrifuge for 3 min at 50 x g . (viii) Aspirate the supernatants, gently resuspend the pellets in each tube in 40 ml fresh cold wash medium using a sterile wide-bore pipette and centrifuge for 3 min at 50 x g . Repeat this sequence for a third wash. (ix) After the third centrifugation the supernatants should be relatively clear. (If not, wash the pellets again.) Remove the supernatants as completely as possible and, using a wide-bore pipette, gently resuspend the cells in about 60 ml of the culture medium selected for subsequent culture. Although the cells can be resuspended in cold medium and stored on ice, it is preferable to resuspend the washed pellets in warm medium, dilute to the desired density and proceed immediately with suspension experiments or plating into culture dishes as discussed in later Sections. (x) For specific applications, it may be appropriate to examine whether it is beneficial to further purify cells on Percoll (Kreamer et al., 1986) prior to maintenance in culture.

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10.6. Suspension culture of hepatocytes 10.6.1. Comparison of suspension and monolayer culture

For studies on hepatocyte gene expression, growth, mechanisms of toxicity or other phenomena where periods of study necessarily extend for more than 1-2 h, the most common approach for in vitro studies has been to use culture systems where hepatocytes attach to some (semi-)solid substratum and re-form at least some features of normal liver architecture. For applications where responses of interest occur within a few hours, hepatocyte suspensions may provide a convenient alternative. This section describes the requirements for maintaining hepatocytes in suspension for extended periods of from 4-24 h (cultures of non-adherent ‘spheroids’ are discussed in Section 10.7.1). For experiments with hepatocytes in suspension, acceptable levels of cell integrity can readily be preserved for G 6 h and, with additional measures, up to about 24 h, as discussed in the next Section. A major advantage of suspension culture experiments is simplicity. For incubation of suspensions up to about 5 h, it is desirable to use sterile incubation vessels and media but maintenance of sterility during cell isolation is not essential, so that much of the time-consuming preparation for longer-term culture experiments is avoided. The use of suspensions allows frequent sampling from a uniform population. A number of factors may influence the choice between suspension and monolayer cultures. Longer term monolayer cultures are frequently seen as having advantages in that cells ‘recover’ from the trauma of the isolation procedure. It has been suggested that recovery might include restoration of some hormone receptors (Morin et al., 1982; Goglia et al., 1985)and rates of protein synthesis (Ichihdra et al., 1982), restoration of normal intracellular ions and glutathione levels, repair of DNA damaged during cell isolation (Cesarone et al., 1984), removal of proteolytic enzymes, which may have persistent effects in fresh suspensions (Guzelian and Diegelmann, 1979; Capuzzi et al., 1979) and also removal (as non-attached cells) of any residual damaged

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hepatocytes. Some forms of damage or depletion in fresh hepatocytes can be minimized by refinements to the isolation procedure and, if desired, contamination of suspensions with damaged cells (or nonhepatocytes) can be minimized by centrifugation through Percoll. Cells in suspension inevitably lack various features of the organization of hepatocytes in the intact liver (see Chapter 5 ) . The suspended cells are spherical, entirely covered with microvilli and apparently lack specialized membrane regions such as bile canaliculi (e.g. Wanson et al., 1977; Maurice et al., 1988), although some elements of polarity are retained (Nickola and Frimmer, 1986). In addition, they lack the contacts with extracellular matrix or cell-cell contacts (with either other hepatocytes or with non-hepatocytes) which appear to be important for maintaining differentiated function in monolayer cultures (see Section I I . I). Although the l’ull effects of cell disaggregation per se are not clear it is likely that patterns of gene expression are altered, with increased levels of mRNAs for some cytoskeletal proteins (Clayton and Darnell, 1983; Clayton et al., 1985), transient expression of c-jiis and more sustained expression of c-my. protooncogenes (Etienne et al., 1988)and reduced expression of various genes associated with fullydifferentiated hepatocyte function (e.g. Clayton et al., 1985; Nakamurd and Ichihara, 1985). At least some of these changes may be reversed under appropriate monolayer culture conditions. In addition to the repair of damage, recovery of polarity and restoration of cell-cell interactions, a period of pre-incubation in monolayer culture may also allow better definition of experimental variables by minimizing effects of prior exposure to hormones or nutrients in vivo. On the other hand, for some applications, such as studies on metabolic control or the actions of xenobiotics, the fact that freshly isolated cell suspensions still reflect the metabolic capabilities of hepatocytes in vivo may be important. A major problem with monolayer experiments in many culture conditions has been rapid loss of differentiated functions including some hormone receptors, as discussed in Section I 1.1. The differing contributions of repair or recovery of some functions with loss of others in culture are illustrated

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in some direct comparisons of different hepatocyte functions in suspension and monolayer (e.g. Guzelian et al., 1977; Gurr and Potter, 1980; Kukongviriyapan and Stacey, 1989): such comparisons suggest that the choice of suspensions or monolayers for shorter-term studies should depend on the parameters under investigation. In summary, suspensions are most likely to be appropriate where phenomena under investigation require study periods of up to 5 or 6 h. Control of various liver enzymes with relatively short half-lives, which thus exhibit relatively rapid responses to changes in hormones or nutrients in the medium, have been studied (e.g. Ernest et al., 1977; Edwards and Elliott, 1974,1975; Canepa et al., 1985). Longer incubation periods are possible and have been used to study plasma protein synthesis (Jeejeebhoy et al., 1975, 1980; Chen and Feigelson, 1978; Moshage et al., 1985). Use of suspensions may be relevant when a metabolic profile closely resembling the intact liver is desirable. Examples include studies on mechanisms of xenobiotic toxicity in hepatocytes (Hsia et al., 1983; de Groot et al., 1988)where the freshlyisolated cells retain levels of the enzymes required to activate xenobiotics, comparable to levels found in vivo. For applications requiring longer experimental periods, or cells stably adapted to a defined in vitro environment, or where cell architecture or contacts are likely to be important, monolayer culture systems are preferable for in vitro studies. The two key requirements for extended suspension culture experiments are suitable mechanisms for keeping cells suspended and oxygenated with minimal damage to cells, and choice of a culture medium appropriate to the phenomena under investigation. 10.6.2. Maintenance of oxygenated suspensions of undamaged hepatocy tes

Systematic studies on conditions optimal for preservation of hepatocyte integrity in longer-term suspension culture have employed approaches involving either shaken flasks or spinner (magnetically-stirred)cultures.

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The first approach uses gentle shaking of suspensions under a carbogen atmosphere with a small volume of cells in siliconized-glass or polypropylene flasks (Crane and Miller, 1977; Dickson and Pogson, 1977; Ernest et al., 1977; Kreamer et al., 1986). In our laboratory, widebased polypropylene Erlenmeyer flasks were found to give better preservation of cells than plastic vials or siliconized glass (Scott, 1982). Vigorous cleaning of the flasks between experiments is important. Damaged cells stick to the plastic around the air-liquid interface and, unless completely removed, have adverse effects on cell integrity in subsequent incubations. Careful adjustment of shaking conditions is important for optimal cell preservation. Orbital shaking gives slightly better preservation than reciprocal shaking (Crane and Miller, 1977; Scott, 1982). Optimal shaking speed depends on the geometry of vessels and volume of cells but should be just sufficient to keep cells evenly suspended. Many laboratories have employed shaking speeds of @ 079oscillations/min, where the total volume of incubation vessels is 5-1 0 times the volume of the cell suspension. The use of simple vials or flasks allows a single cell isolation to be split between many parallel incubations, permitting concurrent comparisons of a variety of experimental treatments. Cells shaken in suspension have been maintained so that > 85% of hepatocytes exclude trypan blue for 5-6 h and, according to some reports, for even longer without purification steps (e.g. Chen and Feigelson, 1978). Nevertheless, complete removal of debris and damaged cells by centrifugation of isolated hepatocytes through Percoll (Dalet et al., 1982; Kreamer et al., 1986) apparently gives a marked improvement in long-term integrity of shaken suspension cultures. Some laboratories have used more complex apparatus for long-term preservation of suspensions in spinner culture, coupled with a system for sensing and adjusting 0,-concentrations in the medium, (Jeejeebhoy et al., 1980; de Groot et al., 1988). A disadvantage of these systems is that only a small number of different incubations is possible with each cell preparation. However, the method of Jeejeebhoy et al. (1980) retained undamaged cells for long periods, and high rates

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of protein synthesis, approximating those in vivo, were observed. In this system, cells progressively aggregated from 12-24 h forming groups of 10-50 cells in which re-formation of junctions and bile canaliculi was reported. These aggregates may resemble the ‘spheroids’ recently shown to form in non-adherent culture conditions (e.g. Koide et al., 1990). Cell densities used for suspension experiments have usually been in the range 1-3 x lo6 cells/ml although lower densities (4 x los cells/ml) have been used with good results (e.g. Kreamer et al., 1986). 10.6.3. Media ,for suspension culture

Although simple media have been used for extended incubations in suspension (e.g. Moore et al., 1985), the minimum requirements for maintenance of complex functions such as protein synthesis are a suitable energy source and substantial levels of amino acids. A culture medium such as Eagle’s MEM with relatively low amino acid content does not prevent progressive polysome disaggregation in suspension cultures although it sustains relatively constant levels of protein synthesis (Dickson and Pogson, 1977. 1980). In hepatocytes in simple media such as Krebs-Henseleit bicarbonate saline, protein degradation greatly exceeds synthesis (Section 6.6.8) and high amino acid levels, even in combination with an adequate energy substrate, are necessary to achieve approximate nitrogen balance with optimal rates of protein synthesis and substantial inhibition of degradation (Seglen et a]., 1980b; Schwarze et al., 1982). These workers also found that lactate or pyruvate (up to 20 m M ) are preferable to glucose and much more effective than fatty acids as an energy source in stimulating protein synthesis. Relatively high osmolality (around 400 mmol/kg compared with 307 mmol/kg in serum) and pH 7.6 were optimal for protein synthesis. Other medium components were not investigated in detail (Schwarze et al., 1982). It might be noted that various culture media available commercially such as Waymouth M B 75211 or Liebovitz L-15 apparently support

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relatively high and constant rates of protein synthesis and inducibility of enzymes as well as good preservation of cells in suspension (e.g. Jeejeebhoy et al., 1980; Kreamer et al., 1986). Most laboratories have used media with 1 mM Ca2+or more (but cf. Ernest et al., 1977) and have used bicarbonate-buffered media for suspension experiments. At higher cell densities ( >lohcells/ml), a fall in pH is observed with time when only bicarbonate buffering is used. In our experience it is important to maintain pH in the range 7.3-7.5. HEPES (10-25 mM) has frequently been added to boost buffering capacity, although care may be required. In our studies, HEPES concentrations above 15 mM impaired induction of 5-aminolevulinic acid synthase (Scott. 1982). Many laboratories have added serum (rat, foetal calf or heatinactivated horse serum) or BSA to enhance cell survival (e.g. Jeejeebhoy et al., 1975). In our hands, either lo'%,foetal calf serum or 0.2'%1albumin reduces the tendency of hepatocytes to aggregate at longer incubation times. However a protein supplement is not essential (e.g. Gurr and Potter, 1980; Kreamer et al., 1986). Choice of hormone supplements may depend on the parameters under investigation. Insulin stimulates protein synthesis and inhibits degradation, thus helping to preserve nitrogen balance (Jeejeebhoy et al., 1980; Schwarze et al., 1982), and may contribute to better cell preservation. Glucocorticoids have broad effects in retarding early changes in levels of hepatocyte proteins in monolayers (Colbert et al., 1985) and thus may help preserve normal function during suspension culture. Other hormones influence individual parameters without having documented effects on overall cell integrity in medium-term suspension experiments. 10.6.4. Methods for suspension culture of' hepatoc:,.tes The detailed protocol given below is designed to allow experimental studies extending to around 8 h or possibly longer, while utilizing relatively simple equipment and allowing multiple parallel incubations. It should be noted that for many purposes, satisfactory levels of cell integrity and function can be maintained for periods of 4-5 h with

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quite simple equipment and procedures. A laboratory experiment for undergraduates in our Department is concerned with hormonal control of tyrosine aminotransferase. Cells are prepared using either Protocol 2.2 or Protocol 10.1 without further purification and incubated as in Protocol 8.3.

Protocol 10.2 Prolonged incubation of hepatocyte suspensions (5-8 h ) (i) Polypropylene Erlenmeyer flasks are used as incubation vessels. To ensure these are very clean, brush out with a test tube brush and detergent, rinse and then fill the flasks with chromic acid for 30 min. Rinse thoroughly with distilled water and autoclave to sterilize. Rubber bungs of appropriate size can be sterilized in 70% ethanol, shaken dry and used as stoppers. (ii) Prepare the culture medium by supplementing Waymouth MB752/1 medium with 15 mM HEPES and 0.2% BSA, fraction V. Adjust to pH 7.5 at room temperature and sterilize by 0.2 pm membrane filtration. Store at 0 4 ° C . On the day of the experiment add antibiotics and insulin (from sterile, 200-fold concentrated stock solutions which are stored frozen) to give final concentrations of 60 pg/ml penicillin, 100 pg/ml streptomycin sulphate and 0.3 pM insulin. (Hormone additions will depend on the phenomena under investigation. Dexamethasone, 100 nM, may also be included for suspension culture.) (iii) For each experiment prepare two sterile 50-ml screw-cap plastic centrifuge tubes containing 21.6 ml Percoll mixed with 2.4 ml sterile ten-fold concentrated Hanks’ balanced salts (for 1 I of the concentrated Hanks’ solution, the composition is as follows: NaCI, 80 g; KCl, 4.0 g; MgS0,.7H20, 2.0 g; KH,PO,, 0.6 g; Na2HP0,.2H,O, 0.6 g and glucose, 10 g). Adjust to pH j.4 and keep at 0 4 ° C . (iv) Prepare hepatocytes as in Protocol 10.1. The volumes below are based on a yield of around 3 x lo8 hepatocytes per liver. Resuspend the pellet after the second 50 x g centrifugation in about 50

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ml culture medium at 0--4"C (the cell concentration should be in the range 5-8 x lo6 cells/ml). (v) Using a wide-bore pipette, place 25 ml of hepatocyte suspension into each of the centrifuge tubes containing Percoll/salts. Cap the centrifuge tubes and gently invert 4-5 times to thoroughly mix cells and Percoll. Centrifuge at 50 x g for 10 min at 0 4 ° C . After centrifugation there should be clear Percoll between the upper layer containing debris, damaged hepatocytes and non-parenchymal cells and the pellet containing intact hepatocytes. Carefully aspirate the supernatants and resuspend each of the pellets in 30 ml ice-cold culture medium. Wash the cells twice to remove Percoll, sedimenting the cells at 50 x g for 3 min. (vi) Resuspend the final pellets in a total of about 60 ml culture medium at 37°C using a wide-bore pipette. Transfer the suspension to a sterile 500-ml Erlenmeyer or De Long flask in which the cells can be readily swirled to preserve a uniform suspension. Dilute to about 150 ml, count cell numbers using a haemocytometer (Section 2.6.1) and dilute the suspension so that the final density is about 1.2 X lo6 cells/ml. (vii) Distribution of the cell suspension into incubation flasks is best done by two people, one pipetting and one gassing the flasks. Swirl the flask constantly to keep the cells evenly suspended and provide some oxygenation. The time between suspending cells in warm medium and establishing incubations should be kept to a minimum. Using a sterile wide-bore pipette, dispense 10-ml aliquots of the cell suspension into the 100-ml polypropylene flasks. Gas for 10 s with carbogen, stopper each flask and place in an appropriate clamp or rack in a 37°C water bath with orbital shaking at 80 cycles/min. (viii) Test compounds may be added to flasks at the time of dispensing the cell suspension or after a short period of pre-incubation. If flasks are opened for additions or sampling, they should be re-gassed with carbogen before continuing the incubation. After equilibration at 37°C with carbogen the pH should be 7.4-7.5. If the phenol red

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indicator shows significant fall in pH during incubations, adjust the pH by drop-wise addition of sterile 0.1 M NaOH.

10.7. Other non-adherent culture methods 10.7.1. Multicellular spheroids

An approach to hepatocyte primary culture which falls methodologically between suspension and monolayer culture involves seeding freshly isolated cells into a culture vessel, chosen so that hepatocytes cannot adhere to the surface, but instead re-associate with each other to form multicellular spheroids in suspension. Culture as spheroidal aggregates has been employed for a variety of other cell types: its application for liver cells was first described for neonatal hepatocytes by Landry et al. (1985) and more recently for adult hepatocytes by Koide et al. (1990) and for neonatal cells by Tong et al. (1990). During the 12 h after seeding, the cells form loose associations and subsequently reform junctional complexes to yield tightly-associated aggregates which increase in size from 50 pm in diameter after 1 day to 150-pm spheroids after 1-2 weeks. These have a surface layer of thin (probably mesenchymal) cells but contain mainly cuboidal hepatocytes within the aggregates, in 5-6 layers arranged in small islands, often about a central lumen. Extracellular matrix is present, associated in part with a small number of non-parenchymal cells. Some of the neonatal spheroids contain bile duct-like structures, lined with cells resembling bile duct epithelial cells (Landry et al., 1985). Tong et al. (1990) reported that the size of spheroids was greater in enriched media. It is presumably the re-formation of some of the features of liver architecture in vivo, including the laying down of extracellular matrix and extensive cell-cell interactions (possibly including heterologous interactions), that contribute to the long-term survival of hepatocytes in spheroids for periods of 2-8 weeks. After an initial decline over

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several days, spheroids from neonatal cells maintain stable levels of albumin secretion and tyrosine aminotransferase inducibility at 20-50'%, of initial values (Landry et al., 1985; Tong et al., 1990). Spheroids derived from adult hepatocytes, maintained in a hormonesupplemented medium, preserved high levels of albumin secretion but showed little evidence of PHIthymidine incorporation in response to epidermal growth factor (EGF), suggesting that spheroid culture favours differentiation but not growth. Transfer of spheroids into collagen-coated dishes results in hepatocytes migrating out of the spheroids to form a flattened monolayer where increased ["Hlthymidine incorporation could be observed (Koide et al., 1990). Spheroid cultures of hepatocytes have not yet been studied widely. The following description of method is based mainly on that of Koide et al. (1990). Hepatocytes were isolated by the two-step collagenase perfusion procedure of Seglen ( 1976) which eliminates most nonhepatocytes. It is not yet clear whether a low level of contamination with fibroblasts or other non-hepatocytes is essential for the long-term survival and function of spheroids. Inclusion of serum in media at any stage of isolation or plating prevented spheroid formation in any type of culture dish. Landry et al. (1985) and Tong et al. (1990) used dishes coated with poly(2-hydroxyethyl methacrylate) to block attachment, but for adult hepatocytes Koide et al. (1990) concluded that positivelycharged Primaria dishes (Falcon Labware, Becton Dickinson and CO.) provided the simplest culture surface for spheroid formation. Primaria or bacteriological dishes coated with albumin or a proteoglycan fraction could also be used. Adult hepatocytes (3 x 105 cells/l.5 ml per 35-mm dish) were plated in a variation of Reid's HDM medium (see Table 1 1.11, namely Williams' Medium E supplemented with 10 pg/ml insulin, 0.1 pM CuSO,, 3 nM H,SeO,, 50 pM ZnSO,, 50 ng/ml EGF, 50 pg/ml linoleic acid, 100 U/ml penicillin, 100 U/ml streptomycin and 1 pglml amphotericin B. Half of the medium was replaced by gentle pipetting after 4 h and then every 2 days. While Landry et al. (1985) achieved long-term survival of spheroids of neonatal cells in unsupplemented

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Williams’ Medium E, Tong et al. (1990) reported progressively better survival and preservation of albumin and transferrin secretion after sequential supplementation with the following groups of components: 1 pM dexamethasone, 1 pM glucagon and 0.1 pglml insulin; 50 ng/ml EGF; and a mixture of 20 mU/ml prolactin, 10 pU/ml somatotropin, 10 pg/ml linoleic acid, 0.1 pM H,SeO,, 0.1 pM CuSO, and 50 pM ZnSO,. It is thus possible that inclusion of dexamethasone, glucagon, prolactin and/or somatotropin in the medium of Koide et al. (1990), given above, may be useful. On the limited evidence so far, spheroid culture provides a relatively simple approach for achieving long-term survival of hepatocytes, probably with preservation of near-normal differentiation. It is yet to be shown that a wide range of differentiated functions is preserved or that patterns of transcription are nearer normal in spheroids than in simple monlayer systems. The spheroids may be useful for studying the role of cell-cell and cell-matrix interactions. Disadvantages are that experimental manipulations are less-easily performed than with monolayers, and that non-hepatocytes are present, which introduces some of the same complications as in monolayer co-cultures (Section 11.7). It remains to be established whether the presence of nonhepatocytes has an important influence on survival or function of the hepatocytes in spheroids. 10.7.2. Microencapsulation of hepatocytes

A variation of spheroid culture, which has been introduced for studies on hepatocyte transplantation, involves microencapsulation of hepatocytes in an alginate-polylysine membrane. The method for encapsulation, given in detail by Cai et al. (1989), involves suspension of about 3 x lo6 hepatocytes in 2 ml 0.90/0 NaCl containing 1.5”/0 sodium alginate and 0.045% Type I collagen. Spherical droplets of suspension are extruded from a syringe pump into 1.1% CaCI, solution, where gel spheres form. These undergo a series of washes with sequential exposure to polylysine, alginate and sodium citrate before

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the encapsulated cells are placed into culture vessels or used for transplantation. The spheres formed are about 0.7 mm in diameter with an alginate-polylysine membrane surrounding spherical hepatocytes embedded in a fibrillar matrix, apparently consisting mainly of alginate and collagen. Under the culture conditions used by Cai et al. (1989), encapsulated cells synthesized urea and secreted proteins, such as prothrombin, at substantial rates for 6 7 days and thereafter at declining rates for up to 5 weeks. The porosity of capsule membranes can be controlled so as to admit nutrients and hormones but exclude immunoglobulins and complement, giving effective screening of hepatocytes from a host immune system after transplantation.


CHAPTER 11

Monolayer culture of hepatocytes

11.1. Characteristics of adult primary monolayer cultures: an overview Any attempt to review comprehensively what is now a vast literature on hepatocyte primary culture is beyond the scope of this Chapter. However in order to provide some broader background to consideration of specific techniques for hepatocyte culture, this Section briefly outlines some general observations on the characteristics of hepatocytes in culture and the major variables which influence these characteristics. 11.1.1. Reviews of the literature

Most of the information on hepatocyte culture is for cells from adult rat liver. There is also substantial information on foetal or neonatal rat hepatocyte culture and more limited experience so far with hepatocytes from other species such as humans, mice and rabbits. There are a number of reviews on aspects of hepatocyte culture and its application which should be consulted for a more detailed coverage of the relevant literature. Early findings with rat hepatocyte primary culture were reviewed by Pitot and Sirica (1980) and Bissell and

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Guzelian (1980) and the use of liver in vitro systems in relation to pharmacology and toxicology have been discussed by Sirica and Pitot ( 1980) and Grisham (1979). Guguen-Guillouzo and Guillouzo ( 1983) summarized functional changes with time in adult, foetal and neonatal hepatocytes and discussed effects of soluble factors and specific cellcell interactions on maintenance of differentiated function. Ichihara et al. (1982) and Nakamura and Ichihara (1985) reviewed their studies on hormonal control of various liver-specific functions; and also discussed factors, including cell density, that regulate growth of hepatocytes in culture. Reid and Jefferson (1984) discussed the contributions of media and extracellular matrix components to culture of hepatocytes and other differentiated cells. The book edited by Guillouzo and Guguen-Guillouzo ( 1986) brings together a collection of excellent reviews on hepatocyte culture and this provides valuable information on methods as well as specific applications for cultured hepatocytes. In relation to choice of culture methods, the reviews on growth of hepatocytes (McGowan, 1986), on changes in differentiation in culture and their modulation by media and matrix components (Reid et al., 1986) and on the relevance of homotypic and heterotypic cell interactions (Guguen-Guillouzo, 1986) are particularly relevant. More recently Maher (1988) has provided an excellent short review of the varying approaches to maintaining differentiated function in hepatocyte culture. Finally, some recent original reports on the use of extracellular matrix as substratum for hepatocyte culture compare and discuss the significance of multiple culture variables (e.g. Bissell et al., 1987; Schuetz et al., 1988; Ben-Ze’ev et al., 1988; Guzelian et al., 1989). 11.1.2. Limitations of simple culture systems

The earliest hepatocyte culture systems involved plating isolated cells with added serum onto plastic followed by maintenance with media such as Williams’ Medium E, various Waymouth formulations or

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Leibovitz L-15. Subsequent modifications include coating dishes with Type I collagen which allows better cell attachment and prolonged retention of cells on dishes (e.g. Bissell et al., 1978). Omission of serum from media aids preservation of liver-specific functions (see Reid et al., 1986 for discussion), while addition of insulin and dexamethasone to serum-free media was found to improve attachment and the preservation of cells on dishes, modulate cytoskeletal organization and slow changes in the levels of various hepatocyte proteins (Laishes and Williams, 1976b; Marceau et al., 1986a; Colbert et al., 1985; see Guguen-Guillouzo and Guillouzo, 1983 and Reid et al., 1986 for references). Maintenance of cells on collagen-coated dishes in serumfree medium with insulin and dexamethasone provides a simple culture system, still widely used with minor variations, to examine a variety of questions about hepatocyte function over experimental periods of a few days. With such simple culture conditions the cells initially assume a flattened morphology on the substrate but detach from dishes after 1-2 weeks; and it is now clear that the cells undergo major changes with time in patterns of gene expression and differentiated function. The activity of some non-tissue-specific enzymes such as LDH OF glucose-6-phosphate dehydrogenase change only slowly but many liverspecific functions decline rapidly within 2 or 3 days. Changes include reduced synthesis of serum proteins such as albumin; progressive falls in the levels of gluconeogenic enzymes such as glucose-6-phosphatase; a decrease in various cytochromes P-450, NADPH cytochrome P-450 reductase, cytochrome b, and associated drug metabolizing activities; a decrease in expression of some isozymes characteristic of adult liver (e.g. pyruvate kinase L, aldolase B) and increased expression of isoenzymes typical of non-hepatocytes or foetal liver (pyruvate kinase M,, aldolase A) and of other proteins found in foetal liver such as afoetoprotein and y-glutamyl transpeptidase. At the same time there is increased expression of genes for some cytoskeletal proteins such as actin and tubulin and the extracellular protein, collagen Type I (reviewed in Sirica and Pitot, 1980; Guguen-Guillouzo and Guillouzo,

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1983; Reid et al., 1986; Schuetz et al., 1988). It is an over-simplification to consider all these changes as reflecting ‘de-differentiation’ or ‘foetalization’of hepatocytes. A variety of liver enzymes are hormonedependent in vivo and can be maintained at near normal levels under simple culture conditions if the appropriate hormones are added. (As shown for glucose-6-phosphatase by Spagnoli et al. (1983), for instance.) However there do appear to be some fundamental changes in patterns of gene expression in simple cultures. In particular, decline in liver specific functions reflects greatly diminished transcription of the relevant genes (e.g. Clayton and Darnell, 1983). In the early 1980s it was thus apparent that hepatocyte monolayers on simple substrata such as collagen and in relatively simple media had significant limitations in that normal patterns of gene expression underwent rapid changes in culture. Furthermore, early attempts to study growth found only modest levels of DNA synthesis and suggested cells were unable to enter mitosis (Pitot and Sirica, 1980). 11.1.3. Cell density and cell function in culture

The recognition in the early 1980s that cell density in culture has a significant role in modulating the function of adult hepatocytes on simple substrata has established two main streams of study. On the one hand, the finding that cells at low, but not high, density on collagen or other protein films exhibit at least limited growth in the presence of growth factors has provided the basis for evolving improved conditions for studies on growth (considered in outline in Section 1 1.1.7). On the other hand, the retention of cells on collagen and preservation of increased levels of some liver-specific enzymes is favoured by maintaining cells as tightly-packed confluent monolayers (e.g. Nakamura and Ichihara, 1985; Jauregui et al., 1986). The effects of high cell density in delaying the loss of some differentiated functions and minimizing the accumulation of mRNAs for cytoskeletal proteins may be accentuated if hepatocytes are maintained on a semi-

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mobile collagen gel rather than a protein film (Michalopoulos and Pitot, 1975; Ben-Ze’ev et al., 1988). Such effects may be partly due to more cuboidal, less flattened cell shape at high density, particularly on gels (Ben-Ze’ev et al., 1988). However, Nakamura and Ichihara (1985) concluded that the main factor in reciprocal effects of density on growth and differentiation was contact between hepatocytes. In the context of developing optimal culture systems for either differentiation or growth it would be desirable to understand which of the contacts between hepatocytes influence function and how the mechanisms involved relate to effects of the extracellular matrix and soluble factors discussed below. Unfortunately this area is poorly understood. In the intact liver there are extensive lateral contacts between hepatocytes (see Chapter 5). As seen in the electron microscope, these include gap junctions and tight junctions with associated intermediate junctions and desmosomes (e.g. Hughes and Stamatoglou, 1987). When disaggregated hepatocytes are plated in culture at high density, hepatocytes re-establish both close and looser associations with 3-6 neighbours, with reformation of desmosomes and tight junctions (Chapman et al., 1973; Wanson et al., 1977). In mouse cultures a high proportion of hepatocytes rapidly re-establish gap junctional communication but the extent of communication declines after 2 days (Ruch and Klaunig, 1988). In rat cultures, gap junctions may reform to some extent but disappear over 1-3 days under simple culture conditions (Fujita et al., 1987; Mesnil et al., 1987; Goulet et al., 1988). Gap junctions are partly restored by exposure of pure cultures to proteoglycans or glycosaminoglycans (Fujita et al., 1987) or in co-cultures (Mesnil et al., 1987; Goulet et al., 1988). It is thus possible that reformation of tight junctions or desmosomes may be involved in density effects in simple cultures and that better gap junction restoration may mediate some effects of proteoglycans or co-culture. At the molecular level, the ability of hepatocytes to re-associate in a cell-specific manner apparently involves membrane-associated proteins including the ‘cell adhesion molecules’ (CAMS).For hepatocytes, L-CAM may be important during embryonic development and

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CAMlO5, which appears later in gestation, may have a significant regulatory function in differentiated cells. The level of CAM105 falls by 70% during liver regeneration and is absent from the surface of hepatocellular carcinoma cells. As for cell contacts with extracellular matrix, cell-cell adhesion via such CAMS influences organization of the cell cytoskeleton (Obrink, 1986). Ichihara’s laboratory identified a ‘cell surface modulator’, a high molecular weight protein from the plasma membrane of hepatocytes and some other cells which could be shown to exert reciprocal effects on growth and liver specific proteins when added to cultured hepatocytes (Nakamura and Ichihara, 1985). The relationship of this to known cell adhesion molecules (Obrink, 1986) and the roles of any of these molecules in the junctions seen in the electron microscope are yet to be defined for hepatocytes. 11.1.4. Contribution of complex media to maintaining dfferentiated

function

In attempts to preserve a differentiated state in hepatocytes, use of high cell density at best delays but does not prevent the changes in gene expression seen with relatively simple substrata and media. In the last decade a variety of modifications to this basic culture system have been tested. One approach has involved evolution of more complex media with a variety of hormonal, nutrient and trace element supplements. For primate hepatocytes, the use of complex media alone seems to be adequate to achieve excellent preservation of function over long periods (e.g. Jacob et al., 1989). But for rat hepatocytes, complex media have gone only part-way towards supporting normal function. There is a very large and confusing literature on the significance of medium composition and it seems clear that individual components in media may exert positive effects on one differentiated function while having no effect or negative effects on others. For preserving different

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27 1

functions such as albumin synthesis, or maintaining ligandin or gap junction protein levels, different hormones or nutrients seem to be important (e.g. Jefferson et al., 1984; Spray et al., 1987). Furthermore it seems clear that interactions between nutrients or hormones can produce effects quite different from the sum of their individual actions (e.g. Dich et al., 1988). Despite these complexities some nutrient-enriched and hormonesupplemented media have been developed that have broad beneficial effects on long-term survival and function. Recent observations suggest that a careful balance of dexamethasone, insulin and glucagon concentrations can support survival for 2-3 weeks with near-normal preservation of some enzyme activities and partial preservation of others (Dich et al., 1988) and can preserve a high level of albumin transcription for up to 4 days (Lloyd et al., 1987). Another well-studied example is the hormone-defined medium of Reid and co-workers (Reid et al., 1986)which contains insulin, glucagon, EGF, prolactin, growth hormone, linoleic acid and copper, selenium and zinc salts. This medium preserves rates of synthesis of various serum proteins and levels of corresponding mRNAs at near normal levels for several days but these effects apparently result from stabilization of mRNAs, combined with low levels of transcription, rather than from restoring normal patterns of transcription (Reid et al., 1986). Whether other variations in physiological media can support normal transcription of liver-specific genes for extended periods remains to be established. One recent innovation, the use of a fully-defined modified Chee’s medium (with insulin and dexamethasone), appears to allow survival and preservation of some functions previously possible only with complex substrata or co-culture (Waxman et al., 1990). Further supplementation of hormone- and nutrient-enriched medium with the non-physiological agent, DMSO, has striking effects in preserving levels of some liver-specific proteins and corresponding mRNAs but this effect also seems to be due primarily to mRNA stabilization while transcription of the relevant genes remains abnormally low (Isom et al., 1987).

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11.1.5. Significance of extracellular matrix

Clayton et al. (1985) have shown that hepatocytes cultured in tissue slices, where cell contacts and tissue organization are largely retained, continue tissue-specific transcription at near-normal levels in simple culture media. This suggests that disruption of normal liver architecture makes a major contribution to altered patterns of transcription in hepatocyte cultures. It has been argued that restoring contacts with extracellular matrix components or other cell types, similar to contacts found in vivo, may be necessary to allow normal patterns of transcription (e.g. Reid et al., 1986). One approach has been to test individual components of extracellular matrix, or combinations of these in various forms, with pure hepatocyte cultures. Use of films of individual extracellular matrix proteins (collagens Type 111 or IV, fibronectin, laminin) as substrata does not greatly improve preservation of differentiated functions (Bissell et al., 1987; Sawada et al., 1987; Ben-Ze'ev et al., 1988).Type I collagen in hydrated gel form rather than as a dried film may enhance the stabilization of liver-specific mRNAs (Zaret et al., 1988) and apparently delays, but does not prevent, loss of differentiation (Michalopoulos and Pitot, 1975; Sirica et a]., 1979; Ben-Ze'ev et al., 1988). However when used to maintain hepatocytes in a sandwich configuration between two gel layers, Type I collagen greatly improves survival and preserves albumin secretion (Dunn et al., 1989), suggesting that geometry as well as the chemistry of matrix contacts with hepatocytes may be important. Interestingly, overlaying cells on collagen with laminin also markedly increases albumin transcription (Caron, 1990). Addition of individual proteoglycans or related glycosaminoglycans such as heparin (compounds which are components of liver extracellular matrix) to hormone- and nutrientenriched media has been shown to increase the levels of mRNA for albumin and some other liver-specific proteins, while lowering abnormally high mRNA levels for cytoskeletal proteins such as actin. These effects involve a change towards more normal patterns of transcription in culture (Fujita et al., 1987).

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While these observations suggest that contact with some individual matrix components can help restore normal patterns of gene expression in culture, the most impressive results have been obtained with complex mixtures of extracellular matrix components, notably with liver derived ‘biomatrix’ (Enat et al., 1984) or with ‘Matrigel’, a biomatrix preparation derived from the Engelbreth-Holm-Swarm (EHS) sarcoma. Used as a substratum for cell attachment, Matrigel supports survival of hepatocytes for about 3 weeks but probably no longer (Reid and Jefferson, 1984; Bissell et al., 1987). The cells do not flatten but retain a cuboidal shape and aggregate in clusters or columns that thicken with time (Bissell et al., 1987). Many liver-specific functions appear to be wholly or partly preserved although some are still lost progressively. For instance albumin secretion is maintained at 40-100% of in vivo levels (possibly depending on the medium used), reflecting partial preservation of normal mRNA levels (Bissell et al., 1987; Schuetz et al., 1988). One response on Matrigel which has been difficult to achieve in other variations of hepatocyte culture (but c.f. Waxman et al., 1990) is induction of phenobarbital-inducible cytochrome P-450 isozymes. This induction is mediated by phenobarbital-stimulated transcription to yield the relevant mRNAs in cells on Matrigel (Schuetz et al., 1988, 1990).The abnormally high expression of actin and tubulin genes and appearance of a-foetoprotein frequently observed in simple culture conditions are also suppressed on Matrigel (Schuetz et al., 1988; Ben-Ze’ev et al., 1988). While Matrigel or other biomatrices have been tested mainly as substrata and considered to act partly by preserving more cuboidal, non-flattened cell shape and perhaps partly via cell-matrix contacts in appropriate combinations (see discussion in Bissell et al., 1987; Schuetz et al., 1988; Maher, 1988; Ben-Ze’ev et al., 1988), a recent report suggests that addition of Matrigel to hepatocytes already flattened onto collagen-coated dishes restores substantial levels of albumin transcription without changing cell shape, implying that cell-matrix contacts (possible with specific geometry) may be more important than cell shape per se (Caron, 1990). There has been substantial progress towards understanding the in-

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lSOLATED HEPATOCYTES

teraction between cells and the extracellular matrix, although relatively few studies have focussed on hepatocytes. Contact of cells with matrix components such as fibronectin, laminin or collagens is believed to involve relatively specific cell surface receptors for individual matrix components (see Akiyama et al. (1990) for review). As is the case for points of cell-cell adhesion, the focal contacts between a cell and components of the extracellular matrix are points at which there is a structural interface between external ligands and the internal cytoskeleton of the cell (e.g. Geiger et al., 1987). It seems likely that these focal contacts are important sites of control since they are associated in nontransformed cells, with increased phosphotyrosine and various protein kinases, including protein kinase C (Burridge, 1986; Hyatt et al., 1990). It is likely that receptors for extracellular matrix components and receptors for soluble hormones or growth factors can each generate a variety of intracellular signals that interact to achieve overall control of growth and differentiation. Whatever the mechanism, cell interaction with Matrigel, probably in combination with defined, supplemented media, currently appears to provide the best approach for maintaining near-normal differentiation in pure hepatocyte cultures under well-defined conditions. Although yet to be fully chardcterized, it seems that in vivo patterns of transcription are at least partly restored by culture on Matrigel (Schuetz et al., 1988, 1990; Caron, 1990). 11.1.6. Heterologous cell interactions in co-culture

A second approach, used in attempts to duplicate in culture some of the complexities of cell and matrix contacts in the intact liver, is to co-culture hepatocytes with other cell types, in general related to those in liver in vivo. The best-studied co-culture system is that developed by Guguen-Guillouzo and her co-workers in which hepatocytes are co-cultured with an untransformed rat liver epithelial cell-line, possibly related to bile ductular epithelium (Guguen-Guillouzo, 1986). Coculture with other cell types, including liver endothelial cells, may also

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confer comparable advantages for preserving hepatocyte function (Goulet et al., 1988). In either co-culture system with glucocorticoidcontaining media, hepatocytes are preserved with relatively constant numbers and morphology from day 2 for 1-2 months. Albumin secretion, a number of other liver-specificfunctions and gap junctional communication between hepatocytes are maintained at relatively high levels whereas a-foetoprotein does not appear (Guguen-Guillouzo, 1986; Goulet et al., 1988). After an initial decline as cell-cell and cell-matrix contacts become established, mRNA levels for albumin and some other liver proteins stabilize at about 50% of in vivo values, reflecting transcription rates ranging from 20 to 80% of in vivo levels (Fraslin et al., 1985). Cell-cell contact between the two populations promotes the laying down of reticulin fibres containing fibronectin, Type I collagen and laminin, between the populations and over the hepatocytes. It is suggested that a cooperative interaction of the two cell types in production of the extracellular matrix, and possibly membrane contacts between the cell types, may be major elements in the effect of co-culture on differentiation. However, the mechanisms involved are not fully understood (Guguen-Guillouzo, 1986; Mesnil et al., 1987; Goulet et al., 1988). Monolayer co-culture systems may share some common features with non-adherent spheroid cultures (described in Section 10.7) where hepatocytes are associated with some other cell types and there is extensive extracellular matrix production. Co-culture is clearly one of the more successful approaches to maintaining differentiated hepatocytes although, as for most approaches, there are activities such as induction of phenobarbital-inducible cytochrome P-450isozymes that are not sustained (Schuetz et al., 1988).Also there are additional complexities in experimental manipulation and interpretation introduced by the presence of a second cell type. 11.1.7. Growth of hepatocytes in culture

The second major area of study with hepatocyte monolayers involves efforts to achieve significant levels of DNA synthesis and mitosis.

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Maintenance of cells at lower densities on films of collagen or other proteins remains the basic method for studying growth. Comparison of DNA synthesis in cultures maintained on different protein films suggests Type IV collagen may be the best substratum for achieving growth (Reid et al., 1986). Slightly lower rates are observed on fibronectin or Type I collagen with a further reduction in D N A synthesis in cells on laminin (Sawada et al., 1987). Spreading of cells on the substratum appears to be necessary for substantial levels of replication (Sawada et al., 1987; Ben-Ze’ev et al., 1988). On Matrigel (BenZe’ev et al., 1988), in co-culture (Fraslin et al., 1985) or in spheroids (Koide et al., 1990) very few cells are active in D N A synthesis and in general it appears that substrata (or co-culture interactions) that favour differentiation suppress growth. Possible exceptions are the reports that some liver biomatrix preparations support growth (Enat et al., 1984) and that insoluble protein(s) produced by a feeder layer of liver non-parenchymal cells stimulate hepatocyte D N A synthesis (Shimaoka et al., 1987). The principal advances towards achieving hepatocyte growth have come from evolution of media that meet basal requirements (e.g. nutrients and trace metals) for supporting DNA synthesis and from identifying growth-stimulatory factors for hepatocytes (reviewed by McGowan, 1986; Michalopoulos, 1990). A number of factors including EGF, a-transforming growth factor (a-TGF), serum-derived hepatopoietins A and B, insulin, a,-adrenergic agonists and oestrogens stimulate D N A synthesis alone or in combination with EGF. There are other agents including glucocorticoids, glucagon and some other hormones, which have been included in growth promoting media, but for which there is less consistent or clear evidence of stimulatory effects. For hepatocytes at lower densities on substrata that allow cell spreading, various combinations of the growth factors above can stimulate at least limited growth. A high proportion (about 80%) of hepatocytes will undergo an initial round of DNA replication, many undergo mitosis at least once and a few cells may undergo further

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round(s) of DNA synthesis (e.g. Michalopoulos et al., 1982; Nakamura and Ichihara, 1985; McGowan, 1986; Sawada et al., 1986; Yusof and Edwards, 1990). The extent of mitosis and cytokinesis has been difficult to quantitate precisely in adult cultures, partly because of progressive detachment of cells from monolayers. However, with few exceptions (e.g. Enat et al., 1984),measurements of total cell or nucleus numbers during 6 5 days in culture have shown increases of not more than 50-70'%0 over numbers on day 1 in culture (e.g. Hasegawa et al., 1982; Sawada et al., 1986; Sand and Christofferson, 1988; Yusof and Edwards, 1990). Taken together with studies on changes in cell ploidy in growth factor-stimulated cultures (Tomita et al., 1981),these findings suggest that after DNA replication, many but not all cells in culture may undergo mitosis and that only some of the cells undergo cytokinesis. This situation is not necessarily the result of deficient culture conditions, since in vivo in adult liver, a growth stimulus to mononuclear or binuclear diploid or tetraploid hepatocytes may have a variety of outcomes, involving different combinations of replication, mitosis and cytokinesis (see Styles et al., 1987, for discussion). Little is currently known about factors which control the extent of mitosis or cytokinesis in adult hepatocytes in vivo or in culture. There is debate on whether Ca*+concentrations in culture media influence the extent of mitosis (see Section 11.4.4). Thus, under the conditions tested so far it seems that relatively few cells in primary culture undergo multiple rounds of DNA synthesis and mitosis (e.g. Eckl et al., 1987) and that DNA synthesis stops after 3 4 days. It is interesting that temporary exposure of these quiescent cells to DMSO may restore competence for further DNA replication, suggesting that the cycling of cells between a period of growth (without DMSO) and a more differentiated state (with DMSO) may allow multiple rounds of replication in culture (Chan et al., 1989). As discussed in Section 1 1.10 it has been possible to achieve more extensive growth of foetal or neonatal hepatocytes than of cells from adult rats.

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11.1.8. Summary

A variety of approaches involving media, substrata, different culture configurations and heterologous cell interactions are currently being employed either to preserve differentiated functions or to achieve growth. In efforts to preserve differentiation there may be various ways of achieving related ends, using different combinations of substrata, media (with or without agents such as DMSO) and cell interactions and it is not clear that any approach so far is the method of choice for studying a phenomenon requiring near-normal hepatocyte differentiation. Even for studies on a particular parameter, for example expression of mRNAs for different glutathione-S-transferase subunits, direct comparison of different culture conditions has not necessarily led to simple conclusions about the ‘best’ culture method (Vandenberghe et al., 1990). Thus in the Sections below, some specific protocols are given but methods relating to a wider variety of useful approaches to monolayer culture of hepatocytes are also discussed.

11.2. Attachment of hepatocytes to substrata for monolayer culture 11.2.1. Significance of cell attachment to substrata It is frequently suggested that normal mesenchymal or epithelial cells in vitro must attach to a substrate to survive for extended periods and most approaches to long term culture of hepatocytes utilize a solid or semi-solid support for attachment. For fibroblasts there is dramatic inhibition of complex processes such as DNA, RNA and protein synthesis when cells are detached from a substrate and for most normal cells, growth requires anchorage to a substrate. The relative importance of adhesion via receptors to particular extracellular matrix components and of changes in cell shape and cytoskeletal organization that follow adhesion may vary for different cell types (see Burridge,

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1986 for references and discussion). For hepatocytes, it has been reported that cells in suspension can synthesize proteins at rates comparable with perfused liver (Jeejeebhoy et al., 1980) although in other studies, attachment increased protein synthesis (Agius et al., 1985). Cell adherence to different substrata seems to have a major influence on preservation of liver-specific functions (see Section 11.1.5) and probably on growth, where the ability of cells to spread on a substratum may be important (e.g. Sawada et al., 1986), in line with observations for other cell types (e.g. Folkman and Moscona, 1978). In the intact liver, hepatocytes have contacts to varying extents with a variety of extracellular matrix components including collagen Types I, I11 and IV, fibronectin, laminin and heparan sulphate proteoglycan (Martinez-Hernandez, 1984; Bissell et al., 1987, 1990). Either complex extracts of extracellular matrix or (partially) purified preparations of extractable components have been employed as an alternative to tissue culture plastic as substrates for hepatocyte culture. This Section reviews findings on attachment of liver cells to those commonly used substrata that allow relatively specific adherence and extended maintenance of hepatocytes. 11.2.2. Characteristics of attachment to various substrata Hepatocytes attach poorly to tissue culture plastic and although the presence of serum during plating can provide fibronectin to mediate attachment, longer term maintenance of cultures is more readily achieved by coating dishes with individual matrix proteins or mixtures of matrix components. Typically, dishes are coated with proteins by air-drying dilute solutions on the dishes. The resulting adsorbed protein may, however, be slowly washed off the plastic (Macklis et al., 1985)and although this is not yet thoroughly explored, it is possible that long term maintenance of hepatocytes on protein films will require larger amounts of protein and/or some form of covalent attachment of proteins to plastic (Macklis et al., 1985)or to glass (Aplin and Hughes, 1981). Hepatocytes will attach to dishes coated with

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collagen Types I or IV or with fibronectin or laminin. Substantial cell spreading is observed on all these substrates and is greatest on Type IV collagen and least on laminin (Bissell et al., 1986; Sawada et al., 1987). Consistent with studies on receptors in other cell types (Akiyama et al., 1990),there is some evidence for distinct receptors on hepatocytes for each type of protein (e.g. Bissell et al., 1986; Johansson et al., 1987). A hepatocyte surface protein which can bind Type IV collagen, laminin or heparan sulphate proteoglycan has also been reported (Clement and Yamada, 1990).Cell spreading has been shown to require a higher concentration of matrix proteins per unit surface area than mere attachment (Bissell et al., 1986). Hepatocytes also attach readily to hydrated gels formed from Type I collagen or from complex mixtures of basement membrane components such as EHS-tumour extract (Matrigel). Recent experience in many laboratories suggests that the majority of intact hepatocytes (between 75% and 90%) will attach to any of the substrata above, with little difference in attachment to different proteins (Jauregui et al., 1986; Sawada et al., 1987) except when very low amounts of protein per dish are used (Bissell et al., 1986). 11.2.3. Conditions for hepatocyte attachment It is clear that attachment to protein films or gels can occur in the absence of serum or hormones (e.g. Bissell et al., 1986). Serum, although often included in media during cell attachment, probably does not enhance attachment to protein substrata, at least where hormones such as insulin and dexamethasone are present (e.g. Jauregui et al., 1986), although serum is clearly required for attachment of normal hepatocytes to plastic (e.g. Gjessing and Seglen, 1980).There have been several reports that insulin and/or dexamethasone enhance attachment (see Bissell and Guzelian, 1980; Guguen-Guillouzo and Guillouzo, 1983, for references). In our laboratory, insulin increases attachment efficiency to Type I collagen from 50-60% to above 80% and we routinely include 0.3 pM insulin during plating. Effects of insulin or

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dexamethasone on attachment do not appear to have been investigated in any detail. A variety of culture media with or without antibiotics support similar attachment rates (Williams et al., 1977; Jauregui et al., 1986). To maximize attachment efficiency, Williams et al. (1977) found that larger innoculation volumes were preferable: for instance, for a 50-mm diameter dish (surface area, approximately 20 cm2),a plating volume of 5 ml would be expected to give substantially higher attachment than 2.5 ml containing the same number of cells. As the number of cells plated per cm2culture surface increases, the number of cells that will x lo5 attach to the substratum increases up to about 1.3-1.5 cells/cm2 on collagen-coated dishes. At this density the cells should form a confluent monolayer. Further increasing the plating density to about 2 x lo5 cells/cm2 may yield more densely-packed monolayers although some cells, initially piled up in a double layer, detach after 1-2 days in culture (e.g. Jauregui et al., 1986). Since cells remain rounded on Matrigel it is possible to plate slightly higher cell numbers (about 1.7 x lo5 cells/cm2) yielding cultures containing mainly multicellular aggregates after 1-2 days (e.g. Bissell et al., 1987; Schuetz et al., 1988). Efficiency of attachment declines at higher cell densities and plating excess cells ( > 2 x l o 5 cells/cm2 on collagencoated dishes) leads to poor and patchy attachment. Hepatocytes attach within 30-60 min to the above substrata, at least at lower cell densities, and within 60 min to plastic in the presence of serum (e.g. Rubin et al., 1981; Gjessing and Seglen, 1980; Jauregui et al., 1986).After a suitable period for attachment (up to 4 h but ideally, 1-2 h), the medium with unattached cells should be removed and fresh medium added. Laishes and Williams (1976a) observed that intact hepatocytes attached more rapidly than damaged cells and suggested an attachment period of 1 h to minimize contamination of monolayers with non-viable cells. The following Sections list some additional information on the preparation, use and functional effects of commonly used substrates for hepatocyte culture.

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11 -3. Preparation, use and functional effects of specific substrata 11.3.1. Plastic

Hepatocytes will attach weakly by adsorption to glass or polystyrene. Adsorption, which is rapid and relatively independent of temperature, can occur in the absence of divalent cations (although it is stimulated by Ca2+)and can be blocked by pre-treatment with albumin to saturate adsorption sites (Gjessing and Seglen, 1980). In serum-free media, firm attachment to plastic does not occur within 1 h (Williams et al., 1977) and there is little spreading (flattening) of cells that are merely adsorbed. By 4 h after plating, hepatocytes may synthesize sufficient fibronectin to mediate more specific attachment with flattening of cells and subsequent survival for at least 24 h (Blaauboer and Paine, 1979). Such delayed attachment to plastic is probably stimulated by insulin which has frequently been included in attachment media (e.g. Koide et al., 1990). Hepatocytes will initially form monolayers on negatively-charged tissue culture plastic, on positively-charged plastic (e.g. Primaria or polylysine-coated dishes) or to some extent on hydrophobic bacterial petri dishes. Human foetal hepatocytes (plated with serum) have been maintained in Primaria dishes (Sells et al., 1985) but adult rat cells cultured on bacterial or positivelycharged plastic detach after 1-2 days and may form spheroids of intact cells (Koide et al., 1990). 11.3.2. Collagen films or gels

Hepatocytes attach well to all types of collagen (Rubin et al., 1981) although they have particularly high affinity for Type IV collagen (Bissell et al., 1986). For some cell types, attachment to collagen is mediated by fibronectin (Kleinman et al., 1979) but this is not thecase for hepatocytes (e.g. Bissell et al., 1986). Attachment to collagen is temperature-dependent and maximal in the presence of both CaZ+

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and Mgz+ ions (Gjessing and Seglen, 1980). Only relatively small amounts of collagens are required on dishes for attachment and cell spreading. Where solutions are air dried on dishes, addition of between 0.5 and 15 pg collagen/cm2 gives a film sufficient to support maximal cell attachment (e.g. Rubin et al., 1981; Sawada et al., 1987; Jauregui et al., 1986). Amounts of Type IV collagen as low as 0.05 pgkm2 may be adequate for maximal attachment and cell spreading (Bissell et al., 1986). Bacterial petri dishes have been used for collagen-coating(e.g. Rubin et al., 1981) but in our hands, monolayers detached from collagencoated bacterial plastic after 1-2 days. To establish stable cultures, tissue culture plastic was required. A variety of methods have been used to coat dishes with collagen. The simplest procedures involve air drying a dilute collagen solution containing about 3-10 pg collagen/cm2 onto dishes, or allowing collagen to adsorb to the plastic over a few hours from a solution of about 1 mg collagen/ml and then washing (Kleinman et al., 1987). Such methods give collagen films which promote maximal hepatocyte attachment comparable with films formed by salt-precipitation of collagen (e.g. Jauregui et al., 1986). For studies requiring longer term culture on collagen, there may be merit in cross-linking collagen to dishes (Macklis et al., 1985). Type I collagen in gel form provides a more flexible substrate on which hepatocytes can retain a cuboidal shape, less flattened than on protein films (Michalopoulos and Pitot, 1975). If detached from dishes after cell attachment, collagen gels shrink, allowing close contact between hepatocytes. Gels may also stabilize matrix components such as heparan sulphate proteoglycan released by the cultured cells (e.g. David and Bernfield, 1981). Findings for fibroblasts suggest that use of attached or floating collagen gels has different effects on growth because of differences in collagen organization (Nakagawa et al., 1989) but this has not been explored for hepatocytes. Type I collagen, dissolved at 3-5 mg/ml in dilute acetic acid forms a gel when brought to neutral pH at 2 6 3 7 ° C . This can be achieved with ammonia vapour or by adding a concentrated phosphatehalt solution plus sufficient

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NaOH to achieve a neutral pH (Elsdale and Bard, 1972; Michalopoulos and Pitot, 1975; Kleinman et al., 1987). Type I collagen has been the most widely used substrate for tissue culture, partly because it can be prepared easily and cheaply. The protocols below describe its preparation and use. Type IV collagen can be obtained commercially (e.g. from Sigma) or prepared from EHS tumour grown in P-aminopropionitrile-treatedmice (Kleinman et al., 1982). Type IV collagen can be used as a protein film on dishes but it will not form stable gels.

Protocol 1 1.1 Preparation of Type I collagen from rat tail tendons (i) Rat tails can be collected and stored at -20°C. Four or five tails will yield ample material for many months of collagen coating. Larger preparations are desirable where the collagen is to be used to form gels. (ii) Thaw the tails and wipe with 70% ethanol. Use gloves and instruments swabbed with 70% ethanol to minimize contamination of tendons. (iii) Cut the skin along the length of the tail from base to tip with sharp-nosed scissors. Using forceps, fold the skin back on either side of the cut at the base of the tail. Grip the inner part of the tail with toothed forceps and the skin with a second pair of forceps and strip off the skin by pulling it towards the tip. (iv) Hold the tail at the base and use pliers to grasp two or three joints at the tip. Twist pliers back and forth to crack the adjacent joint. Then pull tendons out from the tail. Place the joints and tendons on paper towel to dry (e.g. in a laminar flow cabinet). Repeat the cracking and drawing procedure until about 3 cm remains at the base of the tail: this can be discarded. (v) Repeat these procedures for several tails. (vi) Allow the tendons to dry for 1-3 h in a laminar flow cabinet. (vii) Cut the tendons away from joints and any blood vessels and cut into 3 4 - c m lengths. Separate as far as possible into fibres by rubbing the tendons between gloved fingers.

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(viii) Place the separated fibres into large pre-weighed petri dishes and place the dishes (tops off) under UV light for 18-20 h to sterilize the fibres (fibres placed 25 cm from a 20 W UV tube have invariably been sterile in our experience). From this point fibres should be handled with full sterile precautions. (ix) Weigh the dishes and determine the weight of collagen fibres. (x) With sterile forceps transfer fibres to a sterile flask containing a magnetic stirring bar. Add 250 ml sterile 0.1% acetic acid/g fibres and stir overnight at room temperature. (xi) There is usually a small amount of undissolved material which can be removed by centrifugation, by filtration through gauze or most simply, by allowing undissolved fibres to settle and decanting the collagen solution. (xii) The stock solution of collagen can be stored at M " C for several months.

Protocol 11.2 Collagen coating of dishes (i) For collagen coating of dishes, dilute the concentrated collagen solution with sterile 0.1% acetic acid. The extent of dilution depends on the method chosen for coating. (ii) Dispense diluted collagen (e.g. a 1 :15 dilution) or commercial Type I collagen at about 200 pg/ml into culture dishes (tissue culture plastic) in a volume that will cover the surface after swirling or rocking the dishes (e.g. 40 pl/cm2 - about 0.4 ml in a 35-mm dish or 0.8 ml in a 60-mm dish). Air-dry dishes at room temperature for 2-8 h or overnight at 40°C. This procedure gives a coating of about +8 pg collagen/cm'. Alternatively a rapid procedure for coating a large group of dishes with collagen is to tip a 1:7 dilution of the concentrated collagen solution from dish to dish, A little care with fingers and avoidance of drips is required to preserve sterility. For 60-mm dishes, for instance, tip about 4 ml diluted collagen from one dish to the next until the re-

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maining volume does not easily cover the surface and is discarded. A thin film of solution remains in each dish (about 0.2 ml) and after air-drying, leaves a collagen coating of 3-4 &cm2 which is adequate to support maximal attachment of hepatocytes. With such thin films, washing is not really necessary before plating cells onto the dishes. Coated dishes can be stored at room temperature for a few weeks.

Protocol 11.3 Formation of collagen gels (i) Undiluted collagen from Protocol 1 1.1 is used. The procedure is based on that of Kleinman et al. (1987) and is appropriate for 60-mm diameter dishes. The volumes can be scaled up or down in proportion to the surface area to be covered. In general gels should be prepared within the 24-h period prior to plating out cells. (ii) Pipette 1.8 ml concentrated collagen into 60-mm tissue culture dishes. Add 0.2 ml of solution consisting of 1 part 1 M NaOH and 5 parts phosphatehalt (0.2 M sodium phosphate buffer (pH 7.4) containing 1.5 M NaCl and 0.05 mg/ml phenol red). Mix thoroughly with the collagen. The phenol red in the resulting mixture should have a salmon-pink colour indicating a pH around 7.0-7.4. With different collagen preparations, it may be necessary to adjust the amount of NaOH used. (iii) A firm gel should form within a minute or two at room temperature but in general, dishes should be left undisturbed at 37°C for 30 min to ensure the formation of stable gels. (iv) Wash the gels with 2 ml culture medium. The wash can be left overnight and removed prior to plating cells onto the gel. Alternatively, if the gels are required for immediate use, washing for a few minutes is sufficient. (v)For experiments with collagen gels, cells are plated into dishes with gels attached to the bottom of the dish. After attachment of cells and a medium change, detach the gels by loosening them from around

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the edge of the dish with a sterile pasteur pipette (flamed to seal the tip). Using gentle agitation, free gels from the plastic so that they float in the medium. 11.3.3. Matrix glycoproteins: fibronectin, laminin

Hepatocyte attachment to dishes coated with fibronectin or laminin probably involves cell surface receptors that are distinct from each other and from receptors involved in binding collagens (Johansson, 1985; Bissell et al., 1986; Johansson et al., 1987; Forsberg et al., 1990). Compared with Type IV collagen, larger amounts of fibronectin or laminin are required for hepatocyte attachment and spreading (Bissell et al., 1986). As for collagens, dishes can be coated by allowing the glycoproteins to adsorb to tissue culture plastic from solution (about 40 pg/ml - Deschenes et al., 1980) or by air-drying dilute solutions onto dishes. For either protein, amounts in the range 0.1-10 pg/cm* have been air dried onto dishes and have supported maximal attachment and spreading (Johansson et al., 1981; Bissell et al., 1986; Sawada et al., 1987). Alternative approaches for achieving fibronectin-mediated attachment are to allow fibronectin from serum to adsorb to culture dishes (Gjessing and Seglen, 1980) or to plate hepatocytes into culture dishes with 10-15% foetal calf serum (e.g. Bonney and Maley, 1975). Cell attachment to fibronectin is temperature-dependent and strongly stimulated by Ca2+or Mgz+or both at around 1-2 mM (Gjessing and Seglen, 1980). Attachment occurs more readily to adsorbed fibronectin or to fibronectin associated with other matrix components (collagen, heparan sulphate) than to soluble fibronectin, suggesting that fibronectin conformation is influenced by association with other components (Johansson and Hook, 1984). Both fibronectin and laminin are available commercially. Fibronectin can be prepared from plasma using gelatin sepharose affinity chromatography (Engvall and Ruoslahti, 1977). Laminin can be prepared from EHS tumour (Kleinman et al., 1982).

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11.3.4. ‘Matrigel’ and other extracellular matrix preparations

Rojkind et al. (1980) pioneered the culture of hepatocytes on a ‘biomatrix’ preparation from liver, designed to approximate the extracellular matrix with which hepatocytes are associated in vivo. Their ‘biomatrix’contained mainly collagens (Types I, 111 and IV),glycoproteins (including fibronectin) and probably glycosaminoglycans. This material, or variations on it, supported 85% attachment of hepatocytes and long term survival with a relatively flattened morphology: some combinations of substratum preparation and medium were reported to support growth (Enat et al., 1984).Partly because of the difficulties of preparing useful amounts of this material in a reasonably homogeneous form for coating culture dishes, ‘biomatrix’ has not been widely used. A more convenient but substantially different source of extracellular matrix material evolved from the finding that the EHS sarcoma, a transplantable mouse tumour, produces large amounts of basement membrane material, which is less extensively cross-linked and more soluble than biomatrix from normal intact tissue (Orkin et al., 1977). EHS tumour extracts (sold commercially by ICN Biomedicals as ‘Matrigel’) are prepared by urea extraction of salt-washed tumour tissue. The tumour contains laminin (15 mg/g wet weight), Type IV collagen (8mg/g) and heparan sulphate proteoglycan (1 mg/g) as major components. Extracts of tumours contain predominantly laminin with smaller amounts of Type IV collagen, heparan sulphate proteoglycan, entactin and some minor components. Relatively small amounts of Type IV collagen are extracted because it is cross-linked by disulphide bonds and by lysine-derived cross-links (Kleinman et al., 1986). A dialysed urea extract alone will form a gel at 37°C and this preparation has been used directly for culturing hepatocytes (Schuetz et al., 1988; Ben-Ze’ev et al., 1988). A variation of this procedure involves extraction of tumours from mice in which collagen cross-linking has been reduced by inclusion of 0.1% 0aminopropionitrile in their drinking water (added freshly each day) from about 2 weeks after tumour innoculation for 2-3 weeks until

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the time ofexcising tumours (Bissell et al., 1987). Reducing the extent of collagen cross-linking may allow better extraction of Type 1V collagen. When the dialysed urea extracts described above are brought to 37"C, only part of the total content of laminin and other proteins participates in gel formation. Some workers have supplemented the extract with Type IV collagen (Kramer et al., 1986) to improve gel formation: addition of extra Type IV collagen and/or heparan sulphate proteoglycan results in increased incorporation of laminin and entactin into the gel structure (Kleinman et al., 1986). At present it is not clear which preparation is either optimal for hepatocyte culture or makes the most efficient use of tumour material. There is considerable variation in the amount of Matrigel per culture dish used by different laboratories. The usual dialysed urea extracts (see Protocol below) contain about 10 mg protein/ml. They can be diluted two-fold and retain gel-forming ability. ICN Biomedicals recommend the use of 0.5 ml of their Matrigel preparation per 35-mm dish. Bissell et al. (1987) and Schuetz et al. (1988) used comparable amounts of undiluted extract: these volumes of Matrigel give gels about 1 mm thick. Ben-Ze'ev et al. (1988) used smaller amounts of material (1.2 mg protein/35 mm dish) and Guzelian's laboratory described the successful use of as little as 100 p1 undiluted extract ( 1 mg) spread on 60-mm dishes (Schuetz et a]., 1988; 1990). Such small amounts of Matrigel may remain uniformly attached to some brands of dish (e.g. Lux) better than others. For longer-term experiments ( > I week), larger amounts of Matrigel may be preferable to preserve the spherical (rather than flattened) shape, characteristic of cells on this lamininrich substratum.

Protocol 1 1.4 Preparation and use of Matrigel The procedure described is the simplest that gives gels for hepatocyte culture and is based on procedures described by Kleinman et al. ( 1986), Kramer et al. (1986) and Schuetz et al. (1988). (i) The EHS tumour is often passaged in female C57BL mice

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although other strains may be used. For transplantation, use tumours which are still growing and free of necrotic tissue (about one turnour to inject 10 mice). Mince and wash the tumour tissue with sterile phosphate-buffered saline (PBS - see Section 1 1.12 for composition). Pass the tissue several times through a 20-ml syringe (without needle) and then through a 15G needle. Wash the tissue twice with PBS, centrifuging for about 30 s at 200 x g. Resuspend the tissue in PBS to make a thick slurry and use this to inject anaesthetized mice (8-12 weeks old) via a 15G needle. Inject 0.2-0.4 ml of the tumour tissue slurry subcutaneously on the back of the mice. The tumour should be palpable after 1 week and, depending on the amount injected, reach 3-5 g in about 4 weeks. (ii) Depending on the condition of the mice, harvest tumours when they reach 3-5 g. Kill the mice and swab fur with 70% ethanol. Using sterile instruments to minimize contamination, cut the skin alongside the tumour, dissect out the tumour and place it in a sterile petri dish. Cut the membrane surrounding the tumour, scrape out the bulk of the tumour tissue and transfer it to a sterile container on ice. When all tumours have been collected, freeze the tissue by dropping small 'balls' into liquid nitrogen, Collect the balls and store at -70°C until beginning the extraction. (iii) To store tissue for future transplantation, wash tissue from the slurry prepared in (i) above with sterile culture medium, suspend in culture medium containing 50% foetal calf serum and 10% DMSO and freeze for storage in liquid nitrogen using the same procedure as for cryo-preservation of cell-lines. (iv) All extraction procedures are conducted at 0-4"C. During and after the urea extraction steps it is essential to keep the extract cool since gel formation will occur at room temperature or above. (v) The following steps are given for 100 g of tumour. Thaw the tumour tissue. Homogenize in 300 m13.4 M NaCl, 0.05 M Tris buffer (PH 7.4) containing 10 mM EDTA and 2 mM N-ethylmaleimide. Centrifuge at 10,000 x g for 15 min and discard the supernatant.

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(vi) Wash the pellet twice more by homogenizing in 250 ml of the same buffer and centrifuging as above. (vii) Homogenize the washed pellet in 100 ml freshly-prepared buffered urea solution (0.05 M Tris-HC1, pH 7.4, containing 2.0 M urea and 0.15 M NaCl). Stir at 0--4"C overnight. Centrifuge at 10,000 x g for 30 min. Keep the supernatant at M " C . (viii) Resuspend the pellet in a further 80 ml of buffered urea solution. Stir for approximately 1 h at W " C , centrifuge at 10,000 x g for 30 min, and save the supernatant. (ix) Combine the supernatants from (vii) and (viii), transfer into boiled dialysis tubing and dialyse against 2 1 Trishaline (0.05 M Tris-HCI, pH 7.4, with 0.15 M NaCl) to which 10 ml chloroform has been added. The chloroform, added to sterilize the extract, is removed by subsequent dialysis steps. Dialyse 2-3 h. (x) Dialyse for a total of 24 h against two further 2-1 vols. of fresh Trishaline. (xi) Finally dialyse for 4 h against 1 1 of serum-free culture medium (e.g. Williams' Medium E). (xii) Flush the outside of the dialysis bag with 70% ethanol and aseptically transfer the contents of the bag to a pre-cooled sterile container in ice. Using chilled pipettes, distribute the extract (Matrigel) into tubes in ice, in amounts so that one or two aliquots are sufficient for an experiment. Repeated freeze-thawing of the extract must be avoided. (xiii) Aliquots of Matrigel can be stored for some months at -20°C. (xiv) For forming gels, thaw the Matrigel, preferably at W " C , or at least so that the temperature is kept below about 10°C. Keep the thawed Matrigel on ice and use chilled pipettes and culture dishes. Dispense 0.3 ml of extract into 60-mm Lux tissue culture dishes and use a sterile, bent pasteur pipette to spread the material evenly over the dish. Place dishes in an incubator at 37°C and leave for about 60 min for firm gel formation. Gels can be kept in a humidified incubator overnight prior to use. As discussed above, larger or smaller amounts of Matrigel per dish

292


may be used and other brands of culture dish may be satisfactory. The procedure given has proved adequate in our laboratory to preserve hepatocytes for several days with spherical shape and responses of the cytochrome P-450system to phenobarbital similar to those observed in vivo.

11.4. Choice of culture media This area is the most difficult of all aspects of hepatocyte culture to summarize adequately. Modifications to culture media have been explored more widely than any other culture variable but several factors make it impossible at this time to identify a single medium as the most appropriate for hepatocyte primary culture. Because primary adult hepatocytes exhibit limited growth, systematic methods for defining optimal media based on measuring clonal growth cannot be applied, although it has been possible to develop systematically media for related cell types (liver epithelial cells, hepatoma cells or foetal hepatocytes, for instance) as noted below. For primary hepatocytes, media have in general been tested by selecting one, or a few, functional end points. It is now clear that particular components in media important for maintaining ‘differentiation’ differ for individual functions. A hormone with a positive effect on one function may have no effect or negative effects on others (e.g. Jefferson et al., 1984). Furthermore, interactions between multiple factors alter the responses observed with factors added individually. Thus the evolution of media, optimal for either growth or broad preservation of differentiated function, has proceeded by a mixture of some logic and considerable trial and error. Various cocktails of nutrients, hormones and growth factors with desirable effects have been identified, although often the importance of individual components for either growth or differentiation is still unclear. In hepatocytes on Matrigel or in co-culture, the pattern of gene expression appears to be less influenced by variations in medium com-

Ch. 1 1 .

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position (e.g. Vandenberghe et al., 1990)and requirements to preserve differentiated function are less complex than when cells are maintained on simple substrata such as collagen-coated dishes. Much of the discussion below relates to attempts to find media optimal for use with simple substrata. Serum may contribute protease inhibitor(s) that aid cell survival (Ichihara et al., 1986) but in general, factors in serum do not aid maintenance of differentiated function and may accelerate changes in culture (e.g. Reid et al., 1986). Factors in serum, particularly from rat serum, can stimulate growth in rat hepatocytes but, as the active components are isolated and characterized (e.g. McGowan, 1986; Michalopoulos, 1990), it should be possible to use purified growth factors rather than crude serum preparations to stimulate growth. It is already possible to achieve growth of foetal hepatocytes under fullydefined conditions (e.g. Hoffmann and Paul, 1990). Because inclusion of sera in culture media introduces unknown factors and may contribute to perturbation of normal differentiated function and to the growth of non-hepatocytes, a major effort in hepatocyte culture has been directed towards devising fully-defined media: the discussion below relates mainly to such serum-free media. 11.4.1. Useful media available commercially

Because of the complexity of complete media required for long-term maintenance and growth of cells in culture, it is obviously convenient to base any specific formulations for hepatocytes on commerciallyavailable media. A number of these have been used extensively with hepatocytes (e.g. Williams’ Medium E; various modifications of Eagle’s minimal essential medium (MEM) or a I:1 mixture of Dulbeccomodified MEM with Ham’s F-12; RPMI 1640; Medium 199; Leibovitz L-15; and Waymouth formulations MB 752/1 or MAB 87/3) and can be assumed to contain many of the basic requirements for cell survival. With a complex substratum such as Matrigel, use of slightly modified versions of Williams’ Medium E (Ben-Ze’ev et al., 1988), Medium 199

TABLE 11.1 Composition of some complex media developed for hepatocyte culture Components

P

Amount (mg/l)* Lanfordl WME

Ammo aciak L-alanine L-arginine HCI L-asparagine H,O Laspartic acid Lcysteine HCI L-cystine, disodium L-glutamic acid ~-glutamine GIycine L-histidine.HCI-H,O L-hydroxyproline L-isoleucine L-leucine L-lysine.HC1 L-methionine L-ornithine.HC1 L-phenylalanine L-proline L-serine L-threonine L-tryptophan L-tyrosine, disodium L-valine

N

90 60.5

20 30 52.1 23.7 50 292 50 20.3 50 75 87.5 15 25 30 10 40 10 43.5 50

Jefferson/ DMEM:F- 12 160 218 125 48 65 118 503 30 I I05 105

197 450 61 126 230 105

119 143 186 165

Reid HDM

lsom RPCD

200 56.8 20

200 56.8 20

59.15 20 300 10 20.26 20 50 50 40 15

59.15 20 300 10 20.26 20 50 50 40

Waxman/ Chee’s 72 168 I20 107

MX-83’ + Hormones 18

(255)’ 50 30 -1

15 20 30 20 5 24.86 20

15

15

20 30 20 5 24.86 20

57.3 118

584 60 84 220 439 282.4 60 132 92 84 190.4 32 89.5 314.4

59 75 58.4 100 56.74 20 I05 105 182.4 30 67.4 66 34 42 95 16 71.9 94

T

m

9

a 4

Ch. 11.

9

v,

2

s

H -s

m

8%

v,

N W

5

22 w

2

8

MONOLAYER CULTURE

2

8

m M

5

8

295

W

2

8

0

TABLE 11.1 Composition of some complex media developed for hepatocyte culture Components

N o\ *o

Amount ( & I ) * Lanford/ WME

Jefferson/ DMEM:F-12

Reid HDM

lsom RPCD

Waxman/ Chee's

MX-83' + Hormones

Nutrients

Deoxycytidine HCI Ethanolamine D-glucose Gluconolactone j3-glycerophosphate, Na Hypoxanthine Linoleic acid Methyl linoleate Oxaloacetic acid Pyruvate. Na Sorbitol Thymidine

12.77 0.061Q: 2000

2

2000

5

2

4500 178 1000

4500

25 0.084

0.044

110

50 110

100 10

0.364

Vitamins, micronutrients

p-aminobenzoic acid L-ascorbic acid Biotin Calciferol Sodium panthothenate D-calcium pantothenate Choline chloride

2000

265 25

x

U

3150

2.04 5§ 0.03

I

1 .o

I .o

2.0 0.5 1.o

0.0037

0.2

0.2

1.o 1.5

2. I 19 8.98

0.25 3.0

0.25 3.0

25 0.03

1.o 0.05 1.o 4.3

4 4

14

r b

Folk acid i-lnositol Lipoic acid Menaphthone. Na bisulphite Nicotinamide Nicotinic acid Protamine sulphate Putrescine.2HCI Pyridoxal HCI Pyridoxine HCI Retinoic acid Riboflavin Tetrahydrofolic acid Thiamine HCI a-Tocop hero1 a-Tocopherol acetate DL-a-tocopherol phosphate. disodium Vitamin A acetate Vitamin BIZ Miscellaneous HEPES Phenol red. Na Penicillin Streptomycin Gentamicin CO. in gas phase

I .o 2.0

2.66 12.51 0.103

I .o 35.0

1.o

4

35.0

7

2.019

1 .o

I .o

0.01 1.o

0.6 18.13 0.2 0.01 4

4

2 1.o

0.08 I 2.0 0.03 1

0.29 1.o

4

4

I .o

K

0

0.1

0.219

0.2

0.2

0.4

1 .o

2.169

1 .oo

0.015 0.4 0.6

1 .o

4

4

0.0012 0.004

0.0I

z

1 C

0.1

0.2

0.68

23838 10

60 I00

0.005 2383 5 60 I00

0.005 3600 5 I00 I00

0.003

0.005

5000

15 39.3 62.5 50

5%

101%)

N \o -4

TABLE 11.1 Composition of some complex media developed for hepatocyte culture h)

Components

%

Amount (mg/l)* Lanford/ WME

Hormones, gronth factors Cholera toxin Lkxamethasone EGF Glucagon Hydrocortisone Insulin Liver growth factor (Gly-His-Lys) Prolactin Somatotropin Thyrotropin releasing ( factor

Jefferson/ DMEM:F- 12

Reid HDM

lsom RPCD

Waxman/ Chee’s

MX-83’ + Hormones

0.0024 0.4 0.025 0.04

0.392

0.05 10

I0

0.06

6.25

0.039 0.14

45

0. I

0.3624

lo§

6.0

1.o 10

0.028 0.18 19:

0.66 10 mU

M)Q:

*Original formulations give amounts for some amino acids or salts using different forms (e.g. hydrates or hydrochlorides). To simplify the table. the most common form is listed and amounts giving final concentrations equivalent to the original formulation are shown. +Referencesfor media: L a n f o r M M E Based on Williams Medium E (Flow Laboratories Catalogue) with supplements (Lanford et al.. 1989) indicated by 5. Jefferson/DMEM:F-12: Based on a 1:l mixture of Dulbecco’s MEM:F-I2 (Flow Laboratories Catalogue) with modifications (Hutson et al.. 1987).Reid HCD: Based on RPMl 1640 (Flow Laboratories Catalogue) with supplcnients (Enat e l al.. 1984). lsom RPCD: Based on RPMl 1640 with trace elemenls (Hutchings and Sato. 1978) plus other supplements (Woodworth et al.. 1986: lsom et al.. 1987). WaxmadChee’s: Chee’s Medium (GIBCO Formulation) plus dexamethasone. insulin. transferrin and Na,SeO; (Waxman et al.. 1990).MX-83 + Hormones: (Hoffmann et al., 1989). In addition to the components in the Table. MX-83 also contains (mg/l): adenine (0,135). D(-) ribose (0.5). inosine (10). malic acid (30). succinic acid (100). dithiothreitol ( I ) . arachidonic acid (6. lo-’),cholesterol (0.06). soybean lecithin (0.12) and sphingomyclin (0.02). Arginine is normally omitted.

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299

(Bissell et al., 1987) or Waymouth MB 752/1 (Schuetz et al., 1988; Guzelian et al., 1989) supplemented with insulin f glucocorticoid have all yielded cultures in which a variety of differentiated functions are well preserved. In co-cultures with rat liver epithelial cells a number of studies, in which albumin synthesis or other differentiated functions were preserved, utilized Ham’s F-12 (e.g. Guguen-Guillouzo et al., 1983) or a 3:1 mixture of Eagle’s MEM/Medium 199 (e.g. Begue et al., 1984). Because the precise composition of media may be much less critical for hepatocytes on Matrigel or in co-culture, the effects of varying the composition of basic culture media (i.e. content of amino acids and carbohydrates, buffer, salts and trace metal ions, vitamins and other micronutrients) has not yet been examined in any detail but there is more information for adult hepatocytes or related cells on collagen-coated dishes. A few off-the-shelf media have been used with defined supplements in combination with simple substrata to support multiple cell divisions in some type of hepatocyte (e.g. foetal, primate) or related cell-types (liver- or hepatoma-derived cell-lines or virus-transformed hepatocytes) and thus must contain most of the basic requirements for hepatocyte growth. Williams’ Medium E was developed by Williams and coworkers to enhance liver phenotypic expression in liver-derived epithelial cell-lines in addition to supporting their growth in the presence of serum (Gunn et al., 1976). It has been used extensively (with supplements) in neonatal or adult rat hepatocyte cultures to support limited DNA synthesis and mitosis (e.g. Nakamura et al., 1988; Yusof and Edwards, 1990) and recently has been used as the basis of a medium devised by Lanford et al. (1989) for supporting growth and extended maintenance of differentiated function in baboon hepatocyte cultures (see Table 11.1). RPMI 1640 is the basis of defined media developed by Reid and co-workers, originally to optimize the serum-free growth of hepatoma cells (Gatmaitan et a]., 1983)and adapted for use with hepatocyte primary cultures, as discussed by Reid et al. (1986) (see Reid HDM in Table 11.1). Sells et al. (1985) used a variation of HDM to support serum-free growth of cells from human

300


foetal hepatocyte cultures. RPMI 1640 is also the basis of RPCD medium used by Isom and co-workers to support serum-free growth of SV40-transformed hepatocytes (Woodworth et al., 1986) and, in combination with DMSO (2%) and EGF, to support extended maintenance of adult rat hepatocytes (Isom et al., 1987) (see Isom RPCD in Table 11.1). A 1:l mixture of Dulbecco-modified MEM/Ham’s F-12 was reported by Sawada et al. (1986) to be the best of several media tested for supporting DNA synthesis in adult rat hepatocytes. It supported multiple rounds of replication in mouse hepatocyte cultures (Sawada et al., 1988) and was the basis for defined media used by Salas-Prato et al. (1988) to support growth of human foetal hepatocytes, and by Shapiro and Wagner ( 1 988) to grow H-35 hepatoma cells. The Dulbecco’s MEM/F-12 mixture was also the basis for an amino acid- and hormone-supplemented medium used by Jefferson and co-workers to preserve high levels of albumin synthesis (Hutson et al., 1987 - see discussion below and Table 11.1). Thus Williams’ Medium E, RPMI 1640 and the Dulbecco’s MEM/F- 12 mixture are three commercially-available media which supply many of the requirements for growth of hepatocyte-like cells and seem to be generally useful as basic media for primary hepatocyte culture. With appropriate modifications they may be appropriate for studies on growth or differentiated function. Undoubtedly further work will identify other media in this category. A new fully-defined medium, MX-83 (see Table 11. l), which supports multiple cell divisions and good preservation of differentiation in foetal rat hepatocyte cultures has recently been described by Hoffmann et al. (1989). MX-83 is not available commercially. Although not shown to be desirable for studies on growth, Leibovitz L-15 with simple supplements, supports relatively good hepatocyte survival (e.g. Sawada et al., 1987) and allows studies with air in the gas phase (i.e. without added CO,). More strikingly, Chee’s medium, a substantially-modified version of Eagle’s MEM, has recently been shown to allow cell survival for 4 - 5 weeks on collagen-coated dishes and excellent preservation of at least some differentiated functions in

Ch. 1 1 .

MONOLAYER CULTURE

30 1

simple cultures where the basic insulin-containing medium is supplemented only with dexamethasone, transferrin and selenium (Jauregui et al., 1986; Waxman et al., 1990). While culture media designed to provide basic requirements of buffer, salts, amino acids, vitamins and other nutrients have mainly been compared as complete media, there have been some systematic studies on the importance of individual components. 11.4.2. Amino acid requirements

Schwarze et al. (1982) and Hutson et al. (1 987) have investigated utilization of amino acids in serum-free cultures and have shown rapid depletion of some amino acids including glycine, proline, arginine and phenylalanine. Both groups found that even in the presence of insulin, added to favour anabolic protein metabolism, amino acid concentrations 5-10 times those in rat plasma are required for N-balance and maximal protein synthesis. To offset depletion of various essential amino acids, high initial levels are required in media. A high amino acid modification of Dulbecco’s MEM/F-12 (1:l) devised by Hutson et al. (1987) is shown in Table 11.1. This medium with insulin and glucocorticoid sustains albumin secretion at rates comparable with perfused liver for at least 4 days. Seglen’s laboratory also formulated a medium (SM- 1) based on requirements for maximizing protein synthesis (Schwarze et al., 1982). However there have been relatively few studies so far on broader effects of these high amino acid media in culture (but see Turner et al., 1988). High amino acid concentrations may be one of the beneficial features of Chee’s medium. In relation to growth, amino acid levels appear to have a complex role in vivo (Fausto and Mead, 1989). In culture, the difference in the ability of different media to support DNA synthesis appears to depend mainly on proline content: 30 mg prolineA(O.29 mM) was found to be optimal (Nakamura and Ichihara, 1985). It appears that high amino acid levels are not required for growth of foetal hepatocytes (e.g. MX-83 of Hoffmann et al., 1989). In adult cultures, some amino

302


acids at high concentration (e.g. alanine, glutamine > 2 mM) inhibit DNA synthesis (McGowan et al., 1984). A common approach to making culture media ‘selective’ for hepatocytes, as opposed to fibroblasts and at least some other possible contaminating cells, is based on the ability of hepatocytes to convert ornithine to arginine. Schwarze et al. (1982) showed that in arginine-free medium, ornithine in the range 0.1 to 1.0 mM was as effective as arginine in supporting hepatocyte protein synthesis and that despite steady consumption of ornithine, arginine remained low in the culture medium. This suggests that there is unlikely to be substantial cross-feeding of any arginine-dependent cells in hepatocyte cultures. Foetal as well as adult hepatocytes survive and grow in arginine-free media containing 0.4 mM ornithine (Leffert and Paul, 1972; Hoffmann et al., 1989). As discussed by McGowan (1986), one reservation about the selectivity of arginine-free media is the possibility that potential contaminants in culture such as Kupffer cells or liver stem cells may have some ability to convert ornithine to arginine. 11.4.3. Carbohydrates

Seglen’s laboratory investigated energy requirements for sustaining protein synthesis and found lactate or pyruvate to be better substrates than glucose in the 5-20 mM range. High glucose concentrations (> 20 mM) result in net glucose utilization and probably reduce depletion of amino acids for energy generation (Schwarze et al., 1982). Carbohydrates included in media have generally been viewed as energy sources and their effects on patterns of gene expression have not been examined in detail. Some effects of high glucose concentrations (around 20 mM) such as induction, in combination with insulin, of L-type pyruvate kinase (Decaux et al., 1989) and repression of ornithine aminotransferase (Spence et al., 1980) may reflect relatively specific controls at either transcriptional or post-transcriptional levels (Decaux et al., 1989; Jacoby et al., 1989). High glucose also helps maintain normal levels of alcohol dehydrogenase (Grunnet et al., 1989),minimizes

Ch. 11.

MONOLAYER CULTURE

303

the appearance of the foetal liver enzyme y-glutamyltranspeptidase in adult hepatocyte cultures (Edwards, 1982) and has been reported to modulate c-myc mRNA levels in hepatoma cells (Briata et al., 1989), observations which suggest wider effects of glucose or its metabolites on gene expression. Synergism between 1 mM glucose and lactate or pyruvate in causing strong stimulation of DNA synthesis has been demonstrated (McGowan et al., 1984) and some complex media such as Chee’s or MX-83 (Table 11.l), recently used with hepatocytes, contain novel combinations of glucose with other carbohydrates. However, whether the particular combination of carbohydrates present in media is important for broad hepatocyte function in vitro is still unclear. 11.4.4. Salts and trace metal ions

Nearly all the media commonly used with hepatocytes contain 1-2 mM Ca2+,approximating plasma free Ca2+concentrations of about 1.2 mM. There have been reports that adult hepatocytes undergo mitosis more readily at low Ca2+concentrations (0.4 mM) than at 1.2 mM (Hasegawa et al., 1982; Eckl et al., 1987) and it was suggested that high Ca2+may favour differentiation rather than growth, as in keratinocytes. However in our hands, the initial round of DNA synthesis (as distinct from mitosis) in EGF- and insulin-stimulated hepatocytes is only weakly dependent on added extracellular Ca2+but optimal at 1.2 mM (Petronijevic and Edwards, 1990); and other laboratories have observed high levels of mitosis at 1 mM Ca2+ (Michalopoulos et al., 1982; McGowan, 1986).The overall importance of Ca2+ concentration to growth and differentiation in primary hepatocytes is still unclear. Table 1 1.1 shows a variety of trace metal ion mixtures included in hepatocyte culture media. Inclusion of Cu*+, Zn2+and selenite in Reid’s HCD was a modification, introduced for primary cultures, of the requirements for serum-free clonal growth of hepatoma cells on collagen (Gatmaitan et al., 1983). The trace metals in Isom’s RPCD and Jefferson’s modification of Dulbecco’s MEM/F-12 are based on

304


requirements for serum-free growth of HeLa cells (Hutchings and Sato, 1978) together, in the latter medium, with the relatively high Fe2+and Zn2+content from F-12. The beneficial effects of selenite at concentrations from 3-100 nM on differentiated functions have been reported by several laboratories (e.g. Newman and Guzelian, 1982; Meredith, 1987; 1988). Increasing Zn2+to pM concentrations (comparable with Zn2+ levels in F-12, Chee’s or MX-83 media) was reported to stimulate mouse hepatocyte DNA synthesis markedly (Kobusch and Bock, 1990)although, in our hands with rat hepatocytes, there was little effect of 10 pM Zn2+compared with the 70 pM Zn2+ present in Williams’ Medium E. At 30 pM, Zn2+ was toxic. Hepatocytes have high levels of transferrin receptors (Sciot et al., 1990) and, although differentiated hepatocytes synthesize transferrin, there is some evidence that added transferrin enhances survival and preservation of normal functions (e.g. Meredith, 1987; 1988).It also enhances the serum-free growth of H-35 hepatoma cells with optimal effects at 5 &ml (Shapiro and Wagner, 1988). Transferrin presumably increases delivery of iron to hepatocytes (Laskey et al., 1988). Iron salts and/or transferrin are present in many of the complex media (but not Reid’s HDM) developed for use with liver cells (see Table 11.1). On the basis of the present data for metal ions it seems reasonable to include Zn2+, Cu2+,Se0:- and Fe2+or Fe3+,possibly with transferrin, in a complex medium for hepatocytes. 11.4.5. Other components

The essential fatty acid linoleic acid bound to serum albumin (or alternatively, methyl linoleate) is included in most media. One documented effect in primary cultures is prolonged preservation of gap junctions (Spray et al., 1987). Low levels of albumin have been included in media, partly to bind linoleic acid, but Gebhardt and Mecke (1979b) suggested that high albumin concentrations (41 mg/ml) greatly reduced the formation of lipid droplets, frequently observed in monolayers after 3 4 days. Ethanolamine (1-65 pM) has been included in some recent for-

Ch. 11.

305

MONOLAYER CULTURE

mulations (e.g. Lanford et al., 1989; Salas-Prato et al., 1988; Chan et al., 1989) and ascorbate (0.3 mM) has been used as a routine supplement by some laboratories (Bissell and Guzelian, 1980; Bissell et al., 1987; Schuetz et al., 1988; Ben-Ze'ev et al., 1988). For these and other micronutrients or vitamins listed in Table 1 1.1 there are, understandably, few detailed studies on the importance of the individual components to overall hepatocyte preservation and function. Inclusion of phenol red provides a convenient check on the pH of media but is known to have oestrogenic activity (e.g. Dumesic et al., 1989), which may be undesirable in some studies. 11.4.6. Antibiotics

=

A combination of penicillin (100-160 unitdm1 60-100 pg/ml 3 170-280 pM) and streptomycin (100 pdml = 70 pM)may delay the overgrowth in cultures of any contaminating gram-positive or gramnegative bacteria respectively. Gentamicin (50 pg/ml 90 pM) has been used increasingly as an alternative for preventing gram-negative and -positive bacterial growth. At these concentrations there are no adverse effects on cell integrity (by trypan blue exclusion) and little effect on protein synthesis except for the higher concentrations of penicillin which may inhibit 10-20'%, (Schwarze and Seglen, 1981: Vonen and Mcirland, 1982; Jeejeebhoy et al., 1975). Kanamycin (60 pg/ml) has also been used for long-term maintenance of hepatocytes (Miyazaki et al., 1985). Although it is usually unnecessary to add a fungicide, fungizone (amphotericin) at 2.5 pg/ml may be added to cultures, preferably for short periods up to 1 day, to minimize yeast and fungal growth. Amphotericin (0.5-1.0 pg/ml) has been routinely included in media by a few laboratories (e.g. Miyazaki et al., 1985; Vintermyr and Doskeland, 1987).

=

11.4.7. Oxygenation

While hepatocytes can apparently survive for a few hours at O2partial pressures as low as 0.1 mmHg or 0.013 kPd (Anundi and de Groot,

306


1989), Jeejeebhoy et al. (1975) showed that for hepatocytes in suspension culture, a PO, as high as 20 kPa was optimal for plasma protein synthesis. For confluent monolayer cultures in 60-mm plastic dishes, Holzer and Maier (1987) found that the PO, in the culture medium was about 60 mmHg (8 kPa) when cultures were maintained in CO,/air (1:19): this PO, falls within the range of 0,-concentrations in the liver lobule in vivo of 9 kPa in the periportal zone down to about 4 kPa in the pericentral zone. It seems that standard culture conditions fortuitously provide appropriate oxygenation of hepatocytes. Bissell and Guzelian (1980) noted that the depth of medium overlying monolayers was important and should not exceed 3.0 ml per 60-mm dish, presumably to allow adequate gas exchange. Lowering the 0,-concentration was shown to improve cell survival slightly and to influence carbohydrate metabolism and some other properties of cultures (Wolfle et al., 1983; Holzer and Maier, 1987), whereas an increased proportion of 0, in the incubator atmosphere led to markedly decreased survival after 7 days, apparently as the result of greater formation of reactive oxygen species (Miyazaki et al., 1989). 11.4.8. Hormonal supplements useful for maintaining dcferentiated functions

While numerous hormones have been shown to influence specific activities in hepatocytes there is reasonable evidence of relatively broad effects of six or seven different hormones, individually or in combination, on hepatocyte survival and differentiated function. All of these (except thyroid hormones) appear in one or more of the media listed in Table 1 1.1. Where hormones are added individually to typical, commerciallyavailable culture media, glucocorticoids are probably the agents with most marked effects on the morphology, function and survival of hepatocytes on simple substrata. Survival of adult (Laishes and Williams, 1976b) and neonatal (Marceau et al., 1982a) hepatocytes in monolayers is extended and retention of a relatively compact, polygonal cell shape with more numerous bile canalicular-like struc-

Ch. I I .

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307

tures is favoured by incubation with dexamethasone or hydrocortisone. Dexamethasone modulates the production of extracellular matrix by hepatocytes, increasing the laying down of fibronectin-rich material on culture dishes (Marceau et al., 1982b). It also favours an ordered arrangement of the intracellular cytoskeleton, more like that in the intact liver than for cultures without glucocorticoid (Marceau et al., 1986a). Studies in which more than 3,000 liver proteins were resolved on giant two-dimensional gels showed that synthesis of 80 or more proteins changed during the first 20 h after placing hepatocytes in culture. Addition of 100 nM dexamethasone retarded or reversed about half of these changes in addition to inducing or repressing a further group of proteins (Colbert et al., 1985). As suggested by these effects on patterns of protein synthesis, glucocorticoids directly influence a wide range of activities in culture, in general favouring better preservation of tissue-specific functions including albumin synthesis (see Guguen-Guillouzo and Guillouzo (1983) and Reid et al. (1986) for references). Glucocorticoids also have permissive effects in the action of other hormones (see Guguen-Guillouzo (1983) for references) and in effects of xenobiotics on hepatocytes (e.g. Edwards et al., 1984, 1987a). Reid et al. (1986) summarized evidence that effects of glucocorticoids in culture are predominantly on mRNA stability rather than transcription. There have been conflicting findings on the extent to which glucocorticoids influence transcription in serum-free cultures (e.g. Jefferson et al., 1985; Nawa et al., 1986). Concentrations of dexamethasone ranging from 10 nM to 10 pM have been employed in culture. Actions of dexamethasone such as induction of tyrosine aminotransferase (which are likely to be mediated by the classical glucocorticoid receptor) are maximal at about 30-100 nM although much higher concentrations of natural glucocorticoids such as hydrocortisone (50 pM) are required, presumably because of rapid steroid metabolism in cultured hepatocytes (see Edwards et al., 1987a, for references). Relatively low concentrations of dexamethasone (30-100 nM) are also adequate to suppress fibroblast growth and to contribute, alone or in combination with other hormones, to the preser-

308


vation of hepatocytes (e.g. Edwards and Lucas, 1982; Dich et al., 1988). Pharmacological levels of dexamethasone (0.3-10 pM) may exert additional effects such as induction of y-glutamyltranspeptidase or of cytochrome P-450 isozyme(s) via a mechanism not necessarily involving the classical glucocorticoid receptor (e.g. Schuetz and Guzelian, 1984; Edwards et al., 1987a). Insulin is almost invariably included in media used for hepatocyte culture, both to enhance initial cell attachment (Section 1 1.2.3) and to improve survival and preservation of differentiated functions. When added alone, insulin has been shown to improve survival of hepatocytes on collagen-coated dishes (e.g. Dich et al., 1988). The hormone increases overall protein synthesis (Tanaka et al., 1978) and secretion of albumin (e.g. Hutson et al., 1987), as well as influencing various activities relating to carbohydrate metabolism and lipogenesis (see Guguen-Guillouzo and Guillouzo 1983 for references). These effects appear to be maximal at about 100 nM insulin (e.g. Hutson et al., 1987). Contributions of insulin to preserving hepatocyte differentiated functions may involve synergism with glucocorticoids (e.g. Hutson et al., 1987) and other combinations of hormones and/or nutrients (e.g. Reid et al., 1986; Dich et al., 1988; Decaux et al., 1989; Tong et al., 1990). Although Clayton et al. (1985) found that no combination of hormones could maintain transcription of liver-specificgenes in mouse hepatocyte cultures, Lloyd et al. (1987) reported that in rat hepatocytes, insulin stimulated albumin gene transcription and, in combination with dexamethasone and glucagon, could maintain albumin transcription for a limited period (4 days) at a level comparable with that in freshly isolated hepatocytes. Glucagon alone does not improve hepatocyte survival (e.g. Dich et al., 1988). In addition to relatively specific effects in the induction or repression of regulatory enzymes in carbohydrate, lipid and amino acid metabolism (e.g. Ichihara et al., 1982), there is evidence that when combined at appropriate concentrations (0.1-1 nM) with glucocorticoid and insulin (Nawa et al., 1986; Lloyd et al., 1987; Dich et al., 1988) or with more complex hormone mixtures (Reid et al., 1986),

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glucagon may contribute to preserving a variety of liver functions including albumin synthesis. EGF was initially included in hepatocyte culture media for its effects on growth. However it also appears to contribute (at concentrations of 25-50 ng/ml) to preserving hepatocyte morphology and tissue-specific functions such as albumin synthesis (Jefferson et al., 1984; Isom et al., 1987; Tong et al., 1990). With respect to other hormones included in media for hepatocyte preservation, there is less information on their individual contributions. Growth hormone (somatotropin) and prolactin apparently have useful effects in the culture systems of Reid et al. (1 986) and Tong et al. (1990), although their individual contributions to cell preservation are not well documented. Growth hormone at concentrations of 1 6 - 5 0ng/ml (approximately equivalent to 1 2 - 6 0 pU/ml) increases synthesis of insulinlike growth factor-1 (Johnson et al., 1989), modulates synthesis of various gender-specific, and xenobiotic-inducible, cytochrome P-450 isozymes (Guzelian et al., 1988; Schuetz et al., 1990)and helps preserve alcohol dehydrogenase activity after several days in culture (Grunnet et al., 1989) but does not influence albumin synthesis (Schuetz et al., 1990). Receptors for tri-iodothyronine (T,) are depleted at early times in culture but restored after a few days (Goglia et al., 1985). In addition to induction of some specific enzymes (e.g. Ichihara et al., 1982), T, (1 FM) was shown to stimulate total protein synthesis in 6-day-old cultures (Gallo et al., 1987). 11.4.9. Hormones and growth factors with growth-stimulatory effects

Hepatocytes from foetal, neonatal or suckling rats undergo one or more rounds of cell division in culture and studies with serum-free media have defined hormones and growth factors which support at least limited growth (see Section 11.10.). With foetal hepatocytes for instance, MX-13 medium, containing EGF, insulin and hydrocortisone, supports multiple cell divisions.

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In adult rat hepatocyte culture it is possible to stimulate DNA synthesis in almost all the cells in cultures on films of collagen (Types I or IV) or fibronectin, if cells are plated at lower densities. However, as noted in Section 1 1.1.7., these cells do not necessarily complete the classical cell cycle. At the present stage of studies on growth in adult hepatocytes, identification of growth-stimulatory factors is based mainly on ability to stimulate DNA synthesis rather than cell division. Stimulatory hormones and growth factors have been discussed by McGowan (1986) and Michalopoulos (1990) and these excellent reviews should be consulted for more detailed information. Michalopoulos ( 1990) makes the distinction between ‘complete mitogens’ such as EGF which can act alone to stimulate DNA synthesis and ‘co-mitogens’ which act only in combination with EGF or other mitogens. A number of hepatomitogens have now been identified. EGF was the first growth factor shown to cause strong stimulation of hepatocyte DNA synthesis (Richman et al., 1976) and it has had a central role in most studies on hepatocyte growth. For a period of about 16 h after isolation and plating in culture, adult hepatocytes are relatively insensitive to growth-stimulatory effects of EGF but addition of EGF thereafter at concentrations in the range 10-50 ng/ml (1.7-8.3 nM) stimulates initiation of DNA synthesis within about 20 h (e.g. Sand and Christoffersen, 1987; Vintermyr and Duhkeland, 1987; Wollenberg et al., 1989). Serum from various sources has been shown to stimulate DNA synthesis, with rat serum exhibiting the largest effects (McGowan, 1986). Michalopoulos et al. (1982) showed that rat serum (30-50% v/v) stimulated DNA synthesis in a high proportion of cultured cells and these investigators subsequently isolated two active components, hepatopoietins A and B. Hepatopoietin A is a 105,000 kDa heterodimeric protein, apparently identical to serum factors including ‘hepatic growth factor’, isolated by other workers (see Michalopoulos, 1990, for discussion). Hepatopoietin B is a small glycolipid-like molecule which is a complete mitogen but interacts synergisticallywith EGF and hepatopoietin A. Other hepatomitogens include a-TGF and

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basic fibroblast growth factor, both active at 100 ng/ml (Hoffmann and Paul, 1990) and acidic fibroblast growth factor (also called heparin binding growth factor-1) which is active at 50 ng/ml, only in the presence of 10-50 pg heparin/ml (Houck et al., 1990). Synthetic or recombinant forms of the last three factors are becoming available. However at present, limited availability or high cost restrict the use of all the mitogens other than EGF for extensive studies on growth. Although less potent than EGF, some prostaglandins may also be hepatomitogens, effective at relatively high concentrations (about 10 pM) (Skouteris et al., 1988). Findings in our laboratory suggest that prostaglandins act synergistically with EGF and that the major prostaglandin in liver, prostaglandin D,, is more potent in stimulation of DNA synthesis than prostaglandins E, and F,a (Lee and Edwards, unpublished). In media with appropriate nutrients, EGF by itself stimulates DNA synthesis. However its effect is increased in a synergistic fashion by exposure of cells to insulin (optimal concentration, 100 nM). Insulin is most effective when added at early culture times (e.g. Sand and Christoffersen, 1987) and the presence of insulin for 2 4 h during cell attachment is sufficient to increase substantially the subsequent response to EGF (Yusof and Edwards, 1990). Inclusion of insulin in media after attachment further augments the response to EGF, although insulin alone has only a small effect (e.g. McGowan, 1986; Sand and Christoffersen, 1987) and it is thus considered a ‘co-mitogen’. In the presence of mitogens, some further co-mitogens cause strong stimulation of DNA synthesis. Norepinephrine (optimal concentration, 10-30 pM), or other agonists acting via the a,-adrenergic receptor, exhibit strong synergism with EGF. Their effect is greatest on the first day in culture and declines with time (Cruise and Michalopoulos, 1985), probably due to progressive loss of a,-receptors in culture (Nakamura et al., 1984). Oestrogens are also effective co-mitogens. Concentrations of oestradiol(30 pM) or ethinyl oestradiol(20 pM) optimal for stimulation of DNA synthesis (e.g. Shi and Yager, 1989; Lee and Edwards, 1990)are greatly in excess of levels

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normally adequate to bind the oestrogen receptor. However the requirement for such high concentrations may reflect the very rapid metabolism of oestrogens in hepatocytes (Standeven et al., 1990). Some other agents with weaker co-mitogenic action include vasopressin (McGowan, 1986) and prolactin (Lee and Edwards, unpublished). Somatotropin has been included in some media designed to support hepatocyte differentiation and growth (Table 1 1.1) but in our laboratory, it had little effect on DNA synthesis in adult hepatocytes. Two other hormones frequently included in growth media (Table 11.1) are glucagon and glucocorticoids. The response to these agents may depend on factors such as cell density or other media components present. Glucagon was reported to act synergistically with insulin to increase EGF-stimulated replication (e.g. McGowan et al., 1981) but subsequent studies reported variable stimulatory or inhibitory effects, possibly dependent on the nature of the experimental conditions (e.g. Bronstad et al., 1983; McGowan, 1986; Kubin et al., 1989). Studies in our laboratory under a variety of conditions suggest that glucagon or dibutyryl cyclic AMP, in general, inhibit DNA synthesis (as stimulated by EGF and various co-mitogens) except for small stimulatory effects when added in the first few hours of culture. Glucocorticoids (e.g. 30-100 nM dexamethasone) have been reported to exert both stimulatory (McGowan, 1988; Yusof and Edwards, 1990) and inhibitory (e.g. Vintermyr and DBskeland, 1989) effects on DNA synthesis. It appears likely that glucocorticoids influence DNA synthesis only indirectly so that their net effects depend on the particular combination of nutrients, hormones and cell density present. Our laboratory has reported stimulatory effects of 30 nM dexamethasone in combination with EGF and some other growth stimulators (Yusof and Edwards, 1990) but the same level of dexamethasone is inhibitory in combination with oestrogens or 20 mM pyruvate. The choice of hormones and growth factors for studies on growth will obviously depend on the object of the investigation. No combina-

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tion of factors so far tested supports continuous proliferation of adult rat hepatocytes but one or two rounds of DNA synthesis can be induced in a high proportion of cells (about 80%) by various combinations of EGF, insulin and serum-derived factors. Oestrogens or a,-agonists may favour earlier and more synchronized entry of hepatocytes to S-phase and, depending on other factors present, increase the number of cells active in replication. Glucocorticoids appear to act synergistically with some growth stimulators but block the effects of others. 11.4.10. Effects of proteoglycans or glycosaminoglycans

Reid and her co-workers reported that supplementation of HDM medium (Table 1 1.1) with certain proteoglycans or corresponding glycosaminoglycans resulted in more compact, less flattened monolayers, induced restoration of gap junctional communication between hepatocytes, increased cytoplasmic mRNA concentrations for albumin, ligandin and some other liver-specificfunctions and decreased the mRNAs for cytoskeletal proteins, actin and tubulin. The effects on mRNA levels were due to both transcriptional and posttranscriptional regulation of the mRNAs (Reid et al., 1986; Spray et al., 1987; Fujita et al., 1987). Chondroitin sulphate proteoglycan (10 pglml, from bovine nasal cartilage) and dermatan sulphate proteoglycan (10 pg/ml, from bovine articular cartilage) are much more effective than their corresponding glycosaminoglycans, dermatan sulphate and chondroitin sulphate. Heparan sulphate, the glycosaminoglycancomponent of the most common liver proteoglycan, is inactive but heparin from various sources is effective in increasing liver-specific gene expression and gap junctional communication. With high concentrations of heparin (100 pg/ml), monolayers tend to contract and detach in sheets. The effect of heparin is dependent on its source. A heparin preparation from bovine liver at 20 pglml restores gap junctional communication between nearly all cells in culture

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whereas preparations from bovine lung or pig intestine are less effective (Fujita et al., 1987; Spray et al., 1987). Reid et al. (1986) consider that additions of proteoglycans or glycosaminoglycans restore contacts with biologically-active matrix components that are present in vivo. There is evidence for uptake of glycosaminoglycans and their transport into the nuclei of liver epithelial cells (Ishihara et al., 1986).It is not clear whether the effects of heparin depend on uptake or binding to cell-surface receptors (Kirch et al., 1987). 11.4.11. Selective media

The potential for outgrowth in primary cultures of non-hepatocytes, present as contaminants in the original cell preparation. was outlined in Section 10.3.3. The growth of fibroblasts or liver epithelial cells is favoured by serum. The use of serum-free media reduces, but does not necessarily eliminate, the potential for overgrowth by contaminating cells (e.g. Edwards and Lucas, 1982). A widely used approach to prevent fibroblast growth and possibly that of liver epithelial cells has been to replace arginine with ornithine in culture media (Leffert and Paul, 1972) as discussed in Section 11.4.2. More recent studies have suggested that complex serum-free media with hormonal supplements may be effective in minimizing growth of non-hepatocytes, even without the use of arginine-free media. For instance, Reid et al. (1986) using HDM medium (Table 11.1)and Isom et al. (1987) using RPCD medium (Table 11.1) with 2% DMSO reported good preservation of hepatocytes with minimal contamination. Glucocorticoids have been reported by several laboratories to suppress fibroblast growth (e.g. Marceau et al. 1980; Edwards and Lucas, 1982) but they allow the growth of liver epithelial cells (Miyazaki et al., 1982). Tsao and Liu (1988) reported that EGF inhibits proliferation of most newly-derived or early-passage liver epithelial cells. Thus, omission of serum and the presence of EGF and

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glucocorticoid may provide conditions under which the most significant potential contaminants in hepatocyte cultures cannot grow.

11.5. Non-physiological supplements to culture media 11.5.I. DMSO

Isom et al. (1985) first reported that adult hepatocytes, maintained on collagen-coated dishes in medium supplemented with hormones and 2% DMSO, survive and preserve substantial levels of albumin synthesis for several weeks - much longer than when cells are cultured in hormone-supplemented medium alone. In cultures with DMSO, a number of the cells plated initially detach within 2-3 days but the remainder are well preserved in islands with more compact, cuboidal morphology than corresponding cultures without DMSO. DMSO prevents DNA synthesis in adult hepatocytes, apparently by blocking cells in Go or early G, phases of the cell cycle (Chan et al., 1989). The effects of DMSO on morphology and survival, detectable at 0.5%) and optimal at 2%, are dependent on the presence of dexamethasone and possibly insulin (McGowan, 1988). Isom et al. (1987) found that inclusion of EGF (25 ng/ml) further improved preservation of hepatocyte morphology in cultures maintained with RPCD medium (Table 11.1) plus 2% DMSO. These cultures undergo some initial changes in gene expression but then maintain relatively constant function from about 6-40 days in culture. The levels of mRNA for albumin after 40 days are 45% of in vivo levels and for other liverspecific genes examined, range from &72%1 (Isom et al., 1987).DMSO also helps preserve the glutathione-S-transferase subunits typical of adult liver while preventing appearance of transferase subunit 7 (Vandenberghe et al., 1990) and also of y-glutamyltranspeptidase(Edwards et al., 1987b), both of which are typical of foetal liver. Despite its effects on long-term survival and partial preservation of normal differentiated function, DMSO did not support induction of the

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mRNAs for the phenobarbital-inducible cytochrome P-450isozymes, even on Matrigel (Guzelian et al., 1989). As discussed by Isom et al. (1987), the ability of DMSO to improve the retention of many liver-specific proteins may involve some effects at transcription but for some genes, including that for albumin, the major effect is on mRNA stability so that patterns of RNA metabolism unlike those in vivo underlie apparent normality at the protein level. While DMSO is known to be a differentiating agent in erythroleukemia cells and some other cell-lines, its mechanism of action is not understood. Based on its solvent properties, effects on hepatocyte membrane properties and transport have been suggested. Along with other agents which favour hepatocyte preservation (phenobarbital, butyrate), DMSO has been shown to inhibit lipid peroxidation in cultures (McGowan, 1987). Consistent with its effect on cell shape, DMSO was recently shown to have marked effects on the cytoskeletal organization of hepatocytes, in parallel with a rapid elevation of cytoplasmic Ca2+possibly involving both release from intracellular stores and influx across the plasma membrane (Yamamoto, 1989). DMSO action differs from effects of co-culture in that it does not promote the laying down of extracellular matrix (Goulet et al., 1988). Despite being a non-physiological effector, DMSO currently provides one simple tool for achieving relatively stable cultures with many near-normal functions. As recently discussed by Chan et al. (1989), DMSO may also be a useful tool in studies on growth since it apparently promotes re-acquisition of mitotic competence in adult hepatocytes that have become quiescent after one or two rounds of DNA replication in culture. A number of other agents, when added to primary cultures, aid survival and at least partial preservation of some differentiated functions. These do not have obvious advantages over DMSO for broad preservation of function but they appear to work by differing mechanisms. Exploring their effects should help clarify the reasons for altered hepatocyte function in culture and perhaps indicate means for correcting it.

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11.5.1. Phenobarbital

Miyazaki and Sat0 and their colleagues (Miyazaki et al., 1985; 1989) have shown that inclusion of phenobarbital (3 mM) in culture media, particularly in combination with insulin and dexamethasone, allows preservation of hepatocytes on plastic or collagen-coated dishes for several weeks. The cells retain morphology similar to that on day 1 in culture and proliferation of fibroblasts or clear epithelial cells is minimal even in medium with 20% bovine serum. There is an initial decline in activities such as albumin secretion and glucose-6-phosphatase but after 7 days these are stabilized at 1&200/0 of initial values. Miyazaki et al. (1989) suggest that the effect of phenobarbital may result from reduction of oxidative stress in hepatocytes. The structural specificity for stabilization of cultures by barbiturates (Miyazaki et al., 1987a; 1987b) appears to differ somewhat from structural specificity relating to the induction of liver drug-metabolizing enzymes or liver hyperplasia in vivo (e.g. Nims et al., 1987). Also, the concentration of phenobarbital effective in stabilizing cultures (2-3 mM) appears to be slightly higher than that optimal for more specific effects on hepatocyte function in culture (0.75-2 mM), such as induction of cytochrome P-450b (e.g. Schuetz et al., 1988; Waxman et al., 1990) or stimulation of DNA synthesis (e.g. Yusof and Edwards, 1990). At the higher concentration (3 mM), phenobarbital resembles DMSO in inhibiting DNA synthesis while also improving the competence of hepatocytes for DNA synthesis after removal of the inhibiting agent (Chan et al., 1989).

11.5.3. Nicotinamide Some of the effects of nicotinamide on hepatocytes appear to be relatively selective. For instance improved preservation of total cytochrome P-450 reflects effects, predominantly on a single isozyme, possibly by inhibition of its degradation (Steward et al., 1985). The possibility that nicotinamide and some related compounds may have broader effects on hepatocyte function is suggested by findings that

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10-25 mM nicotinamide delays the appearance of foetal liver proteins such as y-glutamyltranspeptidaseand pyruvate kinase K in adult cultures (Althaus et al., 1982; Edwards, 1984) and extends survival and partial expression of differentiated functions for up to a month (Inoue et al., 1989). Althaus et al. (1982) suggested that the effects of nicotinamide may be related to inhibition of ADP ribosylation, an activity shown to increase markedly in hepatocytes during the first 3 days in culture. ADP-ribosylation of nuclear proteins modifies chromatin structure and also rapidly depletes cellular NAD levels: either of these events may influence the differentiation and/or growth of cells (see Althaus et al., 1982; Inoue et al., 1989 for discussion). 11.5.4. Butyrate

At 5 mM, addition of butyrate to media retards flattening of hepatocytes and favours preservation of polyhedral morphology in monolayers (Gladhaug et al., 1988). It approximately doubles the period during which hepatocytes survive on plastic or collagen substrata (Staecker et al., 1988); and consistent with effects favouring differentiation in various cell types, it helps preserve basal tyrosine aminotransferase activity and reduces the rate of appearance of yglutamyltranspeptidase (Edwards et al., 1987b; Staecker et al., 1988). Butyrate ( 5 mM) acting synergistically with dexamethasone, also favours preservation in culture of the EGF receptor levels, particularly high-affinity receptors, typical of freshly isolated hepatocytes. This preservation of high-affinity EGF receptors is associated with inhibition of EGF-stimulated DNA synthesis (Gladhaug et al., 1988). With respect to possible mechanisms of butyrate action, butyrate is known to inhibit histone deacetylase and may influence gene expression via increased histone acetylation. In hepatocytes, addition of butyrate results in histone acetylation that is maximal after 3-6 h but then returns to normal as the butyrate is metabolized. It is not clear whether this pulse of histone acetylation accounts for butyrate effects on differentiation (Staecker and Pitot, 1988).

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11.6. Homologous cell interactions Excellent evidence suggests that tissue organization makes a major contribution to the control of patterns of gene expression in hepatocytes (Clayton et al., 1985). In addition to the importance of contacts with extracellular matrix components discussed in Section 1 1.1.5, contact of hepatocytes with each other, or with other cell types present in the liver sinusoids or associated with bile ductules or ducts, may also be significant in determining patterns of gene expression in the intact liver and hence in culture as well. The importance of cell-cell interactions has been reviewed in detail by Guguen-Guillouzo (1986) and more briefly by Maher (1988). 11.6.1. ESfects of hepatocyte density on preservation of differentiated function

In deciding on the cell density to use for monolayer culture experiments, a general principle which currently appears appropriate, regardless of substratum or medium used, is that cell survival and preservation of differentiated function are favoured at high cell density while growth-related activities are favoured at lower density. Our experience in line with that of others (e.g. Jauregui et al., 1986) is that, on simple substrata, survival is maximal where hepatocytes form a tightly packed confluent monolayer in which cells have relatively compact morphology. Survival is particularly poor at very low densities where groups of 1-3 cells are widely spaced and extensively flattened on the substratum. The suggestion that high cell density is associated with better preservation of differentiated function is probably an over-simplification since liver-specific albumin secretion may be higher at low density (Guguen-Guillouzo, 1986) and appearance of the foetal liver enzyme y-glutamyltranspeptidase is more rapid at high density (Edwards et al., 1987b).Clearly, on simple substrata, high density alone is not sufficient to achieve more than partial preservation of differentiated function (see Section 1 1.1.3). Nevertheless a vari-

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ety of liver-specific activities are better preserved at high density on collagen films (Nakamura and Ichihara, 1985) or on collagen gels (Ben-Ze’ev et al., 1988). In co-culture, effects of hepatocyte density are apparently similar to those seen in pure monolayers. The good preservation of hepatocytes in spheroid culture (Koide et al., 1990) may be partly the result of homologous cell interactions promoted by close packing. There is little published information on effects of cell density on Matrigel, although on this substratum, the tendency of hepatocytes to aggregate into columns or islands with multiple cell layers maximizes homologous cell contacts. The relatively high plating densities used to date (e.g. Bissell et al., 1987; Schuetz et al., 1988) are probably optimal. Assuming a plating procedure that achieves rates of attachment of 80% or better (see Section 11.2), seeding 1.2-1.5 x lo5 cells/cm2of culture surface should yield confluent monolayers on plastic, collagenor fibronectin-coated dishes or on collagen gels. It may be possible to achieve more tightly packed monolayers by plating up to 2 x lo5 cells/cm2. On Matrigel, plating 1.7 x lo5 cells/cm2 gives maximal density. 11.6.2. Effects of hepatocyte density on growth-related parameters

In general to ensure that a high proportion of hepatocytes in culture enter S-phase in response to EGF and other growth factors it has been necessary to culture cells at relatively low densities. Maximal percentages of cells active in DNA synthesis (up to 80%, depending on media) have been observed after plating about 0.1-0.2 x lo5 cells/cm2.The percentage of active cells decreases progressively with increasing density up to about 0.8 x los cells/cm2. In such near-confluent monolayers, few cells respond to growth factors (e.g. Michalopoulos et al., 1982; Nakamura and Ichihara, 1985; Sawada et al., 1986; Vintermyr and DBskeland, 1989). Lower densities on simple substrata also favour higher levels per cell of total RNA and protein synthesis, amino acid transport and glucose-6-phosphate dehydrogenase induction

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(Nakamura and Ichihara, 1985; Guguen-Guillouzo, 1986).The effect of density on DNA synthesis is similar with a variety of simple substrata including plastic and dishes coated with Type I or Type IV collagens, fibronectin or laminin (e.g. Sawada et al., 1986) and apparently in cells on collagen gels (Ben-Ze’ev et al., 1988), whereas in cells on Matrigel (Ben-Ze’ev et al., 1988) or in most co-culture systems (Guguen-Guillouzo, 1986) little DNA synthesis is observed even at low density. At very low densities (less than 0.1 x los cells/cm2),replication is less clearly related to density. Vintermyr and DBskeland ( I 989) reported a sharp decline in the proportion of cells active in DNA synthesis at densities < 0.2 x 105 cells/cm2, with levels of replication dependent more on total cells per dish than on local density. One explanation for poor growth (and survival) at very low density could be that modification of the culture medium by hepatocytes (i.e. ‘conditioning’)is required for optimal growth. Production of autocrine growth factor(s) is important in neonatal cultures but apparently not in adult cultures (Nakamura et al., 1988). Conditioning of media by high density cultures is probably not significant in improving differentiation (Nakamura and Ichihara, 1985), although one element in inhibition of DNA synthesis at high density may be depletion of arginine in the medium by membrane-associated arginase (Koji and Terayama, 1984). As discussed in Section 11.1.3, the main factor in reciprocal effects of density on growth and differentiation appears to be contact between hepatocytes.

11.7. Heterologous cell interactions 11.7.1. Rationale and characteristics of co-culture systems

During embryonic development of the liver, endodermal tissue differentiates to form the hepatic parenchyma while mesenchyme gives rise to the endothelial cells lining the liver sinusoids. Tissue interac-

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tions between hepatic endoderm and mesenchyme appear to be necessary for differentiation of hepatocytes. When maintained in culture, mouse hepatic endoderm present alone degenerates, but when maintained in close contact with mesenchyme (either hepatic or pulmonary), endodermal cells proliferate and develop the ability to store glycogen and synthesize a-foetoprotein (Houssaint, 1980).The importance of such heterologous cell interactions during development suggests that related interactions might be important for maintaining hepatocyte differentiation in adult tissue (Bissell, 1983; GuguenGuillouzo, 1986) and this has been tested by co-culturing hepatocytes with a second cell type. The usual procedure for co-culture studies involves allowing hepatocytes to attach at near- or sub-confluent density, then adding cells from a suitable epithelial or mesenchymal cell line. In general these proliferate to fill the spaces between groups of hepatocytes within 2 4 - 4 8 h then stop growing, although in one system a liver cell line continued to grow over and under the hepatocytes (Vajta et al., 1986). Thereafter the co-cultures are maintained in defined culture media with added insulin and glucocorticoids. In various co-culture systems hepatocytes form colonies or islands of cells with compact, polygonal shape, granular cytoplasm and 1-2 nuclei. This morphology is preserved for 1-2 months (Guguen-Guillouzo et al., 1983; GuguenGuillouzo, 1986; Goulet et al., 1988) and is associated with at least partial preservation of in vivo-like patterns of transcription, liver specific mRNAs, maintenance of total cytochromes P-450and restoration of functions such as albumin secretion and gap junctional communication (Fraslin et al., 1985; Guguen-Guillouzo, 1986; Goulet et al., 1988). In successful co-culture systems a common feature is the production of extensive reticulin fibres (containing fibronectin, Type I collagen and laminin) that form a ‘cocoon’ around and over the hepatocytes and infiltrate epithelial cell areas. Fibre formation, which reaches a maximum after 7-8 days, may provide important extracellular matrix contacts for the hepatocytes. Synthesis of fibres

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depends on heterologous cell interactions since few fibres are observed in pure cultures of either cell: the interaction is not via tight or gap junctions but may involve membrane components such as cell adhesion molecules (Guguen-Guillouzo et al., 1983; Mesnil et al., 1987; Goulet et al., 1988). While only the outer cells in hepatocyte islands have contact with the second cell type, this is adequate to allow improved survival and albumin synthesis in all hepatocytes in the colony (Guguen-Guillouzo, 1986). Independent of heterologous cell contacts, the density of hepatocytes in co-cultures influences functions such as albumin and total protein synthesis in the same way as in pure culture (Guguen-Guillouzo, 1986). 11.7.2. Choice of cell type for co-culture with hepatocytes

The cell type optimal for use in co-culture with hepatocytes is still controversial. Guguen-Guillouzo and her co-workers (Clement et al., 1984; Guguen-Guillouzo, 1986) have suggested that co-culture requires relatively specific cell interactions. They argue that in vivo, only one cell type has direct contact with hepatocytes, namely the biliary epithelial cells which form the canals of Hering. These canals allow bile transport from bile canaliculi to the ductules and ducts. The rat liver epithelial cells, isolated according to Morel-Chany et al. (1978) and used after a small number of cell passages in culture, or the epithelial cells from an established cell-line (Mesnil et al., 1987),which have been used in co-cultures by the Guguen-Guillouzo laboratory, are generally considered to be derived from these biliary epithelial cells (Grisham, 1983; Guguen-Guillouzo, 1986), although Marceau et al. (1986b) have suggested they could be derived from Glisson’s capsule. More recently, there has been increasing evidence that cells other than those derived from biliary epithelium may be useful for co-culture. Morin and Normand (1986) showed that plating hepatocytes from suckling rats onto primary cultures of liver sinusoidal (mainly endothelial) cells resulted in improved preservation of hepatocytes with high levels of albumin production, although the co-cultures

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deteriorated after 6-7 days. An endothelial cell-line established from the sinusoidal primary cultures allowed longer-term maintenance of suckling, or adult, rat hepatocytes with high albumin secretion and preservation of gap junctions for at least 4 weeks (Morin and Normand, 1986; Goulet et al., 1988). Although co-culture effects of mesenchymal cells require further investigation, findings of Goulet et al. (1988) suggest that some mesenchymal cells including mouse embryonic fibroblasts (NIH-3T3 cells) and rat dermal fibroblasts also have beneficial effects in co-culture. These authors concluded that heterologous cell interactions required for hepatocyte preservation in co-culture are not tissue, organ or species specific. Senoo et al. (1989) also observed improved survival and albumin production by hepatocytes co-cultured with tendon fibroblasts, although the merits of fibroblast co-culture for overall preservation of hepatocytes may depend on the origin of the fibroblasts (Michalopoulos et al., 1979; Goulet et al., 1988). If more extensive studies confirm the value of NIH-3T3 cells, the wide availability of this established cell-line could facilitate greater use of co-culture. At present, co-cultures with early passage rat liver epithelial cells or endothelial cell-lines provide the best-characterized systems (Guguen-Guillouzo, 1986; Goulet et al., 1988). 11.7.3. Plating and culture procedure for co-culture

The benefits of co-culture for preserving hepatocyte differentiation appear to depend on close contact between hepatocytes and the heterologous cell. Thus the procedure for plating the two cell types into dishes is designed to achieve relatively high hepatocyte densities together with extensive heterologous cell contacts. The procedure outlined here is based on those of Guguen-Guillouzo et a1 (1983), Goulet et al. (1988) and Conner et al. (1990). Freshly isolated hepatocytes are plated at O.GO.8 x los cellslcm2 into plastic or collagen-coated dishes in culture medium containing. 10 pg/ml insulin, with or without 5% foetal bovine serum. After 3 4 h for attachment

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of the hepatocytes, the second cell type is added in sufficient numbers to yield confluent dishes after 24-48 h. If rat liver epithelial cells are used (isolated according to Morel-Chany et al. ( 1 978) or as described recently in more detail by Furukawa et al. (1987)), it is apparently important to use early-passage cells, prior to transformation: 0.8 x lo5cells are plated in fresh medium containing insulin, 5-1094 foetal bovine serum and 3.5 pM cortisol hemisuccinate and allowed to grow to fill the dish. Serum-free medium containing insulin 10 pg/ml and glucocorticoid (3.5 pM cortisol hemisuccinate or 1 pM dexamethasone) is then used to maintain cultures. The relative merits of different basic culture media for co-culture have not been reported (but see Section 1 1.4.1). It seems clear that the second cell type in co-cultures with hepatocytes ‘conditions’ the medium, increasing lactate levels for instance (Agius, 1988). If ‘conditioning’ of the medium were important it might be appropriate to use small volumes of medium or infrequent medium changes. However it is not clear that ‘conditioning’ has any major role in preserving hepatocyte differentiation (discussed by Guguen-Guillouzo, 1986; Goulet et al., 1988). Thus volumes of media used per dish (1 m1/35-mm dish or 2-3 ml/60-mm dish) and frequency of medium changes (daily) have in general been similar to those used for pure cultures. In relation to achieving hepatocyte growth, co-culture with rat liver epithelial cells and a number of other cell types suppresses DNA synthesis, regardless of the density at which hepatocytes are plated (Guguen-Guillouzo, 1986; Shimaoka et al., 1987). By contrast, coculture with primary cultures of rat liver non-parenchymal cells was reported to increase markedly the [’Hlthymidine labelling index of EGF- and insulin-treated hepatocytes, with maximal labelling achieved at low hepatocyte density (Shimaoka et al., 1987; Yamamoto et al., 1989). In this system it is not clear whether heterologous cell contacts (Shimaoka et al., 1987) or release of protein factor(s) by the nonparenchymal cells (Yamamoto et al., 1989) are more important for the observed growth stimulation.

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11.8. Selecting appropriate methods for monolayer culture As outlined in Section 11.1, a variety of approaches to hepatocyte culture have been used, with varying success, to preserve differentiated functions or to achieve growth. So far, separate lines of investigation have identified a variety of useful media components, substrata in various configurations and cell-cell interactions but the best ways of combining these elements remain to be fully explored. Various investigators have noted ‘plasticity’ in the contribution of different factors to hepatocyte function in that a particular pattern of gene expression may be achieved by differing combinations of substratum interactions, media and cell-cell interactions. The continuing challenge of studies on hepatocyte culture is to define combinations of these factors which achieve desired function in culture systems that are experimentally convenient and, ideally, do not involve high-cost materials. Against this background, the designation of specific protocols below involves some rather arbitrary choices. For the investigator contemplating the use of monolayer culture it is, of course, important to relate the chosen protocol to the problem under investigation. For many studies on functions that do not undergo rapid change in culture, confluent monolayers of hepatocytes on collagen-coated dishes, using various commercially-availablemedia such as Williams’ Medium E or a Dulbecco’s MEM/F-12 (1:l) mixture, supplemented with insulin (100 nM) and dexamethasone (30-100 nM), may prove adequate. For investigations involving liver-specific functions, initial studies suggest that more complex media, supplemented with insulin and dexamethasone (and possibly glucagon) can allow excellent preservation of some specific functions for at least a few days. These media include Chee’s medium as supplemented in Table 1 1.1 and used by Waxman et al. (1990); the high amino acid medium developed in Jefferson’s laboratory (Table 11.1) and used by Hutson et al. (1987) or in slightly modified form by Lloyd et al. (1987); and the modified Waymouth Medium with insulin, dexamethasone and glucagon, used by Dich et al. (1988).

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Investigations requiring longer-term preservation of differentiated functions and patterns of transcription more like those in vivo apparently require the use of more complex substrata or co-culture. Protocol 11.5 given below is based on the use of the basement membrane-like substratum prepared from EHS tumours, which is available commercially as ‘Matrigel’ (at high cost) or can be prepared according to Protocol 11.4. At present there is limited data on the most appropriate medium for use with Matrigel, since the complex media developed to optimize hepatocyte preservation on collagen-coated dishes have not been tested extensively with Matrigel. The medium used in Protocol 11.5 is related to the modification of Williams’ Medium E developed by Lanford et al. (1989) (see Table 11.1) for longterm maintenance of primate hepatocytes. The medium for Protocol 11.5 is supplemented with a number of the nutrients and hormones which appear to enhance the preservation of hepatocytes as discussed in Section 1 1.4, but some factors included by Lanford et al. (1989) in their medium are omitted. This medium, although yet to be tested widely with Matrigel, resembles that used successfully with EHS tumour-derived substrata by Ben-Ze’ev et al. (1988). Studies on hepatocyte growth have identified a variety of agents which stimulate or support growth. As noted for the preservation of differentiated functions, optimal conditions for growth are not necessarily achieved by combining all the known stimulatory agents. Various combinations of nutrients, hormones and growth factors can be used to achieve at least one round of DNA replication and mitosis in a high proportion of cells. Protocol 11.6 is based on one effective combination currently used in our laboratory. As noted in Section 1 1.4, other basic culture media, high pyruvate concentrations, serumderived factors and oestrogens might also be used in appropriate combinations to achieve high levels of replication and mitosis.

Protocol 11.5 Culture method for studies on dgferentiated function All equipment used in the following Protocol should be sterile and

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appropriate sterile technique must be used in all steps. Manipulations of the cells after isolation should be carried out in a laminar flow cabinet. (i) For preparation of the base culture medium, purchase Williams' Medium E in powdered form (e.g. from ICN Biomedicals or Sigma). High quality water should be used in the preparation of all media. De-ionized water, glass distilled (in a still, cleaned out regularly) or water from a commercial water purification system, which yields conductivities less than 0.08 pS/cm at room temperature should be suitable. Dissolve the powdered medium together with the following substances in 1 1: 2.2 g NaHCO,, 2.38 g HEPES, 17.3 pg Na,SeO, and 50 mg gentamicin. Adjust the solution at room temperature to pH 7.4, make up to final volume and sterilize by filtration through a 0.2-pm membrane filter. The base medium can be stored at 0-4"C for 6 8 weeks. (ii) Prepare the following supplements for the base culture medium as sterile 1000-fold concentrated stock solutions: 100 pM dexamethasone, 100 pM insulin, 1 pM glucagon, 5 pM EGF, 1 mM ethanolamine and transferrin, 5 mg/ml. These solutions are made up as follows: 0.039 mg dexamethasone in 1 ml DMSO (self-sterilizing); 6 mg insulin dissolved in 2 mlO.01 M HCl then diluted to 10 ml with 0.15 M NaCI; 0.035 mg glucagon, 0.30 mg EGF, 0.61 mg ethanolamine and 50 mg transferrin, each dissolved in separate 10-ml vols. of 0.15 M NaCI. Sterilize the stock solutions (except for dexamethasone) by filtration through 0.2-pm membrane filters. Use of low protein-binding filters such as Millipore 'Durapore' filters is desirable since significant amounts of the peptide hormones may be lost during filtration as a result of binding to standard cellulose nitrate or cellulose acetate filters. Alternatively, a number of the agents above are available as sterile solutions or as sterile, lyophilized preparations which can be re-constituted using sterile 0.15 M NaCl. The aqueous stock solutions may be stored at -20°C or at -70°C for several weeks. Repeated freezing and thawing should be avoided. In our laboratory, stock solutions of dexamethasone are prepared freshly each week and stored at -20°C.

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(iii) Prepare a sterile 100-fold concentrated stock solution of linoleic acid/albumin which is also used as a supplement to the base culture medium. Dissolve 5 g of fatty acid-free BSA in 100 ml 0.15 M NaCl, add 50 mg linoleic acid, stir to allow the fatty acid to bind to the albumin, adjust to pH 7.4, then sterilize by filtration through a 0.2-pm membrane filter. Store 10-ml aliquots at -20°C. (iv) Prepare Matrigel as described in Protocol 11.4 or purchase it from ICN Biomedicals. (v) The day before cell isolation, use the Matrigel preparation to form gels in 60-mm Lux tissue culture dishes as described in Protocol 11.4 (xiv). Keep the dishes overnight in a humidified incubator at 37°C. To fully utilize a normal cell preparation yielding about 3 x lox hepatocytes, 4 0 - 6 0 dishes will be required. (vi) Isolate hepatocytes as described in Protocol 1 1.1. (vii) While cell isolation is in progress, supplement about 300 ml of base culture medium with insulin (100 nM) and equilibriate at 37°C. (viii) After the final wash of hepatocytes, resuspend the cell pellet in 40 ml base culture mediudinsulin using a wide-bore 10-ml or 25-ml pipette to achieve a uniform suspension. From this point, the hepatocytes should be plated into dishes as soon as possible so that cells do not develop ‘blebs’ as a result of becoming anaerobic. (ix) Transfer all or part of the cell suspension (depending on the numbers required for the experiment) to a 500-ml De Long or Erlenmeyer flask and add additional warm medium (with insulin) so that the cell density is about 1.5 x 10bcells/ml. For a full preparation (3 x lo8 cells), the total volume would be about 200 ml. Swirl the flask to form a uniform suspension and continue to swirl the flask at 20-30-s intervals so that the cells are kept in suspension and do not become anaerobic. (x) Use a haemocytometer to count cell numbers in the suspension (Section 2.6.1) and, based on the count obtained, dilute with additional warm medium (with insulin) to give a final cell density of 1.3 X lo6 hepatocytes/ml. (xi) Using a wide-bore 10-ml or 25-ml pipette, add 3 ml cell suspen-

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sion per Matrigel-coated 60-mm dish (actual surface area, 23 cm’). This should give a plating density of 1.7 x lo5cells/cm’. To achieve consistent cell density in all dishes it is most important to swirl the flask containing the cell suspension constantly so that a uniform suspension is maintained. Without continual shaking, the hepdtocytes settle rapidly. (xii) To achieve a relatively uniform cell density on the Matrigel in each dish, the manner of handling culture dishes immediately before they are placed in the incubator is important. Usually cell suspension is added to dishes placed on an incubator tray. When cells have been added to all dishes on the tray, carry the tray to the open incubator and immediately before inserting the tray, hold it horizontal and gently shake four or five times ‘north-south’ and then three or four times ‘east-west’(a circular motion tends to concentrate the cells in the centre of the dish). (xiii) Incubate at 37°C in a humidified atmosphere containing 5% CO, in air. (xiv) The cultures should be left undisturbed for at least 1 h. A period of 2 h for attachment is recommended, although periods from 1-4 h may be used before the medium is changed to remove non-attached cells. (xv) For the medium change after attachment (and subsequent changes), prepare complete medium by supplementing the base medium from the stock solutions prepared in (ii) and (iii). Use 0.1 ml of the 1000-fold concentrated stocks/100 mi medium and 1.0 ml of the 100-fold concentrated stock solution/100 ml medium. The complete medium should then contain Williams’ Medium E with 10 mM HEPES, 100 nM Na,SeO,, 50 pg/ml gentamicin, 100 nM dexamethasone, 100 nM insulin, 1 nM glucagon, 5 nM EGF, 1 pM ethanolamine, 5 pg/ml transferrin and 5 pg/ml linoleic acid with 0.5 mg/ml fatty acid-free BSA. Equilibrate the medium at 37°C. (xvi) After the 2 h attachment period, remove all medium plus nonattached cells by aspiration. The old medium should be removed as completely as possible without the aspirator tip touching the Matrigel.

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For aspirating the medium, a 14-15G, 60-mm long stainless steel needle is convenient, since this can be sterilized by flaming in a bunsen flame at regular intervals during the medium change. The needle is connected via silastic tubing to a side arm flask that acts as a reservoir for aspirated medium and this is joined to a vacuum line via a second flask to act as a trap for any overflow of medium. (xvii) To streamline medium changes, the old medium may be aspirated from multiple dishes on a tray, then fresh complete medium pipetted into the dishes. However, monolayers should not be left for more than 3 - 4 min without medium. Use 2.5 mi mediud60-mm dish. (xviii) Continue incubation of the monolayers at 37°C in a humidified atmosphere containing 5% CO, in air. (xix) At 24-h intervals, change the medium by aspirating the old medium and adding 2.5 ml fresh complete medium per dish. (xx) Immediately after attachment, the hepatocytes on Matrigel remain rounded and are present as single cells or small groups which might occupy 70-80% of the surface area of the gel. With time, the hepatocytes progressively aggregate into larger, multi-layer groups, usually in cords which form a network on the Matrigel. Although some experimental studies might begin immediately after attachment, it is generally desirable to leave monolayers for a minimum of 24 h to allow time for the cells to repair any damage suffered during isolation and to establish contacts with the extracellular matrix and other hepatocytes. Studies in our laboratory on enzyme induction have usually been initiated by adding test agents with the medium change after 48 or 72 h in culture.

Protocol 1 1.6 Culture method for studies on growth This Protocol, which outlines one set of conditions for achieving substantial DNA synthesis and mitosis in monolayer culture, shares many steps with Protocol 11.5. Those steps which differ are emphasized below.

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(i) Prepare the base culture medium and concentrated stock solutions essentially as described in Protocol 11.5 (i) to (iii). A glucagon solution will not be required and a 30 pM stock solution of dexamethasone should be prepared by diluting 0.3 ml of the 100 pM dexamethasone solution with 0.7 ml of DMSO. (ii) One or more days before cell isolation, coat tissue culture dishes of the desired size with Type I collagen as described in Protocol 11.2. The procedure below is given for 60-mm diameter dishes (actual surface area, 23 cmz). A full hepatocyte isolation will yield sufficient cells for more than 300 lower-density cultures so, in general, the number of dishes used will be determined by the experimental design rather than the availability of cells. (iii) Isolate hepatocytes and resuspend the washed cell pellet in 40 ml warm base culture medium plus insulin (100 nM) as described in Protocol 11.5 (vi) to (viii). (iv) From a cell preparation yielding about 3 x los hepatocytes, 7-10 ml of the resulting suspension would contain sufficient cells to establish 100 low-density cultures. Using a wide-bore 10-ml pipette, transfer an appropriate volume of suspension (e.g. 8 ml for 100 dishes) to a 500-ml De Long or Erlenmeyer flask and add warm base mediudinsulin to give a cell density of about 1.8-1.9 x lo5 cells per ml (e.g. 320 ml to dilute 8 ml of suspension). Swirl the flask frequently to keep the cells in a uniform suspension. (v) Use a haemocytometer (Section 2.6.1) to count cell numbers in this suspension and then adjust the density to 1.6 x lo5 hepatocytes/ml with base mediudinsulin. (vi) Use a wide-bore pipette to add 3 ml/60-mm collagen-coated dish and follow the measures outlined in Protocol 11.5 (xi) and (xii) to ensure uniform distribution of the cells. (vii) Allow 2 h at 37°C in an atmosphere of 50/0 CO, in air humidity) for cell attachment. (xiii) After attachment, change the medium as in Protocol 11.5 (xv) to (xvii), except that the volume should be 2 ml medium per dish and the complete medium for growth experiments should contain Williams’

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Medium E with 10 mM HEPES, 100 nM Na,SeO,, 50 pg/ml gentamicin, 30 nM dexamethasone, 100 nM insulin, 5 nM EGF, 1 pM ethanolamine, 5 pg/ml transferrin and 5 pg/ml linoleic acid with 0.5 mg/ml fatty acid-free BSA. (ix) Norepinephrine can be used to provide a further growth stimulus. Immediately before use, prepare a stock solution of 3 mM norepinephrine by dissolving 6.2 mg norepinephrine HCl in 10 mlO.15 M NaCl, filter to sterilize and add 1 ml of this stock solution per 100 ml culture medium used for the medium change after cell attachment. (x) Incubate as above and change the medium at 24-h intervals. (xi) This Protocol illustrates culture conditions which should result in the majority of hepatocytes entering S-phase after about 40 h in culture, with subsequent mitosis in many cells.

11.9. Variations on monolayer culture In this Section some variations on monolayer culture, in general designed to fit a particular set of experimental priorities, are briefly described. The references given should be consulted for detailed methodology. 11.9.1. Monolayer culture on microcarriers

Conventional monolayer culture in dishes has some possible disadvantages for metabolic studies because it is a static unstirred system, which may develop gradients of substrate concentration, 0, or pH. Also, the number of cells per unit volume of medium is limited by the surface area of the dish. Culture of hepatocytes on microcarriers has been developed to provide a stirred system in which greater surface area per unit medium volume is possible. Microcarrier cultures combine some of the ease of experimental manipulation and sampling, characteristic of suspension cultures, with the longer cell survival possible in monolayers. They may also be useful for incorporation into perifusion systems.

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Hepatocytes will attach to Cytodex 1 microcarriers (positivelycharged N,N-diethylaminoethyl-substituteddextran beads, from Pharmacia) in the presence of serum or to Cytodex 3 (dextran beads with a covalently-linked Type I collagen surface layer) in the absence of serum. Agius et al. (1985) and Athari et al. (1988) have described slightly different procedures for achieving attachment and for subsequent incubation. Attachment requires a compromise between the relatively static conditions necessary for cells to attach to a culture surface and agitation to promote both even attachment over the microcarriers and adequate oxygenation. Athari et al. (1988) incubated 0.7 X lo8 cells and 0.3 g (dry weight) of pre-swollen, sterile Cytodex 3 microcarriers in 100 ml Medium 199 supplemented with 4% foetal bovine serum, 1 nM insulin, 0.1 p M dexamethasone and 0.2% BSA in a siliconized 1-1 flask under a carbogen atmosphere, at 37°C. The flask was gently stirred by hand every 10 min for a total of 2.5 h. This procedure resulted in greater than 80% attachment. After a medium change, the microcarriers were maintained in serum-free medium ( 160 ml) using a Wheaton Magnaflex 1 1 spinner vessel, stirred with a suspended magnet at 40-60 rpm. Agius et al. (1985) reported that, depending on attachment conditions, up to 8 h was required for hepatocytes to establish firm attachment to beads, sufficient to withstand washing or shaking (1-2 m1/25-ml Erlenmeyer flask) in a reciprocating water bath. Microcarrier cultures have been incubated for up to 48 h and exhibit metabolic activities and hormone responses comparable to those of dish cultures (Athari et al., 1988) and rates of protein synthesis substantially greater than comparable suspensions of hepatocytes alone (Agius et al., 1985). 11.9.2. Culture on filters or membranes

Savage and Bonney ( 1978) cultured hepatocytes on MF-Millipore filters, which are formed from mixed esters of cellulose acetate and cellulose nitrate. Sterilized stainless steel rings were placed over 0.45-pm pore filters in bacteriological plastic petri dishes to hold them stationary

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during attachment of the hepatocytes. Cells were plated on to dry filters in the presence of insulin and 15%) foetal bovine serum and after attachment for 2 4 h, the filters were allowed to float in fresh, serumfree me‘dium. Monolayers were reported to survive for 6-8 days and to be better preserved than on plastic dishes. The technique has possible advantages in allowing rapid transfer of monolayers from one culture environment to another, in providing an attachment surface which is easily manipulated for techniques such as electron microscopy or for co-culture experiments concerned with exchange of soluble factors between cell populations. Thin gas-permeable membranes have also been used for hepatocyte culture. Petzinger et al. (1988) plated hepatocytes on collagen-coated ‘petriperm’ dishes (from Heraeus, Hanau, FRG). These dishes contain a thin (1 pm), highly transparent ‘biofilm’ on which the cells attach and survive for at least 4 days. The thin, transparent substratum was considered to be desirable for studies involving micropuncture of bile canaliculi, or electron microscopy. 11.9.3. Perijiused monoluyer culture

In conventional culture, the most common practice is to use daily medium changes. Because of the active metabolism of numerous substrates, drugs etc. by hepatocytes, interpretation of experiments may be complicated by progressive changes in metabolite levels between medium changes. In a number of situations it is necessary to add high non-physiological initial concentrations of test agents to achieve effective concentrations for a reasonable proportion of the period between medium changes. Gebhardt and Mecke (1979a) have described a perifusion system to overcome these problems. Hepatocytes were allowed to attach to collagen-coated glass microscope slides in a lucite chamber fitted with multiple inlet and outlet ports. After attachment under static conditions, medium equilibriated at 37°C with 5% CO, in air was pumped through the chamber at the rate of about 0.5 ml/h per 106 cells. The design of the system allowed a series of

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parallel perifusions with different media and incorporation of an infusion pump for studies with hormones. Under perifused conditions, flattening and association of hepatocytes into trabeculae occurred more rapidly than in the conventional culture system. Perifusion with serumcontaining medium (but not serum-free medium) could be continued for up to 8 days with better cell survival than with daily medium changes and substantially better preservation of some liver-specific transaminases and urea cycle enzymes (Gebhardt and Mecke, 1979a). Hormonal induction of some urea cycle enzymes by glucagon (in combination with dexamethasone) could be observed in perifused but not conventional cultures (Gebhardt and Mecke, 1979b). 11.9.4. Culture of periportal or perivenous hepatocytes

The liver parenchyma is considered to be organized into functional units, termed acini, related to the microvascular architecture of the liver. Various metabolic processes are known to be heterogenously distributed in these acini. For instance, periportal cells surrounding the parenchymal branches of the portal vein are more active in gluconeogenesis, amino acid catabolism and urea synthesis and are the site of earlier DNA synthesis during liver regeneration, whereas pericentral cells around the hepatic (central) venules appear to be more active in glycolysis and in oxidation and conjugation of various xenobiotics (e.g. Gumucio and Chianale, 1988). The introduction of digitonin-collagenase perfusion procedures (see Section 12.8), in which either periportal or perivenous cells are first destroyed with digitonin and then the remaining intact cells isolated by normal collagenase perfusion, allows populations enriched in periportal or perivenous cells to be prepared and studied in culture. The two populations attach equally well to collagen-coated dishes and exhibit similar survival, with both populations undergoing changes in differentiated function with time in culture, such as loss of alanine aminotransferase and cytochrome P-450-dependent drug metabolizing activity and elevation of y-glutamyltranspeptidase (Suolinna et al.,

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1989), similar to changes reported for mixed cultures. Various differences in metabolic activity between periportal and perivenous populations (urea synthesis, gluconeogenesis, drug metabolism) are retained for several days in culture (Poso et al., 1986; Quistorff et al., 1986; Suolinna et al., 1989) and periportal cells are also more active in DNA synthesis. Lower DNA synthetic activity in the perivenous cultures is due to very low activity in a subpopulation of these cells which, in vivo, is located in 1-3 cell layers immediately around the terminal hepatic venules (Gebhardt, 1988).This phenotypically-distinct population (about 8% of all hepatocytes) is distinguished by high glutamine synthetase activity that is preserved when cells are placed in culture (Gebhardt and Mecke, 1983).

11.10. Culture of hepatocytes from foetal, neonatal and suckling rats 11.10.I. Foetal liver

In rats the initial time of liver formation is around 10-1 1 days gestation. At days 11-12 there is an apparently uniform population of liver epithelial cells, smaller than mature hepatocytes, which may represent a population of progenitor cells capable of differentiation into hepatocytes or biliary epithelial cells. Significant numbers of these bipotential cells are still present at day 15 but their numbers decline to very low levels at birth. From day 15 the predominant epithelial cell type is the immature hepatocyte (see Germain et al., 1988 for discussion and references). Marceau and his co-workers have microscopically dissected livers from 12-day foetuses and disaggregated fragments using a mixture of collagenase, dispase and hyaluronidase. Epithelial cells were separated at higher centrifugal speeds (200 x g ) than normally employed in hepatocyte isolation and plated onto fibronectin-coated coverslips in the presence of insulin and dexamethasone. When maintained with

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I SO LATED H EPATOCYTES

serum, insulin, dexamethasone and DMSO these cells underwent phenotypic changes typical of differentiation to hepatocytes, while with further addition of sodium butyrate, the cells acquired bile duct epithelial cell markers (Germain et al., 1988). In related experiments with ‘hepatoblasts’ from 13-day mouse liver buds, cultures contained a variety of cell types but 10-20‘% of the cells in medium containing serum, insulin and cortisone underwent differentiation to hepatocytes in a manner similar to liver cells in vivo (Bennett et al., 1987). Methods appropriate for isolation of cells from day 11-14 foetal livers, as well as the characteristics of the resultant cells in culture, are likely to differ somewhat from those for cells from livers of foetuses at 15 days or later, where most epithelial cells are committed to one differentiation pathway and the predominant cell type is the immature hepatocyte. Most studies on foetal hepatocytes have involved these later stages of foetal liver development. Methods for cell isolation were discussed in Section 3.5 and aspects of 15-20 day foetal hepatocyte culture have been reviewed by Guguen-Guillouzo and Guillouzo ( 1983) and Yeoh (1986). Cell isolates, particularly from earlier stages of gestation, contain many erythroid cells but their presence in cultures can be minimized by allowing relatively short times of 2-3 h for cell attachment (Yeoh, 1986). In the presence of 10%)serum, Hoffmann et al. (1989) found about 70%) attachment to plastic or laminin and 85-90% attachment to films of fibronectin or collagens Type I or IV. Although isolated as single cells, most foetal hepatocytes reaggregate into clumps during attachment. Many studies with foetal hepatocytes have involved incubation of cells on plastic or collagen-coated dishes in media with 10%)foetal bovine serum. Under these conditions overgrowth with fibroblasts is a major problem, particularly where hepatocytes are plated at lower densities (Yeoh, 1986). Overgrowth can be minimized by using a selective medium in which ornithilie is substituted for arginine (Leffert and Paul, 1972) and/or by the inclusion of glucocorticoids in the medium (e.g. Murison, 1976). In serum-containing media, epithelial cells with relatively clear, non-granular cytoplasms grow out at the edges of

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hepatocyte colonies after 3 - 4 days and it is these cells which proliferate most readily at later culture times (Leffert and Paul, 1972; Yeoh, 1986). These cells exhibit some hepatocyte functions such as albumin synthesis. However, it is still unclear whether they represent the predominant immature hepatocytes undergoing de-differentiation or result from preferential growth of a smaller subpopulation of (progenitor?) cells. Such uncertainty about the origin of cells applies to some detailed studies on foetal cell growth after about 2 weeks in culture (e.g. Leffert, 1974). In serum-containing media, foetal cells exhibit phenotypes dependent on the developmental stage at which they were isolated. Existing differentiated functions such as synthesis of albumin or a-foetoprotein are preserved for 3 - 4 days, but decline over a week in culture. Cells isolated at 15-19 foetal days undergo further changes in differentiation in culture allowing developmental events to be studied in vitro (Guguen-Guillouzo and Guillouzo, 1983; Yeoh, 1986; Shelly et al., 1989). Recently, Hoffmann et al. (1989) have described serum-free media enriched with various nutrients plus insulin and EGF which maintain the levels of mRNAs for albumin and a-foetoprotein at relatively constant levels for at least 10 days. Such serum-free conditions, which can also support growth (see below), apparently provide a good basis for studying differentiation under well-defined conditions. Either in serum-containing media (Leffert and Paul, 1973; Yeoh, 1986) or in defined media (Hoffmann et al., 1989), foetal cells proliferate for some days in culture. Depending on the conditions, they may undergo 2-3 population doublings. In the absence of serum, Dulbecco’s MEM did not support any growth of foetal hepatocytes (Hoffmann et al., 1989) while Williams’ Medium E did allow at least one population doubling in the presence of EGF (Henderson et al., 1989). This difference might reflect a requirement for proline as noted for adult cells (Nakamura and Ichihara, 1985). As is the case for neonatal cells, some growth of foetal hepatocytes appears to occur in the absence of any hormone in prolinecontaining, enriched media. Such media support further growth

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stimulation by EGF, a-TCF or basic fibroblast growth factor, acting as complete mitogens. Insulin or IGF-I have smaller stimulatory effects (Hoffmann et al., 1989; Hoffmann and Paul, 1990). Although foetal mouse hepatocytes were reported to grow better on laminin than on other substrata (Hirata et al., 19831, growth of rat cells was not enhanced by substrata other than plastic. Proliferation was greatest with an initial plating density of about 0.7 x 104/cm2although cells in higher density areas of monolayers could be stimulated to synthesize DNA (Hoffmann et al., 1989). In contrast to findings for adult hepatocytes, growth and aspects of normal differentiation in foetal cells seem to be preserved under similar conditions. 11.10.2. Neonatal hepatocytes Although enzymic digestion of minced tissue has been used to prepare hepatocytes from 1-7-day-old rats, Deschenes et al. (1980) and later Nakamura et al. (1987) adapted the two-stage collagenase perfusion procedure for use with the 0.25-g livers of neonates. Although technically difficult, this approach gives higher yields (20-180 X lo6 cells/g liver) than earlier methods, with lower levels of contamination by non-hepatocytes (5-1 5%) and a high proportion of cells that exclude trypan blue (90-95%). The resulting cells attach with 90--100O/u efficiency to fibronectin-coated dishes within 1 h in the absence of serum or hormones (Deschenes et al., 1980) or to collagen-coated plastic in the presence of insulin and dexamethasone within 1-2 h (Nakamura et al., 1987). To achieve nearly confluent monolayers, the neonatal cells are plated at densities of up to 4 x lo5 cells/cm2. Neonatal hepatocytes when isolated from 0-7-day-old rats express some liver differentiated functions such as albumin synthesis and hormone-inducible tyrosine aminotransferase but lack others such as tryptophan oxygenase or glucokinase which appear progressively in rat hepatocytes in the 2-3 weeks following birth (Nakamura et al., 1987). In culture, existing functions appear to be relatively well preserved for up to 10 days (Acosta et al., 1978). The process of terminal

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differentiation can be modulated, notably by contact of immature cells with adult hepatocytes (Nakamura et al., 1987). More attention has so far been given to growth of neonatal cells than to differentiated functions. Prior to weaning, nearly all hepatocytes are diploid in contrast to adult hepatocytes which are predominantly of higher ploidy. Proliferation is thought to occur more readily in diploid hepatocytes (Baribault et al., 1985 and references therein). Like cells from foetal liver, hepatocytes isolated within a few days of birth are capable of active DNA replication and mitosis. Proliferative activity declines progressively in cells from older rats. When cells are isolated from rats more than 12 days old, minimal proliferation is observed in basic culture media with or without serum (Guguen-Guillouzo et al., 1980; Nakamura et al., 1988). The growth control of neonatal (and pres1.imably late foetal) hepatocytes apparently involves clear differences from control in adult cells. Cells isolated within a few days of birth are clearly capable of autonomous growth. Nakamura et al. (1988) reported that after attachment in the presence of 10% serum, insulin and dexamethasone, neonatal cells maintained in Williams’ Medium E alone and without medium changes, nearly all underwent at least one round of DNA synthesis and apparently divided to form confluent monolayers. Since the major peak of DNA replication was delayed until 60 h in culture, growth did not seem to result from commitment of cells to progression through the cell cycle prior to isolation. Growth was considered to involve autocrine control since neonatal (but not adult) hepatocytes produced an unidentified growth factor that stimulated growth of neonatal hepatocytes or 3T3 cells but not of adult hepatocytes. Autonomous growth declined as cells acquired markers of terminal differentiation (Nakamura et al., 1988). Studies with mixed cultures of neonatal hepatocytes and stromal cells suggest that a subpopulation of hepatocytes may continue to grow beyond 4 days in culture in the presence of serum and that addition of EGF and a variety of other factors can stimulate growth of additional cells (see Armato et al., 1985, for discussion).

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Growth in neonatal cells appears to be much less density-dependent than in adult hepatocyte cultures, with DNA replication occurring in dense areas of monolayers (Armato et al., 1978; Nakamura et al., 1987). This difference may be related to the ability of mature hepatocytes, but not other cell types, to prevent growth and cause premature differentiation in neonatal cells (Nakamura et al., 1987). 11.10.3. Hepatocytes from suckling rats

Hepatocytes isolated from rats aged about 14 days seem to represent an intermediate stage between neonatal and adult cells. The 14-day cells have developed most of the differentiated functions of adult liver but continue to synthesize a-foetoprotein. Only about 10% of the cells appear to retain the capacity for growth in the absence of added growth factors. However, the cells are all diploid and undergo DNA synthesis and mitosis more readily than adult cells when stimulated by combinations of growth factors (EGF plus insulin, glucagon and/or triiodothyronine) in proline-containing media (Baribault et al., 1985; Baribault and Marceau, 1986; Handa et al., 1986). Distinct effects of dexamethasone and DMSO on growth and differentiation have been described (Baribault and Marceau, 1986).

11.11. Culture of hepatocytes from other species l l . I I . I . Adult hepatocytes from humans or other primates

As with cell isolation from adult rat liver, isolation methods for primate liver involving the mincing and subsequent enzymic digestion of biopsies have in general given low yields and resulted in cultures with short survival times (e.g. Tokiwa et al., 1986). High-yield isolations have employed a two-stage collagenase perfusion procedure. GuguenGuillouzo et al. (1982) utilized whole livers from kidney donors and perfused 5-10% of the organ via a cannula inserted in the anterior

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branch of the left portal vein. An alternative approach for liver biopsies, described by Reese and Byard (1981) and Strom et al. (1982), involves inserting multiple catheters into vascular orifices on the cut surface of a biopsy, otherwise surrounded by liver capsule (see Section 3.4.6 for details of isolation). Using the latter approach with small biopsies for the preparation of human cells for culture, Strom et al. (1982) and Gomez-Lechon et al. (1990) reported yields of between 10 X lo6 and 40 x lo6 cells/g wet liver, with 70-95% cells excluding trypan blue. Attachment efficiences to collagen- or fibronectin-coated dishes were about 70%. To establish confluent cultures, 8-1 1 X 104 cells/cm2are plated in media containing insulin and 2-10% foetal or newborn calf serum (Strom et al., 1982; Ballet et al., 1984; Guillouzo et al., 1985; Gomez-Lechon et al., 1990). Similar plating conditions are employed for non-human primate hepatocytes (Jacob et al., 1989). Lower rates of attachment were obtained on uncoated plastic dishes (Gomez-Lechon et al., 1990; Butterworth et al., 1989). In general attachment periods of about 4 h have been used, although Butterworth et al. (1989) reported that human hepatocytes from older donors required longer times for attachment. When maintained on collagen or fibronectin in media containing insulin with or without glucocorticoid, human hepatocytes survive in culture for 1-2 weeks (Strom et al., 1982; Gomez-Lechon et al., 1990). Preservation of cells is improved by the addition of insulin and glucocorticoid (Butterworth et al., 1989) and by use of high cell density (Michalopoulos et a]., 1986). In general, under these culture conditions differentiated functions of human hepatocytes change with time in parallel with many of the changes observed in rat hepatocytes, although some changes such as a decline in total cytochrome P-450 and associated drug metabolizing activities or in albumin synthesis occur more slowly (Strom et al., 1982; Ballet et al., 1984; Guillouzo et al., 1985; Grant et al., 1987; Gomez-Lechon et al., 1990). Lanford and his co-workers have recently developed a hormone-supplemented medium, based on Williams’ Medium E but fortified with numerous hormones and other factors, that allows survival of baboon or chim-

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panzee hepatocytes for 3 months on collagen-coated dishes, with excellent preservation of synthesis of apolipoproteins and other plasma proteins. The chimpanzee cultures also support de novo replication of hepatitis B virus (Lanford et al., 1989; Jacob et al., 1989). Guguen-Guillouzo and her collaborators have shown markedly improved preservation of differentiated functions when human hepatocytes are co-cultured with rat liver epithelial cells (Guillouzo et al., 1985; Clement et al., 1984). These workers further reported that addition of 1.5% DMSO to pure cultures slightly increased survival and improved the functional stability of cultures; while in co-cultures, DMSO improved the extent to which cells could be infected with hepatitis B virus (Gripon et al., 1988). DMSO, however, is not uniformly effective in preserving specific functions in human hepatocytes (Vandenberghe et al., 1988). Conditions for observing extensive DNA replication in primate hepatocytes have not been widely studied, although Lanford et al. ( 1 989) reported that extensive proliferation of most baboon hepatocytes in culture occurred in their complete hormone-rich medium, whereas a less-enriched medium supported proliferation only in a few cell foci which emerged after 2 weeks in culture. Chan et al. (1989) recently reported that while human hepatocytes fail to exhibit increased DNA synthesis in response to EGF under various conditions where rat hepatocytes respond, a period of exposure of human cells to DMSO may allow them to acquire mitotic competence, since stimulation of DNA synthesis by EGF f norepinephrine was observed after removing DMSO from cultures incubated for 7 days with EGF plus DMSO. 1I.11.2. Human foetal hepatocytes

Foetal hepatocytes have been prepared from human foetuses at 9-23 weeks gestation by digestion of minced tissue with collagenase (Guguen-Guillouzo et al., 1984; Sells et al., 1985), a mixture of collagenase, hyaluronidase and trypsin (Salas-Prato et al., 1988)or dispase

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(Tokiwa et al., 1987). For cells prepared with collagenase, yields of 6 5 x lo6 hepatocytedg liver, with 70-90% intact cells (by trypan blue exclusion) were reported. In the presence of 10% foetal bovine serum, 20% of hepatocytes (but few other cell) types attached to Primaria dishes. There was little attachment in the absence of serum (Sells et al., 1985). The preparations of Guguen-Guillouzo et al. (1984) attached to plastic within 3 h with up to 80%)efficiency when plated with serum at 12 x 104 cells/cm*. Serum is probably not essential for attachment of human foetal hepatocytes to collagen or fibronectin (Salas-Prato et al., 1988). In the presence of serum (Sells et al., 1985) or in complete medium without glucocorticoid (Tokiwa et al., 1987), fibroblastic cells rapidly overgrow foetal hepatocytes whereas cultures with epithelial morphology are preserved in serum-free media supplemented with glucocorticoid f other hormones. In conventional cultures, cells retained normal morphology for about 4-6 days but then, in most studies so far, progressively detached from various substrata over periods of 2 4 weeks, depending on media and substratum. The use of Type I collagen gels and/or hydrocortisone-containing media improved the survival of cells (Guguen-Guillouzo et al., 1984; Tokiwa et al., 1987).The a-foetoprotein synthesis characteristic of foetal cells declined in the first few days of culture in a manner almost independent of medium or substratum, while production of albumin, more typical of adult hepatocytes, increased to a maximum around IO- 14 days. Albumin synthesis was dependent on hormone-supplements and possibly substratum (Salas-Prato et al., 1988; Tokiwa et al., 1987). Guguen-Guillouzo et al. (1 984) reported that co-culture with rat liver epithelial cells preserved hepatocyte morphology for several weeks and maintained albumin synthesis better than in pure cultures with hydrocortisone as the only hormone supplement. Human foetal hepatocytes, if plated at cell densities around 1-3 x lo4 cells/cm*, may proliferate at least during the initial days in culture so as to form a confluent monolayer (in medium supplemented with EGF, glucagon, hydrocortisone, ethanolamine, selenous acid and

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0.5% dialysed foetal bovine serum) (Salas-Prato et al., 1988).Paracrine action of IGF-I may be involved in controlling growth of the foetal cells (Strain et al., 1987). Sells et al. (1985) established long-term proliferating cultures of hepatocyte-like albumin-producing cells in a serum-free medium with hormone and nutrient supplements related to the hormone-defined medium of Reid and her co-workers (see Table 11.1). 11.11.3. Hepatocytes from mice, rabbits and other species

In general, methods employed for isolation and culture of hepatocytes from species other than rats and humans have been based on experience with rat hepatocyte cultures and species-specific requirements have not been investigated in detail. Because the genetics of mice have been characterized in much greater detail than is the case for other species, studies with murine hepatocytes may be of particular interest in areas such as carcinogenesis. Methods for isolation of mouse cells have been discussed in Section 3.4.1. Mouse hepatocytes apparently survive in culture for longer periods than rat cells but, like rat hepatocytes, lose various differentiated functions when maintained with basic culture media, with or without serum (Klaunig et al., 1981b; Maslansky and Williams, 1982; Clayton and Darnell, 1983). In a recent study, Sawada et al. (1988) fractionated cells on Percoll prior to culture. Their findings suggest that hepatocytes from adult mice may be useful for studies on growth, since nearly all the mouse cells replicated two or three times by the 7th day in culture under defined conditions where only low levels of rat hepatocyte mitosis were observed. Murine hepatocyte growth occurred in a Dulbecco’s MEM/F-12 mixture supplemented with EGF plus insulin, transferrin and selenium and gave rise to ‘immature’ hepatocytes that synthesized a-foetoprotein (Sawada et al., 1988). Rabbit hepatocytes have been reported to preserve total cytochrome P-450 and rates of bile acid synthesis better in culture than rat cells and methods for rabbit hepatocyte culture for studies on these

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parameters have been described (Maslansky and Williams, 1982; Daujat et al., 1987; Whiting et al., 1989). Methods have also been described for culture of hepatocytes from hamsters (Maslansky and Williams, 1982), pigs (Kwekkeboom et al., 1988), fish (Klaunig, 1984) and ducks (Fourel et al., 1989).

11.12. Some commonly used experimental methods in studies with hepatocyte monolayers Any attempt to discuss specific research applications of hepatocyte monolayer culture is beyond the scope of this book. This Section briefly considers some basic procedures which might be required in a range of culture applications. In many cases, methods appropriate for use with hepatocytes in suspension are also applicable, with minor modifications, to monolayers. For simplicity, all the procedures given below employ quantities appropriate for 60-mm culture dishes with a surface area of about 23 cm2.Quantities for other culture vessels should vary roughly in proportion to surface area. The solution widely used for washing cells is Dulbecco’s PBS, which contains per litre: 8 g NaCl, 0.2 g KCl, 0.2 g KH2P0,, 1.15 g Na,HPO,, 0.133 g CaC1,.2H20 and 0.1 g MgC1,.6H20, adjusted to pH 7.4. 11.12.1. Assessment of hepatocyte integrity in monolayers

In monolayers, most damaged cells eventually detach from the substratum or are easily seen as rounded cells still attached to the substratum or to other flattened hepatocytes. Thus assessment of the proportion of intact cells in monolayers has been less of a concern than in hepatocyte suspensions. Nevertheless in some studies, such as those on chemical cytotoxicity in cultured hepatocytes, a relatively simple measure of the numbers of damaged or intact cells is desirable. Methods have been discussed by Jauregui et al. (1981) and Chao et al. (1988).

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At least at early times in culture, trypan blue can be used to detect cells which have lost plasma membrane integrity but remain attached to the culture surface. Monolayers are washed twice with medium or PBS to remove unattached cells and then exposed for 5 min to trypan blue solution. Williams et al. (1977) used 5 ml serum-free medium plus 0.2 ml 0.4% trypan blue. This low final trypan blue concentration (0.016%) was reported to give staining similar to higher concentrations (0.2-0.3%) usually employed with isolated hepatocytes (Section 2.5.1). Jauregui et al. (1981) exposed monolayers to 0.2% trypan blue in saline. The trypan blue solution is removed prior to counting stained and dye-excluding cells in the dishes. An inverted phase-contrast microscope is used for counting, as described by Williams et al. (1977). After a few days in culture the boundaries between hepatocytes in monolayers may be poorly defined and counting is impractical. Since it is difficult to achieve quantitative release of hepatocytes as intact single cells from substrata, counts of trypan blue-stained cells cannot be used after 2 or 3 days in culture. Alternative approaches used by Jauregui et al. (1981; 1986) and Chao et al. (1988), involve measurements of LDH in culture media to assess the extent of4eakage due to cell damage and assays of intracellular LDH (releped from washed monolayers using the detergent, Triton X-100) to provide a measure of numbers of intact cells attached to the substratum. 11.12.2. Release of hepatocytes from culture substrata

A variety of methods for release of hepatocytes from different substrata using different combinations of collagenase, trypsin, exposure to chelators and mechanical disruption have been reported. From most substrata it is possible to release hepatocytes in sheets by incubation with collagenase but in our experience it has not been possible to achieve disaggregation of sheets to single cell suspensions suitable for counting or in which most cells are undamaged. Methods employing crude collagenase seem to be adequate to separate cell aggregates from plastic protein films or collagen gels. Con-

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centrations of collagenase in PBS ranging from 0.5 to 1.5 mg/ml have been employed. To release hepatocytes, discard the medium and wash monolayers twice with PBS to remove unattached cells. For monolayers on plastic or protein films, add 2 ml collagenase solution (0.5 mgml in PBS) and for cells on collagen gels or gels on nylon grids, use 4 ml collagenase solution (0.5-1.5 mdml). Incubate at 37°C. The progress of cell release can be followed by periodic inspection under a microscope and will require 5-10 min for protein films and up to 20 rnin for collagen gels. Longer incubations were required to release cells from biomatrix (Enat et al., 1984). Release of cells from dishes towards the end of the incubation may be helped by using an automatic pipette, with wide bore tip, to pipette collagenase solution over the surface of the dish. Pipetting can also be used to help break up partly digested collagen gels. Collect the cells by centrifugation for 3 rnin at 100 x g, wash once by resuspending cells in 5 ml PBS and centrifuge for 3 rnin at 100 x g . The cell pellet is then treated as appropriate for parameters under investigation. Variations of this procedure are described by Michalopoulos and Pitot ( 1 979, Sirica et al. (1979), Wilson and McMurray (1981), Enat et al. (1984), Flaim et al. (1985) and Dunn et al. (1989). Alternate procedures for releasing cells from plastic or protein films (not tested in our laboratory) have been described. Williams et al. (1977) reported that 0.25% trypsin released cells without increasing the proportion that were stained by trypan blue and Bissell and Guzelian (1980) reported that a method using trypsin quantitatively converted a monolayer on collagen-coated dishes to single cells. In their procedure, monolayers are washed twice with Calf- and Mg2+-freePBS containing 0.5 mM EGTA and 25 mM Tricine, then incubated for 10 min in the same buffer with 0.1%~trypsin. Soybean trypsin inhibitor is added, followed 4 min later by addition of DMSO to a final concentration of 10'%1 (v/v). Bissell et al. (1986) have also used incubation with 10 mM tetracaine and 1 mM EDTA in Ca?+ and Mg2+-freePBS to release hepatocytes from protein films. To remove cells from Matrigel, Guzelian et al. (1989) wash

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monolayers three times with PBS, then overlay monolayers with 2 ml CaZ+and Mg*+-freePBS containing 5 mM EDTA. The cells plus matrigel are scraped from the plastic dish, transferred to a tube on ice and the cells suspended by pipetting until the Matrigel is dissolved. The cells are then collected by centrifugation. The scraping step may lead to some loss of cytosolic proteins from cells during this procedure. 11.12.3. Preparation of whole cell homogenates

To prepare extracts from monolayers for biochemical assays the most common procedure is to scrape washed monolayers directly from the substratum (plastic, protein films, Millipore filters, etc.) into a small volume of homogenizing buffer. Where cells are maintained on collagen gels or on gels supported on nylon mesh, one approach is to digest the gel with collagenase (as in Section 1 1.12.2) prior to cell harvest (Michalopoulos and Pitot, 1975; Sirica et al., 1979; Dunn et al., 1989). Where a concentrated cell extract is not essential for subsequent measurements, an alternate approach is to homogenize washed gels together with attached cells. In some studies, hepatocytes have been scraped from substrata into PBS and pelleted prior to resuspension in the homogenizing buffer. However cytosolic proteins are lost during this procedure (Edwards and Lucas, 1985). In a typical procedure for cell harvest (for instance, prior to assay of enzyme activities), monolayers are washed twice with PBS or saline to remove dead cells and medium, then drained. Ice-cold homogenizing buffer (1-2 ml) is added to the dish and cells scraped off the dish using a polyethylene scraper or ‘rubber policeman’. Alternatively, where radioactive or toxic reagents are not present, a gloved finger is suitable for rapidly scraping cells from plastic or protein film substrata. Subsequent treatment of this damaged cell suspension will depend on the parameters under investigation. In general, homogenization procedures are designed to ensure that all cells are broken and that a uniform extract for pipetting is obtained.

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Where maintaining the integrity of cell organelles is not required and detergent will not interfere with subsequent assays, lysis with Triton X-100 is effective in disrupting hepatocytes. To eliminate aggregates from the resulting extract, brief sonication (10 s) or homogenization with a Polytron homogenizer (Kinematica GmbH) or Ultraturrax homogenizer (Janke and Kunkel KG) for 10-20 s may be desirable. When 0.1% Triton X-100 was added to unstirred monolayers, up to 30 min was required for complete release of cytoplasmic enzymes (Chao et al., 1988) but 0.1% Triton X-100 combined with homogenization for 10 s with an Ultraturrax homogenizer at two-thirds maximum speed was sufficient in our hands to cause maximal release of cytoplasmic enzymes (Edwards et al., 1987a). Similarly, Schuetz et al. (1990) observed disruption of 9O-100% of cells with 0.2% Triton X-100 in a hypotonic buffer combined with homogenization using a Dounce homogenizer with tight-fitting pestle. In our laboratory (Scott, 1982), treatment of hepatocytes with 0.5%)Triton X-100 was found to be more effective than sonication, Ultraturrax homogenization or freeze-thawing in releasing mitochondria1 enzymes. A variety of other approaches have been used to disrupt hepatocyte monolayers. As described by Gellefors and Nelson ( 1 979) for cells in suspension, it is quite difficult to achieve complete disruption of monolayer hepatocytes by mechanical means. For instance, Gebhardt et al. (1978) found that up to 100 strokes of a Dounce homogenizer were required to achieve complete disruption of cells scraped from monolayer cultures. Sonication has been used by a number of laboratories to prepare extracts. Dich et al. (1988) prepared lysates by sonicating hepatocytes for 10 s at 40 W with a Branson sonifier. Supernatants prepared by centrifugation of these lysates at 10,000 x g were used for a number of enzyme assays. Whole cell lysates prepared by sonication have also been used for spectrophotometric assays of cytochrome P-450 (Bissell and Guzelian, 1990; Dich et al., 1988). A number of laboratories (e.g. Nakamura et a]., 1983a) have used two cycles of freeze-thawing to disrupt cells. For various enzyme assays, 30,000 x g supernatants have been prepared from the freeze-thawed

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lysates. Where a complete freeze-thawed lysate is used for biochemical assays, brief homogenization with Polytron or Ultraturrax homogenizers (homogenization for 20-30 s if collagen gels are present) is necessary to give a uniform homogenate for pipetting. When preservation of mitochondria1 integrity during homogenization is desirable, the use of Dounce, glass-teflon, Polytron or Ultraturrax homogenizers may be appropriate (e.g. Wilson and McMurray, 1981) although any mechanical method is likely to require a compromise between the extent of cell disruption and degree of preservation of mitochondria (e.g. Scott, 1982). For assays where preservation of enzyme activity is not required (e.g. assays of cell protein or DNA) monolayers may be solubilized in 0.1-0.3 M NaOH or KOH (e.g. Nakamura et al., 1983b) or in sodium dodecyl sulphate solution (Dunn et al., 1989). 11.12.4. Preparation of subcellularfractions from hepaioqvte monolqws

Procedures for subcellular fractionation as applied to hepatocyte monolayers do not appear to have been studied extensively. However some methods applied to hepatocytes are listed briefly. Nuclei. For the rapid isolation of nuclei in high yield from monolayers for counting, the procedure of Butler (1984) has been employed for hepatocyte monolayers in our laboratory ( Yusof and Edwards, 1990) with excellent results. After swelling in hypotonic buffer, cell lysis with ethylhexadecyldimethylammonium bromide yields a suspension of single nuclei in high yield, free of debris and suitable for counting. A variety of procedures have been used to isolate hepatocyte nuclei (in lower yield) for transcriptional run-off experiments. A procedure modified from Clayton and Darnel1 ( 1983) is given by Schuetz et al. (1990). Plasma membranes. A procedure involving Dounce homogenization, differential centrifugation and separation of membranes on Percoll is described by Nakamura et al. (1983a). Mitochondria. Wilson and McMurray ( 1981) reported a procedure

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in which hepatocytes were detached from dishes with collagenase, washed with PBS and suspended in 0.25 M sucrose, 50 mM TrisHCl (pH 7.4) and 0.1 mM EDTA. The cells were then homogenized for 15 s using a Polytron and mitochondria isolated by a series of centrifugation steps. The procedures based on digitonin lysis of nonmitochondria1 membranes with or without mechanical disruption as discussed in Section 12.3, or the fractionation of hepatocyte homogenates on Percoll as described by Reinhart et al. (1982), may also be useful in preparing mitochondrial-enriched fractions from hepatocyte monolayers. Microsomes. A method involving sonication of scraped cells and centrifugation has been described by Schuetz et al. (1988). 11.12.5.Expression of relative biochemical activities in monolayer culture

Although values are occasionally expressed on a ‘per culture’ basis, biochemical activities are most frequently related to the total protein or DNA content of cell lysates or subcellular fractions. The procedure of Lowry et al. (1951) is commonly used for protein estimation. For DNA estimation, a frequently used and sensitive method is the fluorimetric procedure of Labarca and Paigen (1980) using Hoechst dye 33258. A recent version of this procedure with hepatocytes is given by Dunn et al. (1989). In cells on collagen-coated dishes or other protein films, a small correction may be necessary for the contribution of the substratum to total protein in cell lysates. For cells on collagen gels or Matrigel, activities must be compared on the basis of DNA content, with a correction for the DNA content of Matrigel (Bissell et al., 1987). 11.12.6. Fixation of cell monolayers

Factors relevant in fixation of hepatocytes are probably similar to those for other cultured cell types. This Section only briefly lists some procedures which have been successfully used with hepatocytes.

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In preparation for light or electron microscopy and for various immunohistochemical procedures, fixation with various combinations of formaldehyde, paraformaldehyde and glutaraldehyde, buffered at pH 7.4, have been employed. In general, shorter initial fixation times (3-30 min) are used prior to immunohistochemistry than in preparing cells for electron microscopy or autoradiography. A procedure for preparing monolayers for immunohistochemistry, used with minor variations in several laboratories, is given by Ratanasavanh et al. (1986). Monolayers are washed twice with cold PBS and fixed for 3-5 min at 0-4"C with a solution of 4% paraformaldehyde/O.l% glutaraldehyde buffered with 0.1 M cacodylate, pH 7.4 (other procedures use periods of up to 30 rnin at room temperature) for fixation. After washing with cold PBS, fixed cells are incubated in 0.1 M lysine with 0.1 M sodium cacodylate (pH 7.4) for 15 min, followed by permeabilization with PBS containing 0.2% saponin for 1 h. Saponin (0.2%) is also included during subsequent exposure to antibodies. A number of laboratories include a 30-60-min exposure to 1% BSA or 10% foetal bovine serum in PBS during or after the permeabilization step to prevent non-specific adsorption of antibody and in some procedures, brief exposure to Triton X-100 (0.2-0.5%) is used in place of saponin to permeabilize cells (e.g. Yamamoto, 1989). In preparation of cells for light or electron microscopy, related fixation procedures (without permeabilization) are used with post-fixation in 1% OsO, (e.g. Bissell et al., 1986; Germain et al., 1988). Some laboratories use overnight fixation in 10% formaldehyde in PBS (pH 7) prior to autoradiography. Fixation of monolayers using organic solvents has been used for some immunohistochemicalassays, for many histochemical procedures and prior to autoradiography. On glass slides, fixation of cells for 1 G 3 0 min in acetone at -20°C is appropriate for a number of histochemical procedures (e.g. Germain et al., 1988). However, acetone dissolves plastic culture dishes! Methanol (95%) and various concentrations of ethanol (50-95%) for periods of 3-30 rnin may also be used for fixation. A detailed procedure for autoradiography following fixation for a total of 30 rnin with acetic acid/absolute ethanol (1:3) is given by Butterworth et al. (1987).

CHAPTER 12

Selected specialized techniques

12.1. Introduction Over the past 20 years numerous interesting and ingenious techniques have been devised for use with the isolated hepatocyte preparation. Indeed, so many novel developments have taken place that it is not feasible to provide a comprehensive description of them here. Instead, we include a representative selection, chosen to demonstrate the wide scope of the available techniques. We have had personal experience of all methods where protocols are given. In other cases we cite pertinent references.

12.2. Subcellular fractionation of hepatocytes During the last 50 years the application of new morphological and biochemical techniques has led to a developing appreciation of the complexity of the living cell. No longer is it pictured as a membranous fluid-filled sac, but rather as a highly organized structure containing a multiplicity of organelles, most of which are rich in membranous structures. Substantial gradients of ions and metabolites exist across

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the bounding membranes of these organelles. This non-homogeneous distribution of certain metabolites tends to invalidate any analytical approach based on extraction of the whole cell, and leads to the conclusion that a better understanding of cellular function will require knowledge of the intracellular distribution of ions and metabolites. This would seem particularly true for the liver where the enzymes of two major metabolic pathways, gluconeogenesis and ureogenesis, are distributed between the mitochondria and the cytoplasm. Fractionation of intracellular components of the liver can be carried out on the solid organ, but unless the complex and time-consuming technique of Elbers et al. (1974) is employed, the metabolite concentrations within the various cellular compartments cannot be determined. On the other hand, when the starting material is a hepatocyte suspension, mitochondria can be separated from cytoplasm rapidly and, with little effort, the concentration of the metabolites in the two compartments can be determined. A major criticism of subcellular fractionation techniques is that they, as is the case with whole cell studies, cannot reveal the true activity of the intermediate or ion under study. It is often assumed that all metabolic intermediates exist in free solution in the various cellular compartments, but there is no definite proof of this. Indeed there is substantial evidence that intermediates are often bound to enzymes or other proteins, and that enzymes are frequently associated with membranes (Wilson, 1980; Clegg, 1984; Srivastava and Bernhard, 1986). The binding of metabolites to enzymes can be extensive since the concentration of an enzyme may sometimes be equal to or greater than the concentration of the intermediates of the reaction it catalyses (Srivastava and Bernhard, 1986). Moreover, the activity and solvent properties of intracellular water, especially within the proximity of membranes, is believed to be very different from that of bulk water (Clegg, 1984). Thus, when we use the term ‘soluble’ in discussing cell constituents, we are generally referring to their solubility following cell fractionation, and the term should not be taken to imply that the constituent under study is necessarily freely soluble in situ. Lardy

Ch. 12.

SELECTED SPECIALIZED TECHNIQUES

357

(1965) has coined the term ‘cytosol’ to describe that fraction of the cytoplasmic constituents solubilized on fractionation and this term has come into general usage. It is important to recognize, however, that the concentration of a metabolite measured in the cytosol may not necessarily reflect its activity within the intact cell. Despite these provisos, much useful information can be obtained from subcellular fractionation, and the isolated hepatocyte preparation has proved a valuable tool in studies employing this approach.

12.3. Methods of subcellular fractionation Whereas the distribution of enzymes within the liver cell can generally be determined by fractionation of homogenates of whole liver, the intracellular disposition of metabolites cannot be ascertained in this manner since, on homogenization, loss of mitochondria1 and nuclear intermediates into the cytoplasmic fraction cannot be avoided. Fortunately, the availability of the isolated hepatocyte preparation has offered a way out of this dilemma. It has proved possible to treat suspensions of hepatocytes in a manner that makes the plasma membrane permeable to the soluble cytoplasmic constituents. Rapid centrifugation through silicone oil is then used to separate a particulate, mitochondria-rich fraction from a supernatant fraction that contains almost all the contents of the cytosol. Both fractions are contaminated with other cellular material (Shears and Kirk, 1984a), but it has been shown that this contamination does not interfere with the main purpose for which the procedure was designed, namely to provide a means of measuring the metabolite content of the cytosolic and mitochondrial matrix compartments (Brocks et al., 1980). 12.3.I . Rationale of method of digitonin fractionation

A number of methods have been devised for separating the mitochondria-rich and cytosolic fractions of isolated hepatocytes. Only

358


one of these, the digitonin fractionation technique, first described by Zuurendonk and Tager (1974) has found general favour. The method relies on the findings that digitonin binds specifically to cholesterol, forming an insoluble complex (Windaus, 1909) that results in the induction of membrane defects. There is a large difference in the cholesterol content of the plasma membrane (0.28 pmollmg protein) and mitochondria1 membranes (0.06 and 0.02 pmol/mg protein in the outer and inner membrane, respectively) in rat liver hepatocytes (Colbeau et al., 1971). Hence, exposing cells to digitonin at an appropriate concentration achieves a selective lysis of the plasma membrane and release of the soluble cytoplasmic contents into the extracellular medium. Optimal conditions for lysis must be rigorously established. The concentration of digitonin, the period of contact with the cells and the temperature of the mixture are all important factors for successful cell fractionation. The technique is rapid and allows the direct measurement of metabolites in subcellular compartments, unlike the metabolic indicator methods (Section 12.3.7) (Holzer et al., 1956; Krebs and Veech, 1970) where distribution is derived from total cell contents. 12.3.2. Preparation of solutions

The fractionation medium, which is kept in a flask on ice at U " C , consists of 0.25 M sucrose, 20 mM MOPS, 3 mM EDTA and digitonin (2.5 mglml in a final volume of 2 ml) at pH 7.0. Due to the temperature response of the MOPS buffer, the medium (digitonin-free) is prepared at 22°C and the pH adjusted to 6.7. At working temperature the final pH is 7.0. A digitonin stock solution (50 mg/ml) is prepared freshly by dissolving 100 mg digitonin (Sigma) in 2 ml of the sucrose/MOPS/EDTA medium, by heating in a boiling water bath. The solution is stored at W " C for 5 3 h. Concentrated stock solution (0.1 ml) is diluted with medium (1.4 ml) so as to achieve a final digitonin concentration of 2.5 mg/ml after the sample has been added.

Ch. 12.


359

12.3.3. Preparation of tubes The initial lysis is carried out in ice-cold glass tubes containing 1.5 ml fractionation medium at M " C . In addition, microcentrifuge tubes (total volume 2.2 ml) are prepared with 0.2 ml 8.5% (w/v) PCA at the bottom of the tube and an overlay of 0.65 ml silicone oil mixture, p P 1.03 (see Protocol 6.5). Once prepared, the tubes are centrifuged for 15 s at 12,000 x g in a microcentrifuge (Eppendorf, Model 5415) to sharpen the interface between layers, and then stored in an ice slurry until needed.

Protocol 12.1 Method of digitonin fractionation (i) Before any attempt is made to fractionate samples, it is essential that all tubes and solutions be set up and arranged in an orderly way. We have found it necessary to allow 2 min for the processing of each sample. Hence, incubations are commenced at intervals of 2 min. Add a 0.5-ml sample of the hepatocyte incubation mixture (25-30 mg wet weight) to the fractionation medium contained in the glass tube and mix rapidly while holding the tube partially immersed in a slurry of ice. (ii) After 15 s, layer a 1.4-ml portion of this mixture above the silicone oil in the microcentrifuge tube (see Protocol 6.5). Complete the layering of sample within 30 s and then centrifuge the tube for 20 s in the microcentrifuge. (iii) Transfer 1.2 ml of the resultant supernatant fraction, representing soluble cytoplasmic contents, to a microcentrifuge tube containing 0.2 ml 50% PCA and mix thoroughly. (iv) Remove the remaining portion of the supernatant carefully together with the silicone oil layer and discard. Wipe the walls of the tube clean, using a cotton bud (or wash as in Protocol 6.5 (v)). Add a further 0.4 ml 8.5% PCA to the acid layer containing the centrifuged pellet (mitochondria-rich fraction) and mix vigorously. (v) Centrifuge both acidified fractions to remove precipitated pro-

360


tein and remove samples for neutralization with 2 M KOHl0.3 M MOPS/ 0.2 M KCl. Store the neutralized samples on ice for 30 min and then remove the precipitated potassium perchlorate by centrifugation. Metabolite concentrations in these samples can then be measured. (vi) To estimate the total intracellular and extracellular metabolite levels, omit the digitonin fractionation step and directly layer a measured portion of the suspension onto the silicone oil and centrifuge for 20 s. Repeat steps (iii)-(v). (vii) Contamination with entrained medium of the pellet fractions from both the digitonin and non-digitonin treated samples can be in the measured by including 3H,0 and (hydr~xy['~C]methyl)inulin incubations (Protocol 6.3 or Protocol 6.5). (viii) Calculate the quantity of any cytosolic metabolite by subtracting the amount found in the pellet of digitonin-treated cells from that measured in the pellet of untreated cells. Alternatively, subtract the amount in the supernatant of untreated cells from that found in the supernatant of digitonin-treated cells. The amount of metabolite found in the pellet of digitonin-treated cells reflects the quantity contained in the mitochondrial fraction. (ix) Check that all metabolites have been recovered by comparing the total of the cytosolic and mitochondrial fractions with that obtained by PCA-precipitation of untreated cells. 12.3.4. Validation of digitonin fractionation

The above conditions were established on the basis of the initial work of Zuurendonk and Tager (1974), and modifications introduced by Akerboom et al. (1978) and Zuurendonk et al. (1979), but have been refined for our own particular requirements. Readers are referred to the cited publications, and to those of Siess and Wieland (1976) and Brocks et al. (1980), for a detailed description of the validation of the basic procedure. Our conditions, like those recommended by others, were established by varying both the concentration of digitonin and the time of

Ch. 12.


361

exposure to the digitonin buffe1. Because of variations in the quality of digitonin, this approach is strongly recommended. A critical feature of the procedure is the need to maintain controlled near-freezing temperatures at the time the cells are exposed to digitonin. For this reason, it is important to ensure that the ratio of the volumes of fractionation mixture to sample is at least 3: 1. Higher solution temperatures lead to mitochondrial lysis and hence invalidate the procedure. It is important also to ensure that the cytosolic component is diluted at least 100-fold and that EDTA is added to block all kinase activity. Obviously, if all these precautions are carried out the final concentration of any metabolite in either fraction is likely to be low. Hence, the approach must be combined with refined techniques for metabolite determination (Bergmeyer, 1983). The degree of cross-contamination of cytosolic and mitochondrial fractions caused by digitonin damage to the mitochondrial membrane can be assessed by determination of the distribution of enzyme activities between the two compartments separated by the silicone oil layer. For this purpose it is assumed that LDH is entirely confined to the cytoplasmic compartment, and glutamate dehydrogenase to the mitochondrial matrix. The distribution of adenylate kinase, located in the mitochondrial intermembrane space (between the mitochondrial inner and outer membranes) can be used as an indicator of damage to the outer mitochondrial membrane. Enzyme levels are measured in each fraction, using the protocol described above, except that the PCA in the bottom of the tube is replaced with 0.5 M sucrose/ 1% Triton X-100. We find that for a maximal retention of LDH and a minimal contamination of glutamate dehydrogenase in the cytosolic fraction, an initial incubation of 15 s at 0--4"C with a digitonin concentration of 2.5 mg/ml is optimal. However, it is suggested that research workers interested in using the digitonin-fractionation technique undertake their own validation procedures. These may also include recovery of intracellular metabolites (Akerboom et al., 1978), or determination of the distribution of added radiolabelled species (Siess and Wieland, 1976).

362

I SOLATED H EPATOCYTES

12.3.5. Variations

Janski and Cornell (1980), in a variation to the standard approach, employed conditions in which lysis of the plasma membrane could be achieved within 2-3 s. The main modifications introduced were extensive purification of commercially available digitonin, and its use in combination with a non-ionic detergent, Kyro EOB. A further modification was the substitution of a mixture of brominated hydrocarbons (but see Section 6.2.3) in place of the silicone oils used by others. In view of the temperature sensitivity of the cells to lysis by digitonin, Brocks et al. (1980) carried out the procedure at -5°C.This was achieved by working in a cold-room, and by including toluene and DMSO in the fractionation medium to prevent it freezing. With respect to compartmentation of adenine nucleotides, glutamate and citrate, the results obtained were very similar to those observed when fractionation was performed at M " C . However, the mitochondrial amounts of aspartate and malate recovered were about twice as high as found in the conventional method. This was attributed to continued activity of the malate and aspartate transporters at 0 4 ° C . It would appear that the conventional procedure yields satisfactory results for most metabolites. 12.3.6. Alternative methods of cell fractionation

Since some cholesterol is present in mitochondria1 membranes, the possibility exists that under certain circumstancesleakage of mitochondria] contents may follow exposure to digitonin. Hence, a method that avoids such exposure could offer some advantages. Tischler et al. (1977) have devised a cavitation method that involves mechanical shearing of the plasma membrane of hepatocytes, accomplished by forcing a cell suspension rapidly through a fine needle. This method has not achieved widespread popularity, possibly because of the requirement for special apparatus, and will not be described in detail. A combination of the techniques of digitonin lysis and rapid mechanical disruption has been described by Shears and Kirk (1984a).

Ch. 12.


363

An interesting approach has been adopted by Elbers et al. (1 974) who have employed fractionation of freeze-clamped liver tissue and density-gradient centrifugation in organic solvents. This method has also been applied to isolated hepatocytes (Soboll and Sies, 1983). The cell suspension should prove a better starting material in that a more rapid and effective cooling is likely to be obtained than can be achieved by freeze-clamping whole liver. 12.3.7, Use of the metabolite indicator methodfor determining metabolite

distribution

An alternative method of determining the intracompartmental distribution of metabolites is known as the metabolite indicator method. For example, it may be of interest to know the cytoplasmic concentrations of ATP, ADP and Pi, in order to determine the cytoplasmic phosphorylation potential (AGJ defined as follows: AGp = AGO' + RT In [ATP]/[ADP][Pi] where AGO' is the standard free energy change at pH 7.0, R is the gas constant and T the absolute temperature. The principle of the method is based on the assumption that nearequilibrium conditions exist for many redox and phosphorylation reactions occurring in the cell. The first step is to identify a reaction or reactions involving the nucleotides and known to occur only in the cytoplasmic compartment. The particular reactions usually chosen for this purpose are those catalysed by glyceraldehyde-3-phosphate(GAP) dehydrogenase and 3-phosphoglycerate (3-PG) kinase (Veech et al., 1970,1979; van der Meer et al., 1978; Groen et al., 1982b). On the assumption that these enzymes catalyse 'near-equilibrium' reactions, AGp can be derived from measurement of the concentrations of the other metabolites according to the following equation: "AD+] [ATPI = K,. [ADPI [Pi1 [ NADH]

[GAP] [ 3-PGI

K, is the combined apparent equilibrium constant of the glyceraldehyde-3-phosphate dehydrogenase and 3-phosphoglycerate

364


kinase reactions and can be determined experimentally. The cytoplasmic [NAD+]/[NADH] ratio is determined from the ratio of [pyruvatejl[lactate] and the equilibrium constant of the LDH reaction, and [GAP] is normally calculated from the measured dihydroxyacetone phosphate concentration and the equilibrium constant of the isomerase reaction (van der Meer et al., 1978). This approach has the attraction of obviating the need for cell fractionation, and thereby avoids the risk of alteration of the levels of labile intermediates during the fractionation procedures. Nevertheless, the metabolite indicator method is based on a number of somewhat dubious assumptions, not the least of which is that the reactions chosen maintain near-equilibrium. In addition, corrections have to be made for the degree of binding of certain metabolites to the structural elements of the cell, and there is no certain way of knowing the degree of this binding. For example, the cytoplasmic AGp determined by the metabolite indicator method gives quite different values from that measured by NMR, mainly because no free ADP can be detected by the latter method (Iles et al., 1985). The metabolite indicator method has proved useful in demonstrating relationships between cellular compartments (Williamson et al. 1967; Mehlman and Veech, 1972), but its value for providing information concerning the absolute concentrations of metabolites within cell compartments is doubtful.

12.4. Homogenization of isolated liver cell suspensions Once hepatocytes are separated into a suspension of single cells, they are remarkably difficult to homogenize. The shearing forces which act to break cells in intact liver tissue differ from those acting on isolated cells. Whereas a Potter-Elvejhem homogenizer will disrupt cells in a tissue matrix (such as intact liver), it will not readily break isolated cells. Greater force, such as that exerted by a Dounce homogenizer, is required. Krack et al. (1980) used a Dounce homogenizer to disrupt hepatocytes in a 0.25 M sucrose solution buffered to pH 7.4 with 3

Ch. 12.


365

mM imidazole. The separation of the particulate components was then achieved by differential centrifugation. Alternative methods of disrupting hepatocytes include vacuum filtration of the cells through a scintered filter (Hogg and Kornberg, 1963), sonication (Gellerfors and Nelson, 1979), freezing in liquid N, (see Protocol 6.3), homogenization with an Ultraturrax (Janke and Kunkel KG) or addition of 0.1% Triton X-100. All these methods are too vigorous to allow satisfactory subcellular fractionation, so that if measurement of the distribution of metabolites is required, the method of choice remains digitonin lysis (Protocol 12.1).

12.5. Preparation of cell fragments ( ‘cytospheres’) An unexpected ‘spin-off from the development of the technique for hepatocyte isolation has been the finding that substantial quantities of cell fragments, which we have termed ‘cytospheres’, can be obtained by high speed centrifugation of isolated hepatocytes. These cell fragments have been characterized (Grivell et al., 1986) and shown to consist almost entirely of organelle-free cytoplasm, bounded by plasma membrane (Fig. 12.1). The cytospheres are formed as a result of shearing stresses applied to the cells during centrifugation, leading to cell elongation (Fig. 12.2) and a budding off of the cytosphere from the remainder of the cell body. The cell is irreversibly damaged by this process, but the cytosphere can remain intact for several days. Not unexpectedly, it is found to contain enzymes that would normally be present in the cytosolic fraction, but metabolites are present in very low amounts compared with the situation in vivo. Because the cytosphere suspension may prove useful for a variety of studies, a protocol for the preparation of these bodies is presented here. 12.5.1. Preparation of solutions and cell suspension An isotonic 90% (v/v) Percoll stock solution is prepared by diluting

366


Fig. 12.1. Interference contrast micrograph of cytospheres. (Bar 10 pm, mag. x 1470).

9 vols. Percoll with 1 vol. 2.5 M sucrose solution, and the pH adjusted to 7.4. All other Percoll media used are prepared by dilution of this stock solution with 0.25 M sucrose solution. All sucrose solutions are buffered at pH 7.4 with 0.02 M Tris-HC1. Cells are prepared according to Protocol 2.2 except that they are washed in bicarbonate-free phosphate saline (Protocol 2. l), rather than washing medium (i.e. Ca2+are omitted from the washing medium).

Protocol 12.2 Preparation of cytospheres (i) Transfer the cell suspension to four tubes for centrifugation in

Ch. 12.


367

Fig. 12.2. Micrograph of an isolated hepatocyte immediately following centrifugation at 260,000 x g for 1 h. A membrane-bound bleb apparently devoid of organelles has formed. (Bar 10 pm, mag. x 1650).

a Beckman LS-50centrifuge at 25-30°C using a 55.2 Ti rotor which has been prewarmed to 30°C (a 60 Ti rotor is also satisfactory). Polycarbonate centrifuge bottles sealed with aluminium screw-caps, supplied by Beckman, are used with the ultracentrifuge rotor. However, it is essential to inspect the bottles closely to ensure that they are placed within the rotor so that the force-bearing surfaces of both bottle and aluminium cap are co-linear. Without this care, lateral stress at high g-forces fractures the bottles at the base of the threaded neck, leading to loss of contents during the initial or, more critically, during the subsequent centrifugation step in Percoll-medium. (ii) Accelerate the rotor slowly and linearly over 20-30 min to 50,000 rpm (260,000 x g) using the slow acceleration facility of the ultracentrifuge at setting 1. Centrifugation is continued at 50,000 rpm for 30 min. This centrifugation programme evolved from many preliminary experiments as the one providing the greatest and most reproducible yield of cytospheres whilst simultaneously minimizing the contamination of the final preparation by intact cells. The maintenance of a temperature between 25°C and 3 0 T , and gradual

368


acceleration of the rotor are essential to achieve reasonable yields of cytospheres. While the full yield of cytospheres is produced by the time that acceleration is complete, the further 30-min operation at maximal speed is associated with destruction of intact cells (i.e. those cells capable of trypan blue exclusion). This greatly facilitates the subsequent isolation of a pure, cell-free preparation of cytospheres, since damaged cells are much easier to separate from cytospheres than are intact cells. (iii) All subsequent procedures are performed at ambient temperature (approx. 22°C). Resuspend each pellet, comprising some intact cells, damaged cells, cellular debris and cytospheres, in 5 ml of 0.25 M sucrose solution, with the aid of a glass rod. Use a further 10 ml of 0.25 M sucrose solution to transfer all the suspensions to a Dounce homogenizer with a loose-fitting pestle (Wheaton; type B). Complete the resuspension of the partially dispersed pellets with five very gentle down and up strokes. (iv) In order to remove intact cells, adjust the combined suspensions to 40 ml with 0.25 M sucrose solution, transfer the contents to two centrifuge tubes (30 ml capacity) and centrifuge for 3 min at 17 x g. Retain the supernatant fractions, resuspend the pellets in 40 ml 0.25 M sucrose solution and centrifuge at 17 x g for 2 min. Discard the pellets that now consist largely of intact cells. (v) To separate cytospheres from other contaminants, combine the supernatant fractions derived from both of these centrifugation steps. Divide the combined supernatant into four equal portions and centrifuge for 5 min at 400 x g. Gently but thoroughly resuspend the pellets containing cytospheres, damaged cells and debris in a small volume of a solution containing 25% Perco11/0.25 M sucrose ( p = 1.063),and then transfer the suspension to a 100-ml measuring cylinder. Add an additional amount of Percoll/sucrose medium to bring the volume to 70 ml. (vi) To harvest the cytospheres, gently mix the suspension and distribute it evenly between four 38.5-m1 ultracentrifuge tubes. Underlay each aliquot with 7 ml 700/0Perco11/0.25 M sucrose medium

Ch. 12. (p


369

= 1.122). Centrifuge the tubes at 10,000 x g for 10 min, in a

Beckman LS-50 Ultracentrifuge. Remove the upper debris layer from each tube, and then pool the density-interface regions (each representing a volume of about 4 ml and containing over 80% of the cytospheres). Dilute the pooled cytosphere suspension with 3 vol0.25 M sucrose solution to reduce its density, and centrifuge at 400 x g for 5 min. Discard the supernatant. Resuspend the pellet in 10 m10.25 M sucrose solution in a tared plastic tube and again centrifuge at 400 x g for 5 min. Remove the supernatant fluid as completely as possible without disturbing the pellet, and weigh the tube plus pellet to provide an estimate of the pellet wet weight yield of cytospheres. These low speed centrifugations remove essentially all the Percoll from the suspension. Resuspend the final cytosphere pellet in 2 - 4 ml 0.25 M sucrose solution to provide a suspension containing about 15 mg protein/ml. Cytospheres are counted in an Improved Neubauer haemocytometer. Fig. 12.3 summarizes this preparative procedure. It was found that preparation of cytospheres at ( 3 4 ° C resulted in very poor yields, and that yields at 20°C or 37°C were 10-15%0 less than at 25°C or 30°C. For this reason, ultracentrifugation at 25-30°C is recommended with subsequent steps at ambient temperature. Once purified, however, cytospheres are relatively stable to storage at ( 3 4 ° C for at least 2 days. 12.5.2. Properties of cytospheres Cytospheres are spherical, have a mean diameter of 9.2 pm (S.D. = f 3.2) and a protein content of 225 =t 12 mg/g wet weight. About 3% of the protein from the original isolated hepatocyte suspension is recoverable. Transmission electron microscopy shows cytospheres to possess a trilaminar membrane and a finely granular hyaloplasm generally devoid of organelles, filaments and microtubules (Fig. 12.4). Freeze-fracture studies reveal a membrane structure typical of a plasma membrane. Ouabain and wheat germ agglutinin-binding studies in-


370

Cell suspension from 1-2 rats (100mg wet weightlml) x g,30min, after uniform acceleration for 30 min) Subsequent steps at 22%

Pellet resuspended in, 0.25M sucrose solution

Pilet Supernatant

Resuspended in 0.25M sucrose solution (17 x g, 2 min)

Supernatant Combine Supernatants

1

- - - - - - - - (400 x g , 5 min) Pellet, consisting of 0s heres, Intact and damaged cells and debris, resus ended in%mPof 25% percolVO.25Msucrose solution. Dlvbed into four centnfu e tubes, each underlayed with 7ml70% PercoiV!.25M sucrose solution.

- - - - - - - - (10,000 x g. 10 mln) damaged cells, debris

intact cells Approximately 4ml density-interface removed; diluted with 3 VOI 0.25M sucrose solution - - - - - - - - (400 x g, 5 min) Pellet drained; resuspendedin lOml 0.25M sucrose solution in tared conical centrifugetube

- - - - - - - - (400 x 8. 5 rnin) Pellet drained, weighed; resuspended in 2ml0.25M sucrose solution

Fig. 12.3. Flow diagram of routine procedure for the preparation and purification of hepatocyte cytospheres.

Ch. 12.


Fig. 12.4. Transmission electron micrograph of a cytosphere. (Bar 5 pm, mag.

371

X

5000).

dicate that the original orientation of the plasma membrane is maintained throughout the formation of the cytospheres. The cytospheres have also been characterized biochemically. Cytospheres are enriched in the enzymes normally associated with the hyaloplasm, whereas the activities of enzymes localized in organelles are greatly diminished. Lipid analysis of the cytosphere membrane indicates that it is derived from the plasma membrane of the hepatocyte. Cytospheres are sensitive to changes in the osmolarity and ionic composition of their environment. Cytospheres should therefore prove a useful preparation for the study of organelle-free cytoplasmic metabolism and of plasma membrane receptor and permeability pro-

312


perties. It may also prove possible to obtain a purified preparation of plasma membranes by osmotic lysis of cytospheres followed by differential centrifugation, perhaps combined with Percoll fractionation (Reinhart et al., 1982).

12.6. Preparation of plasma membrane fractions Nicotera et al. (1985) have devised an ingenious method for preparing purified plasma membrane fractions from isolated hepatocytes in suspension, by binding the cells to polyacrylamide beads coated with polyethyleneimine (Affgel731, Biorad). To lyse the attached cells and remove intracellular components, the beads are resuspended in a small volume of Tris buffer and vortexed for four periods of 5 s. After four washes, the beads with attached cell plasma membranes are stored on ice until required. Where a highly purified preparation, free from intracellular membranes, is required, it may be feasible to use cytospheres as the starting material.

12.7. Permeabilized cells 12.7.1. Introduction

Many studies of intracellular processes in hepatocytes involve the use of agents which do not readily penetrate the plasma membrane. Obviously, such studies would be greatly facilitated if the permeability of the membrane to these compounds could be increased. It could also be advantageous to render the plasma membrane selectively permeable to low molecular weight compounds, such as ions, cofactors and metabolites, normally contained within the cell. To a limited extent this has been achieved. Permeabilized hepatocytes have been used to study a number of processes such as CaZ+fluxes in intracellular organelles (Burgess et al., 1983; Hughes et al., 1987), the mechanism

Ch. 12.


373

of action of inositol 1,4,5-trisphosphate in releasing Ca2+from the endoplasmic reticulum (Burgess et al., 1984; Joseph et al., 1984), the properties of mitochondria in situ (Allan et al., 1983), the process of autophagy (Gordon and Seglen, 1982) and the kinetics of enzymes in situ (Boon and Zammit, 1988). There are two methods of hepatocyte permeabilization which have been used with a modicum of success. These are the treatment of the cells with a chemical agent such as saponin, filipin or digitonin, or exposure of hepatocytes to a series of high voltage electric discharges (electro-permeabilization). Saponin, filipin and digitonin produce permanent changes in plasma membrane permeability whereas electric discharge appears to cause only a temporary increase in permeability. Ideally, the permeabilizing process should have no structural effects on components of the cell other than the plasma membrane. Unfortunately, it would appear that most permeabilizing treatments, especially those involving chemicals, do affect other structural components of the cell. Accordingly, in drawing conclusions from permeabilization studies, it must be appreciated that in many cases, the results have been obtained with hepatocytes with lesions not confined to the plasma membrane.

12.7.2. Rationale for permeabilization of hepatocytes with saponin Saponins are glycosides which occur naturally, and are found predominantly in plants (Tschesche and Wulff, 1973).These molecules form complexes with cholesterol in the plasma membranes of animal cells. This results in the formation of holes ranging in diameter from 0.1 to 1 pm (Brooks and Carmichael, 1983). Treatment of cells with an appropriate concentration of saponin is largely selective for the plasma membrane because, relative to intracellular membranes, it is enriched in cholesterol (Colbeau et al., 1971). Similar considerations apply to treatment of cells with digitonin (Section 12.3). The incubation medium for the permeabilization of hepatocytes (permeabilization medium) is designed to reflect the intracellular ionic

374


environment. Permeabilization medium contains 20 mM NaCI, 100 mM KCl, 5 mM MgSO,, 0.96 mM NaH,PO,, 25 mM NaHCO,, and 2% defatted and dialysed BSA. The medium is gassed with carbogen and adjusted to pH 7.2 (Burgess et al., 1983).

Protocol 12.3 Permeabilization of hepatocytes using saponin (i) Isolate and wash the hepatocytes in the normal manner. Centrifuge the hepatocytes at 50 x g and resuspend in permeabilization medium at a density of 2 x lo6 cells/ml. Incubate the cells for 15 min at 37”C, add saponin (50 pglml) and incubate for a further 5 min. Centrifuge the cells at 50 x g for 1.5 min at room temperature and resuspend the pellet in an equal volume of permeabilization medium (free of saponin, but containing 1.4 mM Na,ATP2-, 4.7 mM creatine phosphate and 25 pg/ml creatine kinase as an energy source for the permeabilized cells) at a cell density of 2 x lo6 cells/ml. For the particular experimental conditions employed in a given laboratory, it will be necessary to determine the optimal ratio of saponin to cell density required to render the plasma membrane permeable to small molecules with minimum loss of intracellular proteins. This can be done by monitoring the loss of “Rb from cells pre-loaded with 86Rband the leakage of LDH (Burgess et al., 1983; Hughes et al., 1987). The optimal concentration of saponin is that which induces a loss of about 75% 86Rbwith little loss of LDH. (ii) The degree of permeabilization of the cells can also be assessed using trypan blue and a marker for the volume of the extracellular space such as [,Ta]EGTA. More than 90% of the hepatocytes should be permeable to trypan blue with a loss of only 20% of the LDH activity over a period of 2 h. The volume of apparent extracellular space accessible to [4SCa]EGTAshould increase from approx. 0.35 to approx. 0.7 pl/mg pellet wet weight (Burgess et al., 1983; Hughes et al., 1987), as the marker enters the permeabilized cells. A significant advantage of the saponin method for the permeabilizd-

Ch. 12.


315

tion of hepatocytes is that it is relatively simple and does not require expensive equipment such as that used for electro-permeabilization. However, a disadvantage of the saponin method is that it is difficult, indeed perhaps almost impossible, to obtain a satisfactory balance between adequate permeabilization of the plasma membrane and lack of significant damage to intracellular membranes such as those of the mitochondria and endoplasmic reticulum. This also applies to the use of other agents such as filipin, which chemically disrupt the plasma membrane. The disruption caused by saponin is dependent on both the concentration of saponin and the time of exposure of the cells to this compound. There is no evidence that hepatocytes ‘re-seal’ after treatment with saponin. This contrasts to the recovery observed after electro-permeabilization. 12.7.3. Other chemical methods for hepatocyte permeabilization The use of filipin to render the plasma membrane of hepatocytes selectively permeable to small molecules was first reported by Jorgenson and Nordlie (1980). Filipin is a polyene antibiotic which binds to cholesterol in cell membranes. As in the case of saponin, filipin is selective for the plasma membrane of cells because that membrane has a relatively higher concentration of cholesterol than other cellular membranes. The resulting disruption of the membrane renders it permeable to compounds of low molecular weight. The actions of filipin on hepatocytes have been characterized by Jorgensen and Nordlie ( 1980) and Gankema et al. (1981). The latter group incubated cells in the presence of 50 pM filipin at 37°C for 1 min in a medium which contained 250 mM mannitol. The cells were then washed twice with the same medium in the absence of filipin. This procedure was found to render the cells permeable to sucrose, inulin and glycerol 3-phosphz re. A gradual leakage of LDH from the cells was observed. This was partially prevented by the inclusion of glutathione and ATP in the incubation medium. Digitonin has been used not only to achieve sub-fractionation of

376


isolated hepatocytes (Section 12.3.1) but also in an attempt to bring about selective permeabilization of the plasma membrane to ions and metabolites but not macromolecules (Boon and Zammit, 1988). Again, it must be emphasized that the integrity of components of the cell other than the plasma membrane is also highly likely to be affected by such treatment. In our hands, hepatocytes exposed to even mild chemical permeabilization lose their ability to synthesize glucose from lactate. A number of workers have used a low concentration of ATP (approximately 1 pM) and the absence of extracellular Mg2+in attempting to render the plasma membranes of some animal cells selectively permeable to molecules of a low molecular weight (Cockcroft and Gomperts, 1980; Gomperts, 1983; Ratan et al., 1986; Mustelin et al., 1986; Steinberg et al., 1987). Hughes and Barritt (1989) employed this method in an attempt to make the plasma membrane of hepatocytes permeable to an analogue of GTP, guanosine 5 ’-[y-thioltriphosphate (GTP[S]). They found that incubation of hepatocytes in the presence of 3 pM ATP and in the absence of MgZ+in a modified KrebsHenseleit medium (see Section 8.2.2) caused a small enhancement of [14C]sucrose uptake compared with cells incubated in the same medium but without ATP and in the presence of 1.2 mM MgSO,. It was concluded that ATP does not readily induce the permeabilization of the liver cell plasma membrane as it does in a number of other cell types (Mustelin et al., 1986; Steinberg et al., 1987).It was speculated by Hughes and Barritt (1989) that the subsequent uptake of [14C]sucroseand GTP[35S]is due to micropinocytosis (Sasaki et al., 1987). Indeed, use of the property of liver cells to undergo micropinocytosis may provide a mechanism by which agents can be introduced into the cytoplasmic space of the hepatocyte with minimal disruption of the cells. 12.7.4. Electro-permeabilization

Electro-permeabilization (or electroporation) of hepatocytes involves subjecting cells to a single or multiple brief high-voltage pulses (e.g.

Ch. 12.


377

five pulses of 2 kV/cm of 50 ps duration) (Gordon and Seglen, 1986). An increased number of pulses or a higher voltage can cause irreversible membrane damage. The mechanism of permeabilization is not known, but may be related to loss of plasma membrane potential (Bernhardt and Pauly, 1973; Kinosita and Tsong, 1977; Zimmermann, 1982) and the production of pores in the cell membrane (Serpersu et al., 1985). For a period after treatment, small charged molecules pass freely through the membrane while larger charged molecules (e.g. ATP, M , 651) and cytoplasmic proteins are retained within the cell. While the cell is in this permeable state, it is possible to introduce small neutral molecules by diffusion. Moreover, rat hepatocytes in suspension have been transfected with plasmids by electroporation (Tur-Kaspa et al., 1986). Sucrose, lactose or raffinose, non-metabolizable substances, to which the plasma membrane is normally impermeable, can enter electropermeabilized cells (Seglen et al., 1986). The cells are subjected to the high voltage treatment at 37°C and then allowed to stand at M " C for about 1 h, the substance to be introduced being present during both treatments. This appears to be an efficient method of ensuring a high incorporation of the neutral molecules. While held at 0--4"C, cells remain permeable. Subsequent incubation at 37°C for < 15 min 're-seals' the hepatocytes, possibly by restoring the energy-dependent plasma membrane potential (see Section 12.9.1). This results in a trapping of the introduced substances inside the cell. Fusion of two or more electro-permeabilized cells is also frequently observed, especially when a high concentration of cells is used during electro-permeabilization or 're-sealing' steps (TeissiC et al., 1986). Seglen and co-workers (Gordon and Seglen, 1982; Seglen and Gordon, 1984; Gordon et al., 1985) have used electro-permeabilization, combined with centrifugation through Metrizamide and digitonin fractionation, to study autophagy and protein lysosomal degradation. Under optimal conditions, proteolysis is only reduced by 10% in these permeabilized cells. Although electro-permeabilized cells are not

378


permeable to trypan blue (Mr961), dyes of slightly lower molecular weight such as erythrosin B ( M , 880) and eosin (Mr620) are able to enter the cells, thereby providing a rapid method of estimating the efficiency of permeabilization. The salt content and temperature of the medium, the size of the vessel used during electro-permeabilization,the suspension volume, the number of cells per ml and the decay time of the electrical pulse, as well as the pulse number, strength and duration, are all important variables during electroporation. The technique of electro-permeabilization is not easy to establish, optimal electro-permeability being achieved within a narrow range of conditions. Biotechnologies and Experimental Research Inc. (BTX), makers of electro-permeabilizationequipment, provide a valuable consulting service.

12.8. Perifusion of hepatocytes The hepatocyte is an active cell, rich in mitochondria and capable of achieving an 0,-uptake of over 15 pmol/g wet weight per min under stimulated conditions. While this high metabolic rate allows changes to be observed over a relatively short time, the rapid rate of substrate consumption and product accumulation can frequently lead to experimental difficulties. This is perhaps best illustrated by an example. The rate of palmitate uptake and metabolism by hepatocytes from fasted rats is sensitive to fatty-acid concentrations below 2 mM. Rates of removal are relatively rapid, up to 1 pmol/g wet weight per min at a fatty acid concentration of 2 mM in the incubation mixture, and about 0.5 pmol/g wet weight per min when present at 1 mM. It follows that 100 mg wet weight of hepatocytes incubated with 2 mM palmitate in a volume of 2 ml will remove 25% of the added fatty acid in 10 min. Under these experimental conditions, a linear rate of palmitate uptake cannot be maintained in the incubation medium. Other examples of the difficulty of maintaining steady-state condi-

Ch. 12.


379

tions due to changing substrate or product concentration during the incubation period are too numerous to mention. The standard means of incubation of cells in a flask is also not suitable for experiments where it is desired to examine whether or not the effects of a particular agent are reversible. Once added, there is rarely the possibility of removing it, although some substances bind so readily to albumin that its addition to the incubation medium effectively removes the agent. The alternative possibility of centrifuging the cells and resuspending them in fresh medium leaves much to be desired. Another disadvantage of the standard incubation procedure is the difficulty it presents for monitoring gradual or stepwise alteration of substrate concentration or the extracellular environment, e.g. PO, or pH. To avoid these disadvantages of the standard incubation procedure, the technique of perifusion has been introduced. A portion of the cell suspension is incubated in a chamber, which is sealed with a filter to prevent cells, but not medium, from exiting. Incubation medium is pumped through the chamber to provide substrate and 0, to the cells. It may be asked why not perfuse an intact liver. Perfusion of the isolated liver requires the presence of erythrocytes in the perfusion medium or a very high flow rate to maintain satisfactory oxygenation of the tissue. The main advantage of isolated cell perifusion is that a much smaller rate of flow per gram of hepatocytes is required for oxygenation, and hence greater inflow-outflow differences can be maintained across the chamber. Also, the suspension is comprised almost entirely of hepatocytes and sampling of cells during the perifusion is considerably easier than during perfusion of whole liver. The smaller perfusion volumes required lead to a considerable cost reduction particularly if expensive substrates, such as radiolabelled compounds, are being employed. 12.8.1. Approaches to perifusion

Despite the obvious value of the perifusion technique for answering


380

certain kinds of biological question, the method has not gained widespread popularity. Fewer than ten groups (see Crisp et al., 1982) have described the design of a perifusion apparatus and presented data obtained by perifusion of hepatocytes. The probable reason for this is that perifusion of liver cells proves not to be easy. The principle difficulty is the containment of the cells in the chamber without damage while maintaining an adequate flow of perifusion medium. Damage to the cells inevitably leads to the release of DNA and protein, and clogging of the filter. This raises the pressure within the chamber and brings the perifusion prematurely to an end. Van der Meer and Tager (1976) were the first to describe a system suitable for hepatocyte perifusion, based on an Amicon Stirred Cell (model 12). Subsequently Groen et al. (1982a) devised an improved model, which had some significant differences. These were the placement of the filter and the outlet at the top of the chamber, a sloping top to the chamber so that the filter was no longer horizontal (Fig. 12.5) and the use of a different filter. All these changes were designed

samplingport ventilationport

membrane filler cell suspension magnetic stirring bar

fusion ports

wafer jacket

water jacket inlet (outlet on other side)

Fig. 12.5. Perifusion apparatus.

Ch. 12.


38 1

to reduce filter clogging, which still remains the chief difficulty with most perifusion systems. Attempts to avoid this difficulty have been made by suspending the cells in a gel-matrix, thereby obviating the need for a filter. Crisp et al. (1982) have described a relatively simple system that does not require manufacture of a complex chamber, but utilizes chromatography columns. The matrix is Sephadex G-50, and a cellulose suspension is used to plug the orifices. The chief disadvantage of this approach is that sampling of the cells during the perifusion is not possible. The cells can be retrieved from the Sephadex matrix at the end of the perifusion, but not under conditions where metabolite concentrations are meaningful. Another difficulty associated with the use of a gel matrix relates to measurement of the degree of cell damage. A possible approach to this has been adopted by Weigle et al. (1983, 1984) who employed a 10-ml plastic syringe as the perifusion chamber, and used a matrix of Bio-Gel P-2. The degree of cell damage was ascertained by determining the ability of the cells on the column to take up [Wlouabain from a mixture of this isotope and ['4C]sucrose. The latter is excluded by the cells and so provides a reference. Ouabain uptake by hepatocytes can then be related to intact cell mass on the column. The inability to sample cells during or after the perifusion is considered a significant disadvantage of methods that employ a gelmatrix and for this reason the apparatus of Groen et al. (1982a) is preferred. Nevertheless, the approach using a gel-matrix is well worth exploring if sampling of the cells is not required. 12.8.2. Description of perifusion apparatus

The apparatus consists of a water-jacketed, thermostatted perspex chamber (internal volume 12 ml) with a sampling port, including an air-bleeding vent, at the top of the chamber (Fig. 12.5). The perifusion medium enters the chamber at the bottom and leaves via an outlet at the top. Cells are retained in the chamber by a membrane filter, 25 mm in diameter, supported by a sintered-glass disc. Suitable mem-

382


brane filters with a pore size of 12 pm (Cat. No. 400006) are available from Schleicher and Schuell. The Sartorius product with a pore size of 8 pm (Cat. No. 1130125N) is also satisfactory. The cells in the chamber are kept in suspension by a magnetically rotated, cylindrical stirring bar. The rate of stirring must be sufficient to prevent cells from adhering to the membrane filter. The composition of the perifusion medium is identical to that used in standard incubations, but generally does not contain albumin. It is equilibrated with carbogen.

Protocol 12.4 Perifusion procedure (i) Place a membrane filter in position in the dry chamber and start the pump so that perifusion medium gradually fills the chamber. The desired pump flow-rate is set so that 0, and substrate concentrations in the chamber do not become limiting factors. This rate is usually 3-5 ml/min. When medium is flowing from the outlet tube, stop the pump, clamp the chamber-outlet tube to prevent backflow and remove sufficient medium from the chamber via the sampling port to allow introduction of the required volume of cells (e.g. up to 600 mg wet weight cells). (ii) Re-start the pump and fill the chamber completely, ensuring that air-bubbles are bled off through the vent. Close the vent, unclamp the outlet tube and switch on the stirrer to begin the perifusion. (iii) Introduce substrates either by addition to bulk medium (in which case, the pump inlet tube is transferred to the medium of choice) or via a second pump by slow infusion of concentrated solutions of substrates into the perifusion medium just before entry into the chamber. Collect samples of the perifusate from the outlet tube. Because a low degree of leakage of enzyme (e.g. LDH) occurs from the cells in the chamber, deproteinize collected fractions with 1 M PCA to prevent unwanted reactions that might occur prior to or during subsequent metabolite assays. Before assay, neutralize the acidified

Ch. 12.


383

samples with a 2 M KOH/0.3 M MOPS/0.2 M KCl solution. Samples of the perifused cells can be collected during the experiment, via the sampling port. The dilution of cell density brought about by such sampling must, of course, be taken into account. (iv) When required, the 0,-content of the perifusate can be measured by use of a flow-cell and 0,-electrode positioned close to the chamber outlet. Provided that the flow rate is known, the rate of consumption of 0, by the cells in the chamber can readily be calculated. The duration of an experiment can be over 3 h, and the rate of infusion of substrate can be varied so that the cells experience a wide range of steady-state substrate concentrations. The overall flow-rate can also be adjusted. The greater the flow-rate, the shorter the time taken for a steady-state concentration of added substrate to be achieved in the perifusion chamber. With a flow-rate of 3-5 ml/min, a new steady state can be established in approximately 30-40 min. As mentioned previously, the main problem encountered during perifusion is the clogging of the membrane filter with consequent pressure increase in the chamber and inlet line and a decrease in flowrate from the outlet. Early detection of this phenomenon can be obtained by connecting a pressure-sensing device between the chamber and the pump. A simple but adequate device is a manometer consisting of a closed, air-filled glass tube, such as a 1-ml pipette, sealed at one end. Alternatively, the pressure gauge from a sphygmomanometer will suffice. In the event of an increase in pressure above approximately 80 mmHg, the perifusion is interrupted and the membrane filter replaced. A common cause of pressure rise is an inadequate stirring rate. 12.8.3. Use of a small chamber The 12-ml sample volume and the consequent extended time for an infused substrate to reach steady-state is obviously a disadvantage. To overcome this problem, McMenamy et al. (1981) have designed a perifusion chamber with a volume of only 0.7 ml. This allows the

384


use of small amounts of cells (up to 100 mg wet weight). A flow rate of 0.7-1.0 ml/min is readily tolerated. According to the authors the system does not require mechanical agitation of the cells by stirring or by continuous shaking of the whole chamber, so that the risk of mechanical damage to the cells is decreased. Despite these apparent advantages the apparatus does not appear to have been widely used, possibly because of the complexity of manufacture. A good description is given in the original article.

12.9. Measurement of cellular membrane potentials Many, if not all, of the cellular membranes contain transport systems that translocate metabolites and ions. In an actively metabolizing cell, this ion transport leads to the creation of steady-states wherein substantial gradients exist across certain membranes, in particular the plasma and inner mitochondrial membranes. Accompanying the creation of these chemical potentials is the establishment of electrical potentials across the membranes. The measurement of these potentials is of value to workers interested in the electrochemical and electrophysiological properties of the cellular membranes. 12.9.I. Measurement of plasma membrane potential

The potential of the plasma membrane (A\kp) can be determined in cells of solid organs by means of a microelectrode, but the use of such a device is difficult, though feasible, for cells in suspension and is considered beyond the scope of this book. An easier way is to employ an ionic species that passively distributes itself across the membrane under study as a function of the charge across it. A difficulty with this technique is ensuring that the ion selected will discriminate between the inner mitochondrial and plasma membranes. For example, some workers have proposed the use of triphenylmethylphosphonium ion (TPMP') for measuring plasma membrane potential. However,

Ch. 12.


385

this lipophilic cation is avidly taken up by mitochondria, and hence will give a totally erroneous impression of the magnitude of the A q P . Because of the active uptake of cations by the mitochondria, it is necessary to measure A q p on the basis of the mitochondria1 exclusion of a permeant anion. Although thiocyanate has been used for this purpose (Hoek et al., 1980), the anion of choice appears to be CI-. In view of the analytical problems involved in measuring small differences in C1- concentrations, it is necessary to use W l - (Baur et al., 1975; Bradford et al., 1985; Berry et al., 1988; Nobes and Brand, 1989). For determination of Aqp, cells are incubated in the presence of TIand (hydr~xy['~C]methyl)inulin(40,000 dpm of each) and 3H,0 (400,000 dpm). Following a minimum incubation of 10-15 min the cells are separated from the medium according to Protocol 6.3. 36Cl-, 14C and 3H in the supernatant and pellet are measured in a liquid scintillation counter which corrects for quench on the basis of prepared variable quench standards of the three isotopes separately (e.g. Beckman LS 3801). Correction for isotope-energy overlap is made by solving simultaneous equations by means of matrix algebra. Intracellular and entrained medium volumes in each individual pellet can be calculated from the distribution of I4C and 3H (Protocol 6.3) and hence C1- concentration outside [Cl-1, and inside [Cl-li the cells can be determined. The calculation of A q p is based on the Nernst equation: RT [Cl-Ii A q p = __ In F [CI-I, ~

where R is the gas constant, T is the absolute temperature and F is the Faraday constant. Strictly speaking, this equation applies only to equilibrium conditions, but it appears to be valid on the assumption that 'C- is passively distributed as a consequence of the active distribution of other ions. To increase the sensitivity of the technique, the quantities of radioisotopes can be increased proportionately.

386


12.9.2. Measurement of inner mitochondrial membrane potential

The uptake of the lipophilic cation TPMP' has been used extensively in the measurement of the membrane potential in isolated mitochondria (Liberman et al., 1969; Heinz, 1981; Nicholls, 1982). The same approach has been taken for the determination of the mitochondrial membrane potential (A",,,) for whole cells (Hoek et al., 1980; Brand and Felber, 1984; Rafael and Nicholls, 1984; Anderson and Jones, 1985; LaNoue et al., 1986) including isolated hepatocytes (Hoek et al., 1980; Anderson and Jones, 1985; Berry et al., 1988). In whole cells, the measurement of A q m using permeant lipophilic cations is complicated by factors that potentially affect the distribution of the cation. These include A q P , which determines the concentration gradient between the cytoplasm and extracellular medium, and the presence of other cellular organelle compartments. It is fortunate that only the mitochondrial compartment has a sufficient volume and magnitude of membrane potential to bring about measureable cellular TPMP' uptake above that attributable to A"p (Hoek et al., 1980; Berry et al., 1988). 12.9.3. Procedure

For the measurement of A",,, in isolated hepatocytes, 2 pM [14C]TPMP+(spec. act. 7,000 dpdnmol) is included in the incubation medium (2 ml). It is necessary to keep the concentration of TPMP' at this level or below because of the observed inhibitory effects of higher concentrations (> 10 p M ) on electron transport and respiration (Berry et al., 1988). At concentrations < 2.5 pM, no deleterious effects are observed and the cation appears inert. At the end of the incubation period, hepatocytes from a 0.5-ml sample of the whole incubation are rapidly pelleted by centrifugation (10,000 x g, for 30 s in a microcentrifuge) and the extracellular TPMP' measured in a 0.3-ml sample of the supernatant by liquid scintillation counting. Steady-state extracellular levels of TPMP' in

Ch. 12.


387

suspensions of metabolizing hepatocytes, incubated at 37"C, are reached within 15 min. Thus, for meaningful measurements a minimum incubation period of 15 min is required. 12.9.4. Calculation of A*,,,

A q mis derived on the assumption that the outer mitochondrial membrane is freely permeable to TPMP+ and that the cation adopts a passive (Nernstian) distribution in free solution across the inner mitochondrial membrane, on the basis of the active distribution of protons: A*,,, = 61.5 loglo

[ TPMP']

,

[ TPMP']

where the subscripts m and c identify mitochondrial and cytoplasmic compartments, respectively. From the measurement of the extracellular concentration ([TPMP+],), it is possible to determine the amount of TPMP' accumulated by the cell, and from this, the intracellular concentration ([TPMP+Ii),on the assumption that the cell water volume is 2.1 ml/g dry weight (Hoek et al., 1980). Exhaustive digitonin fractionation studies have shown that 95% of the intracellular TPMP+ is located within the mitochondria under these conditions (Berry et al., 1988). Thus, if the mitochondrial matrix space is taken to be 12Yo of cell volume (Hoek et al., 1980), a reasonable approximation to the mitochondrial concentration ([TPMP'],) can be obtained: [ TPMP']

,, P

[TPMP' ] i

0.12

In studies where inhibitors are used to modify cellular energy transduction, the above approximation is only valid if [TPMP+],/[TPMP+l,is greater than 50: 1 . At lower ratios there is a disproportionate increase in [TPMP+] within the cytoplasmic compartment, thereby in-


388

validating the assumption that 95% of the cation is accumulated within the mitochondria1 space. In studies with isolated mitochondria 67% of the TPMP' has been shown to be non-specifically bound (Hoek et al., 1980; Brown and Brand, 1985). Of course, it is by no means certain that the same degree of mitochondrial binding takes place in the intact hepatocyte. In the absence of evidence to the contrary, this figure has been adopted for the calculation of A*,,, in the intact hepatocyte. The mitochondrial concentration of TPMP' in free solution is therefore taken to be 33% of the total [TPMP+],,,. The concentration gradient of TPMP' across the plasma membrane must also be considered in the estimation of A*,,,. In our preparations an average value of A q P of 32 mV was obtained, representing a gradient of TPMP' across the plasma membrane of 3.3 1. A correction is also necessary for the measured binding of 10% of the extracellular TPMP' to albumin which is present in all incubations at a concentration of 2.25% (Berry et al., 1988). Taking these considerations into account: [TPMP'],

P

3.31 x 0.9 x [TPMP'I,

(3)

With all these assumptions and corrections Equation I may be restated as follows:

= 61.5 loglo

[TPMP'Ii x 0.33 [TPMP'], x 3.31 x 0.9 x 0.12

(4)

A full discussion of the of the validity of these assumptions appears in the paper by Berry et al. (1988).

12.9.5. Other methods of measuring A*",

Various lipophilic cations have been used to measure A*,,,. Of these tetraphenylphosphonium ion has been the cation most frequently

Ch. 12.


389

employed (Kamo et al., 1979; Demura et al., 1987). However, it appears to have a higher degree of non-specific binding (Demura et al., 1987). On balance, TPMP' is to be preferred. The need for radiolabelled TPMP+ can be avoided by employing a TPMP+-electrode. This technique is useful in that it can be combined with polarography so that A*,,, and the rate of 0,-uptake can be measured simultaneously in the same preparation. The procedure was first developed for mitochondrial preparations (Brown and Brand, 1985) and will not be described in detail here. The procedure using radiolabelled TPMP+ is probably more straightforward, as the electrodes tend to be somewhat unreliable. A spectrophotometric method for determining A*,,, would obviously be convenient. Akerman and Jarvisalo (1980) have explored the use of safranine. They have shown that there is a slow penetration of safranine into the mitochondria of isolated hepatocytes, and that aggregation of safranine molecules occurs in response to A*,,,. This is accompanied by a spectral response that is reversed by valinomycin, uncoupling agents or KCN. Although the method has the advantage of rapid and continuous monitoring, it appears to be only of qualitative value. 12.9.6. Significance of measurement of A*,,,

It will be appreciated that the various correction factors applied in determining the magnitude of A*,,, are somewhat arbitrary. In general, these corrections cause relatively little change in the calculated value. Nevertheless, it may be disconcerting to workers entering this field to find there is no consensus on such important elements as the degree of non-specific binding (Hoek et al., 1980; Brand and Felber, 1984; Andersson et al., 1987; Berry et al., 1988). It should be recognized, however, that generally the absolute value of A*,,, is of no great import. Rather, it is the degree of change that can be induced under various experimental conditions that provides information of interest. A*,,, is a measure of mitochondrial energy state and its response to

390


agents which affect electron transport and oxidative phosphorylation can be expected to throw light on these mechanisms. It is noteworthy that, in hepatocytes from fasted rats, there is an excellent correlation between the values of A*,, phosphorylation potential (AG,) and mitochondria1 redox potential (E,,,,,), when cells are exposed to respiratory inhibitors or uncoupling agents (Berry et al., 1988).

12.10. Measurement of intracellular p H The measurement of intracellular pH was reviewed in depth by Roos and Boron (1981). At that time, the most reliable and frequently used chemical method was based on the assumption that only the uncharged form of a weak acid or base can penetrate the plasma membrane, across which it equilibrates. Since the degree of dissociation of the acid (or base) is dependent on the pH on each side of the plasma membrane and, assuming the dissociation constant is not changed, measurement of the distribution ratio of the acid allows the calculation of ApH (Rottenberg, 1979). The distribution of the permeant weak acid, [2-i4C]5,5-dimethyl-2,4-oxazolidine-dione (DMO) was employed to measure the pH not only of whole cells (Waddell and Butler, 1959) but also of isolated mitochondria (Addanki et al., 1968). Subsequently, values for the intracellular liver pH of approximately 7.2 were reported for perfused liver as well as the organ in vivo (Calvey, 1971; Iles and Cohen, 1974). Using DMO, an intracellular pH of 7.0 has been determined in the cytoplasm of isolated hepatocytes (Hoek et al., 1980). The use of DMO in conjunction with digitonin fractionation (Protocol 12.l ) has allowed the determination of intramitochondrial pH in isolated hepatocytes; Hoek et al. (1980) obtained a value for intramitochondrial pH of 7.97. The use of fluorescent probes to measure intracellular pH has recently become practical with the development of a new class of lipophilic compounds, permeant derivatives of carboxyfluorescein, represented by the acetoxymethyl ester of 2 ’,7 ‘-bis(2-~arboxyethyl)-5(6)-

Ch. 12.


39 1

carboxyfluorescein (BCECF-AM). BCECF-AM was first synthesized by Rink et al. (1982), but is now obtainable from Molecular Probes. This nonfluorescent substance penetrates the plasma membrane and then is hydrolysed by intracellular esterases to liberate the fluorescent weak acid, BCECF, which is trapped within the cell. The use of BCECF for determination of intracellular pH has proved successful for hepatocytes, either in culture (e.g. Gores et al., 1989)or in suspension (e.g. Moule and McGivan, 1990). Intracellular pH values between 6.93 and 7.36 have been obtained. For cultured cells, fluorescence is measured in single cells by using multiparameter digitized video microscopy; cells in suspension are incubated in a thermostatted plastic fluorimeter cuvette, and fluorescence determined in a dual-wavelength spectrophotofluorimeter. The difference in emitted fluorescence after alternate excitation at two wavelengths provides a measure of pH that is unaffected by changes in intracellular dye concentration due to leakage or bleaching. A problem associated with the use of BCECF in hepatocytes is the secretion of the dye by the cells into the surrounding media. This can be largely overcome in studies on cells in culture by using superfusion techniques (Section 11.9.3). Combining the use of a fluorescent probe (6-carboxyfluorescein) with the distribution of DMO, Strzelecki et al. (1984) obtained a value of 7.72 for mitochondria1 pH. Another means of measuring the intracellular pH of hepatocytes involves the use of microelectrodes. This approach requires the hepatocytes to be immobilized, and is therefore best suited to the determination of pH for cells in culture. Details are given by Fitz et al. (1989). The pH, measured in isolated hepatocytes under control conditions, ranges from 7.1 to 7.3.

12.11. Transplantation of hepatocytes Within a few years of the development of techniques for isolation of hepatocytes in high yield, a number of workers began to explore the

392


feasibility of using transplantation of isolated hepatocytes to restore liver function impaired by disease or congenital defect (reviewed by Fuller, 1988). Although there is now good evidence that prolonged surtrival of transplanted hepatocytes can be obtained, there remains some doubt as to the therapeutic value of the procedure. This is in part due to the complication that injection of liver extract, free from intact cells, may be sufficient to cause significant alteration of liver function in the recipient animal. 12.11.1. Transplantation procedures

Isolated hepatocytes for transplantation are prepared by the standard procedures, described for the rat in Chapter 2, or alternatively for larger animals in Chapter 3. There has been considerable debate concerning the optimal site of transplantation, particularly as it was originally believed that hepatocytes would require a continuous supply of portal blood for prolonged survival. In fact, injection of hepatocytes into the portal vein or liver appeared to result in a high early mortality due to host liver infarction and necrosis (Fuller, 1988). Groth et al. (1977) were able to identify hepatocytes in muscle tissue after intramuscular injection, and Jirtle and Michalopoulos ( 1982) observed proliferation when hepatocytes were injected into fat pads. Mito et al. (1978) demonstrated hepatocytes in the splenic pulp up to 6 weeks after innoculation into the spleen. Innoculation into other areas, such as kidney or abdominal cavity, and injection into the coeliac or renal arteries were apparently not successful. Subsequently, Kusano and Mito (1982) observed survival times as long as 18 months for rat hepatocytes transplanted into the spleen of syngenic animals. As a consequence of this work, the spleen has become the preferred site for innoculation (Nordlinger et al., 1985; Fuller, 1988; Henne-Bruns et al., 1989; Hillan et al., 1989; Maganto et al., 1990). Whereas Mito et al. (1978) and most other groups, have concluded that proliferation of hepatocytes can take place within the spleen, Henne-Bruns et al. (1989) have expressed a contrary view. This is ob-

Ch. 12.


393

viously an important issue to clarify, since the number of cells innoculated (see below) can hardly be sufficient to support adequately the hepatic function of animals in severe liver failure. It must also be appreciated that the majority of the long-term studies have employed syngenic cells, although the introduction of cyclosporine A and other immunosuppressive agents has increased the hope of prolonged survival. Another approach (Cai et al., 1984) has been to encapsulate hepatocytes in an alginate-polylysine membrane that can be selectively porous to nutrients and hormones but excludes immunoglobulin and complement (Section 10.7.2),thereby protecting the hepatocytes from the host’s immune system. By far the greatest number of studies have been undertaken in the rat (Fuller, 1988), but dogs were used by Sommer et al. (1979) and pigs by Nordlinger et al. (1985). The transplantation of hepatocytes is a relatively simple procedure, even in animals as small as the rat. Direct injection through a 25-27G needle of 0.5-1.0 ml of cell suspension containing 10-40 x lo6 hepatocytes, usually in up to three splenic sites, is recommended (Fuller, 1988; Hillan et al., 1989; Maganto et al., 1990). It appears unnecessary to clamp the splenic pedicle during innoculation of the cells. 12.11.2. Measurement of hepatocyte function in transplanted cells

There are a number of different approaches to the functional assessment of hepatocyte grafts. One fundamental requirement is the demonstration of surviving cells in the host, since the possibility cannot otherwise be excluded that improvement in host liver function is a result of transfer of some ‘hepatotrophic factor’, rather than the presence of intact cells (Fuller, 1988). For this reason, direct histological demonstration of intact hepatocytes is highly desirable. The difficulty of distinguishing injected donor cells from the hepatocytes of host liver is another reason why injection into the portal vein or liver is generally avoided in experimental studies. Cells injected into the spleen can be examined for hepatocyte-like morphology,

394


the presence of glycogen granules or specific enzymes such as glucose-6-phosphatase or ornithine carbamyltransferase. Electron microscopic studies can also be helpful by revealing intercellular biliary canaliculi and other ultra-microscopic structures characteristic of hepatocytes. Biochemical or physiological studies can be divided into two main groups. In one type of experiment, evidence has been sought for hepatic function at the transplantation site; for example the production of conjugated bilirubin within the spleen. Woods et al. (1982) obtained conclusive evidence for conjugation of bilirubin by extracting conjugates from the spleens of grafted but not control rats. Demetriou et al. (1986), using hepatocytes attached to micro-carrier beads and injected into the peritoneal cavity of Gunn rats, observed a reduction in plasma bilirubin and the formation of bilirubin glucuronides. When allogenic, rather than syngenic, cells were used, the response was transient, disappearing within 6 days. Imaging techniques employing TC derivatives of iminodiacetic acid have also been successfully employed (CuervasMons et al., 1985). The other main group of experiments involves transplantation of hepatocytes in animals that have received treatments likely to induce life-threatening hepatic failure. Evidence is then sought for recovery or at least prolonged survival. The main dificulty with this approach is the possibility that improvement is at least in part a consequence of trophic factors infused together with the cell suspension. The likelihood that this is the case is supported by the fact that Sutherland et al. (1977) obtained some improvement with isolated hepatocytes prepared by mechanical methods, and therefore almost certainly grossly damaged (see Section 1.3). Indeed, as yet no clear evidence has been produced that transplanted cells can provide adequate support in experimental liver failure (Fuller, 1988). The achievement of this goal may have to await the development of reliable methods for producing long-term survival after allogenic grafting, together with more effective means of inducing the proliferation of transplanted hepatocytes.

Ch. 12.


395

12.11.3. Cryopreservation

If hepatocyte transplantation is ever to be available as a therapeutic intervention in humans with severe and irreversible liver failure, a reliable system of cell preservation is likely to be necessary. Perhaps the greatest possibility for such preservation lies with cryopreservation. Maganto et al. (1990) have described a technique based on that of Makowka et al.( 1980). Pellets of isolated cells are placed on ice and the hepatocytes resuspended in ice-cooled Williams’ Medium E/foetal calf serum (4:1). An equal volume of Williams’ Medium E/DMSO (4:1), also held at 0 - 4 ” C , is added to give a final cell concentration of 2.5-3.0 X lo6 cells/ml. The final suspension is distributed among 5 ml cryotubes (Nunc) and the freezing process commences. This consists of the following steps: 30 min at 0 4 ° C ; 60 min at -20°C; 5-6 h at -80°C; 10-15 days in liquid N, (-196°C). To thaw the cells, the vials are introduced directly into a bath at 37°C. Shortly before the contents are completely thawed, the vials are opened and emptied into a flask containing a volume of Williams’ Medium E twice as great as the volume to be thawed. The cells are centrifuged and washed with Williams’ Medium E, and this step is repeated. According to Maganto et al. (1990), the degree of dye-exclusion for frozen and thawed hepatocytes ranges from 45 to 73% with a mean value of 53%. These authors produce evidence that the cells retain some degree of metabolic competence, but this requires further study.

12.12. Studies with hepatocyte ‘doublets’ Research workers interested in cell-to-cell communication have taken advantage of the fact that the isolated hepatocyte suspension contains a substantial number of doublets, i.e. cell pairs. These doublets are best identified in cells that have been newly plated for culture, and it is feasible that some doublets are formed by re-establishment ofjunc-

396


tional complexes during adherent cell culture (Chapter 1 1). However, it is more likely that the majority of doublets represent cell pairs in which the junctions have not been cleaved by the isolation procedure (Spray et al., 1986).These cells are ‘dye-coupled’in that lucifer yellow spreads from injected cells to their neighbours and a bile canaliculus is maintained between each pair of cells. The doublets lend themselves to both electrophysiological and chemical studies. Doublets or triplets can be separated from single cells by centrifugal elutriation (Bernaert et al., 1979). 12.12.1. Electrophysiological studies

By means of sophisticated electrophysiological apparatus, it is possible to demonstrate that the hepatocytes of doublets are electronically coupled (Spray et al., 1986). The preparation is particularly useful for studying the electrical properties of the gap junction and a considerable body of experimental work has been conducted in this area (see for example, Spray et al., 1986; Boyer et al., 1988; Reverdin and Weingart, 1988). Readers interested in these techniques should refer to the cited papers. 12.12.2. Chemical studies

Recently, hepatic doublets have been successfully used to study biliary transport processes (Kitamura et al., 1990). TR- mutant Wistar rats secrete bile acids but markedly fewer other organic anions from the liver into the bile than do control rats. Fluorescence-image analysis of isolated normal and TR- hepatocyte doublets in monolayer culture can be employed to study the transport of fluorescent-labelled bile acid (fluorescein isothiocyanate glycocholate) and a non-bile acid organic anion (carboxydichlorofluorescein diacetate) into the biliary canaliculus of the doublet. On initial isolation of doublets, the biliary canaliculus is collapsed and non-functional. However, after about 5 h in culture secretory activity can be detected. These studies have

Ch. 12.


397

revealed that normal hepatocyte doublets readily transport both bile acid and non-bile acid anions into the canaliculus, whereas doublets from TR- rats transport only the bile acid anion, due to the functional lack of an ATP-dependent organic anion transport system. The defect in TR- mutant rats is phenotypically similar to that seen in mutant Corriedale sheep and in the Dubin-Johnson syndrome in man.

12.13. Separation of periportal and perivenous hepatocytes Although hepatocyte heterogeneity has been recognized for over 30 years (Novikoff, 1959; Rappaport, 1960), it is only in the last decade that functional studies have been carried out (reviewed by Jungermann and Katz, 1989). Much of this effort has been directed to examination of microdissected liver tissue, primarily by biochemical or histochemical means. However, in the last 5 years, ingenious methods for partially separating periportal and perivenous hepatocytes have been devised (Lindros and Penttila, 1985; Quistorff, 1985). Although these methods are still in their infancy, it is to be hoped that further development of the techniques will lead to the possibility of obtaining much purer samples of periportal and perivenous hepatocytes. 12.13.I . Separation by collagenase-digitonin perfusion

The two methods now commonly in use are based on the principle of local collagenase digestion described by Vaananen et al. (1983) and independently by Quistorff et al. (1984). The principle is based on the ability of digitonin perfusion to destroy selectively one region of the acinus, the particular zone affected being dependent on whether the digitonin solution is perfused through the portal vein or in a retrograde manner through the inferior vena cava. The methods as published are relatively straightforward. After establishment of normal hepatic perfusion, a perfusion medium containing digitonin at 4 mg/ml is infused at 37°C for 4 0 - 6 0 s at 7 ml/min

398


(according to Quistorff, 1985) or alternatively a 7 mM (8.6 mg/ml) digitonin solution is infused at 40°C for 2 5 - 4 5 s at 10 ml/min (according to Lindros and Penttila, 1985). A guide to the length of perfusion is provided by the appearance of the surface of the liver immediately following digitonin perfusion, as illustrated in the publication of Lindros and Pentilla (1985). Perfusion is antegrade if destruction of periportal cells is desired, and retrograde if perivenous cells are to be eliminated. Immediately following digitonin perfusion, the direction of flow is reversed and perfusion continued with Caz+-freeperfusion medium as the initial step of a standard two-step hepatocyte isolation procedure (Section 2.3). The initial cell suspension obtained after hepatocyte isolation contains much cell debris and the initial proportion of cells excluding trypan blue is generally less than 70%. The damaged cells should be removed by treatment with Percoll (Protocol 2.5) or Metrizamide (Seglen, 1976).Quistorff (1985) reported mean yields per liver of 2.02 x loRand 1.22 x lo8 for periportal and perivenous cells, respectively. Although enzymic markers show that these techniques can frequently achieve a good degree of selectivity, it is apparent that considerable overlap can occur. There seems no doubt that the few layers of glutamine synthase-rich cells bordering the central vein (Gebhardt and Mecke, 1983) are consistently destroyed by retrograde perfusion. However, it is doubtful if all the hepatocytes that would normally be designated perivenous when liver is subjected to micro-dissection, are destroyed by perfusion in this manner. Likewise, it is probable that a considerable percentage of the periportal cells are spared during antegrade perfusion, even when a substantial number of the perivenous hepatocytes are concomitantly being destroyed. The main reason for this is that it is not feasible to perfuse each hepatic acinus to exactly the same degree. Nevertheless, this approach shows considerable promise and is likely to undergo refinement in future years.

Ch. 12.


399

12.13.2. Separation by centrifugal elutriation Sumner et al. (1983) have attempted to fractionate periportal and perivenous hepatocytes by the technique of centrifugal elutriation. Hepatocytes, prepared by standard methods and resuspended in a medium containing 2% BSA at pH 7.3, were fractionated using a Beckman JE6-B elutriator rotor on a Beckman 52-21 centrifuge modified for operation at low speed (0-2000 rpm full range). Eight populations of cells were recovered and characterized by fluorescent dye uptake, cytochrome P-450 recovery, enzyme distribution and response to phenobarbital or 3-methylcholanthrene. These researchers found that though there was enrichment of various fractions with periportal or perivenous hepatocytes, complete separation was not obtained. Rather, they considered that the observed metabolic heterogeneity was related more to cell size than the location of cell origin. Fractionation by centrifugal elutriation on the basis of hepatocyte size has also been carried out by Bernaert et al. (1979), but these workers made no attempt to distinguish periportal and perivenous hepatocytes.

12.14. Future developments In 1967 when I was conducting the initial experiments on collagenase perfusion of the liver, my intent was merely to produce a preparation that was more suitable than surviving slices for studying hepatic gluconeogenesis. I was more than content to find that suspensions of isolated hepatocytes could synthesize glucose from lactate at about 60% of the rate attained by perfused liver (Berry and Friend, 1969). It was not until 5 years later that, during a brief stay in Oxford, I was encouraged by H.A. Krebs to use bicarbonate-saline medium. To the surprise of us both, the isolated hepatocytes proved capable in this medium of surpassing rates of gluconeogenesis and ureogenesis

400


observed with perfused liver. There is little doubt that the inital catalyst for the spread of the methodology was the enthusiasm of Professor Krebs for the new technique, an enthusiasm that was in marked contrast to his previous justified scepticism. Since those exciting days I have not ceased to be astonished by the ingenuity shown by researchers around the world in finding new ways to use the isolated hepatocyte preparation. The isolated hepatocyte has proved an extraordinarily resilient cell, capable of resisting all but the most severe insults imposed by the research worker. Because of this, and because of the relative simplicity of the method, it seems highly probable that many more remarkable techniques will be developed for exploiting the preparation. I, and my co-authors, are hopeful that this manual will form a firm foundation for research workers to build future developments in the field, and that we will have the pleasure of incorporating such developments in the next edition!

Appendix 1

Mediu

The preparation of media recommended for the isolation of hepatocytes by the oneand two-step procedures are given in detail in the text. Bicarbonate-free phosphate saline Perfusion medium without collagenase Cell washing medium Perfusion medium with collagenase Medium for dispersion of collagenase-digested liver Media for two-step procedure Percoll medium for removal of damaged cells Fortified media

Protocol 2.1. (i) and (ii) Protocol 2.1. (iii) Protocol 2.1. (iv) Protocol 2.1. (v) Protocol 2.2. (ix) Section 2.3.1. Protocol 2.5. ( i ) Section 10.5.1.

Other media for which recipes are given. Perfusion medium for rat hepatocyte preparation by non-enzymatic technique Medium for preparation of damaged cells Standard incubation media Incubation media for hormone studies Dulbecco’s phosphate-buffered saline

Section 3.3.2. Section 3.7. I . Section 6.4.6. Section 8.2.2. Section 11.2.

Culture Media not defined in Table 1 I . 1 can be found in catalogues from companies such as Flow Laboratories (ICN-Biomedicals).

402


TABLE A. 1 Composition of important basic salts media

Components

NaCl KCI CaCI, , 2H,O KH,PO, MgSO, 7H,O MgCI, 6H,O Na,HPO, NaHCO, D-g~ucose Phenol red. Na

.

.

Amount (mg/l) Hank:

Krebs-Henseleit bicarbonateb

8000 400 185 60

6900 350 373 160 290

100

I00

48 350

2090

1000

20

Hanks medium is normally equilibriated with loo'%, 0, and Krebs-Henseleit bicarbonate-saline with carbogen.

The composition of two other basic media frequently used (with modifications) in the preparation and incubation of isolated hepatocytes are given in Table A.I.

References: "Hanks, J.M. and Wallace, R.E. (1949). bKrebs. H.A. and Henseleit, K. (1932).

Appendix 2

Addresses of suppliers and manufacturers

Amersham International plc. White Lion Road, Amersham. Bucks, HP7 9LL, U.K Telephone: (44-2404) 4444 Fax: (44-2404) 40 Beckman Instruments Inc.. 2500 Harbor Boulevard, Fullerton, CA, 92634, U.S.A. Telephone: (1-714) 871 4848 Fax: (1-714) 7736652 Becton Dickinson and Co.. Clay A d a m Division, 299 Webro Road, Parsippany. NJ, 07054, U S A . Telephone: (1-201) 887 4800 Biorad Chemical Division, 1414 Harbour Way So., Richmond. CA, 94804. U.S.A. Telephone: ( I -41 5 ) 232 7000 Telex: 331-732 or Biorad Laboratories Ltd., Caxton Way, Watford Business Park. Watford, Hertfordshire, WDI 8RP, U.K. Telephone: (44-923) 40322 Telex: 88 I3 192

404


Biotechnologies and Experimental Research Inc. (BTX), 3742 Jewel1 Street, San Diego, CA, 92109, U S A . Fax: (1-619) 4833817 Boehringer Mannheim Biochemicals, P.O. Box 50816, Indianapolis, IN, 46250, U.S.A. Telephone: (1-317) 849 9350 Fax: ( 1-317) 5767525 or Boehringer Mannheim GmbH. Sandhofer Strape 116, Postfach 310120, D-6800, Mannheim 31. Germany. Fax: (49-621) 7592890 British Drug Houses Chemicals Ltd. (BDH). Broom Road, Poole, BH12 4NN England. Telephone: (44-202) 74 5520 Fax: (44-202) 738299 Coulter Electronics, Northwell Drive, Luton, Beds, LU3 3RH, U.K. Telephone: (44-582) 58 2442 Telex: 825074 or P.O.Box 2145, Hialeah, FL, 33012-0145, U.S.A. Telephone: (1-305) 885 0131 Fax: (1-305) 8836820 Eppendorf Geratebau, Netheler and Hinz GmbH, P.O. Box 65 06 70, D-2000, Hamburg 65, Germany. Telephone: (49-40) 53801-1, Telex: 02 174315 epge d Gilson Medical Electronics Inc., 3000 W. Beltline Highway, P.O. Box 27, Middleton, WI. 53562, U S A . Telephone: (1-608) 836 155I Fax: (1-608) 8314451 or 72 Rue Gambetta-B.P. 45, F-95400, Villiers-le-Bel. France. Telephone: (33-1) 39905441 Fax: (33-1) 39945183 Heraeus Equipment, 11 1A Corporate Boulevard, South Plainfield, NJ. 07080, U.S.A. Telephone: (1-201) 755 4800; (800) 441 2554 Fax: (1-201) 754 9494

APPENDIX ICN Biomedicals Inc. (Flow), P.O. Box 19536. Irvine, CA, 92713. U.S.A. Telephone: (1-714) 545 01 13 Telex: 685 580 or ICN Biomedicals Ltd., Free Press House, Castle Street, High Wycombe. Bucks, HP13 6RN. U.K Telephone: (44-494) 443 826 Janke and Kunkel GmbH and Co. KG. Neumagenstraoe 27, D-78 13, Staufen/Br., Germany. Telephone: (49-7633) 831-0 Fax: (49-7633) 83198 Kinematica AG, Luzernerstrabe 147a, CH-6014. Littau-Luzern, Switzerland. Telephone: (41-41) 571257 Fax: (41-41) 571460 Kontes Glass Co., Spruce Street, P.O. Box 729, Vineland, NJ, 08360. U.S.A. Telephone: (1-609) 692 8500 Lux - Division of Miles Lab Inc., 30W 475 North Awrora Road, Naperville, IL. 60566, U.S.A. Telephone: (1-800) 348 7465 Millipore Corporation, 80 Ashby Road, Bedford, MA, 01730, U.S.A. Telephone: (1-617) 275 9200 Fax: (1-617) 8752050 or Millipore S.A., B.P. 307, Saint-Quentin, F-78054. Yvelines Cedex. France. Telephone: (33-1) 30589858 Telex: 698 371 Molecular Probes Inc.. P.O. Box 22010, Eugene, Oregon, 97402. U.S.A. Nunc, Postbox 280 Kamstrup, DK 4000, Roskilde, Denmark. Telephone: (45-42) 359065 Fax: (45-42) 350105

405

406


Perkin-Elmer Corporation. Main Avenue, Norwalk, CT, 06856. U.S.A. Telephone: (1-203) 762 1000 Fax: (1-203) 7626000 or Post Ofice Lane, Beaconsfield, Bucks, HP9 IQA. U.K. Telephone: (44-49) 466 I6 I Fax: (44-75) 3884614 Permutit Co. Inc. (Subs. of Zurn Industries). E49 Midland Avenue, Paramus, NJ, 07652, U.S.A. Telephone: (1-201) 967 6000 Fax: (1-201) 9677540 Pharmacia LKB Biotechnology AB. Box 305. S-16126 Bromma, Sweden. Telephone: (46-8) 7998000 Fax: (46-8) 986364 or 800 Centennial Avenue, Piscataway, NJ, 08854. U.S.A. Telephone: (1-414) 347 7442 Fax: (1-201) 4570557 Plischke and Buhr K.G.. Siemensstrape 6, D-5300, Bonn I , Germany. Telephone: (49-228) 625290 Radiometer A/S, Emdrupvej 72, DK-2400. Copenhagen NV. Denmark. Telephone: (45-I ) 69631 1 Telex: I541 1 or Radiometer-America Inc.. 81 I Sharon Drive, Westlake. OH. 44145, U.S.A. Telephone: ( 1-216) 87 I 8900 Telex: 985276 Sartorius GmbH. Weender Landstrafie 94/108, D-3400. Gottingen, Germany. Telephone: (49-551) 308-1 Fax: (49-551) 308289 or Sartorius Filters Inc.. 30940 San Clemente Street, Hayward, CA. 94544, U.S.A.

APPENDIX Telephone: (1-415) 487 8220 Telex: 338-534 Schleicher and Schuell GmbH, Postfach 4, D-3354 Dassel, Germany. Telephone: (49-5561) 791-0 Fax: (49-5564) 2309 or 10 Optical Avenue, Keene, NH, 03431, U.S.A. Telephone: (1-603) 352 3810 Fax: (1-603) 3573627 Sigma Chemical Company, P.O. Box 14508, St. Louis, MO, 63178-9916, U.S.A. Telephone: (1-314) 771 5750 Fax: (800) 3255052 Wheaton Scientific, 1301 N. Tenth Street. Millville, NJ. 08332, U.S.A. Telephone: (1-609) 8251400 Fax: ( 1-609) 825 I 13 I Worthington Biochemical Group, Halls Mill Road, Freehold, NJ, 07728, U.S.A. Telephone: (1-201) 462 3838 Telex: 3715614 Yellow Springs Instruments Co., Box 279, Yellow Springs, OH, 45387, U.S.A. Telephone: (1-513) 767 7241 Telex: 20-5437

407


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Subject index

Note: page numbers in italics indicate Protocols or experimental details.

A23187 and Ca2+ measurement 218, 219 Abnormal animals, hepatocyte isolation from 7 6 7 7 Acetate metabolism 171, 200 Acetone as solubilizing agent 159 Actin 103, 105, 117, 195 Adenine nuceotides 57, 58, 95, 98, 144--Y47. 189, 363 Adenylate cyclase 204 Adhesion of cells Ca2+ and 3. 4, 10-12, 62 cell adhesion molecules (CAMS) 4, 269-270, 323 ADP-ribosylation 318 Adrenaline, effect on intracellular [Ca2+] 203 Adrenergic agonists 20 I, 203,276.3 1 I , 333 Advantages of isolated hepatocyte preparation I , 14, 116, 158, 179-180, 182, 201. 357

of monolayer culture 252-254 of suspension culture 252-254 Aequorin 224 Aflatoxin 185 Albumin (BSA) effect on cell attachment 261, 282 gene transcription 27 1-275, 307-308, 313, 315 in media 40,58,89,93,155---l57,172, 205, 206, 212, 257, 258, 295, 304, 388 synthesisand secretion 125,254,271, 273, 275, 315, 339, 345 Alcohol dehydrogenase 302, 309 a-Amantin 183 Amino acids depletion 35, 301-302, 321 in media 40, 242, 294, 301-302 metabolism 160, 170, 173-176, 30 1-302 Ammonia metabolism 170 AMP deamination 42 Anaesthesia

442


MS 222 (tricaine methane sulfonate) 69 pentobarbital 25 toxicity 189, 191 Angiotensin II 201, 215 Anoxia 109, 189 Antibiotics in media 210,246,249,258, 297, 305, 330, 333 Apparatus for isolation of hepatocytes collagenase-free 63 foetal and neonatal 74-76 mechanical 78-40

to laminin 280, 287 to matrigel 281, 288-289 to microcarriers 334 to plastic 261, 279-280, 282 Attachment of non-hepatocytes 242-243, 245 Autocrine growth factors 321, 341 Autophagy (autophagocytosis) 109,175, 176. 377 Autoradiography 354

one-step 17-23, 67 sterile 247-248 thin slices 6, 7 two-step 17-23. 32 [3H]Arachidonic acid 208 Arsenazo 111 and Ca2+ measurement 218 Ascorbate 296, 305 Aspartate 1 7 6 1 7 1 Atomic absorption spectroscopy and Ca2+ measurement 216-218 ATP breakdown accumulation of AMP during 145, 146 caused by fructose or glycerol 98, 145, 147 in hypoxia 42, 95, 144, 167 ATP, hepatocyte content of 95, 144-147 ATP/ADP ratio 95. 98, 1*145, 363 Attachment of hepatocytes for monolayer culture after damage during isolation 246 characteristics 279-280 effect of hormones on 280, 282 effect of serum on 261, 280, 345 media for 280-282, 324, 329, 332, 334, 340, 343 significance for cell function 278-279 to collagens 267,286281,282-284 to fibronectin 280. 287

Bacterial petri dishes 261, 282, 283 Barbiturates hexobarbital 181 in culture media 316, 317 in enzyme induction 273, 275, 317 phenobarbital 76, 77, 181, 273, 275. 316. 317 Basement membrane matrix see Biomatrix; Extracellular matrix; Matrigel Bile canaliculi 103, 105. 107, 177, 306, 396 ducts or ductules 243 ductular epithelial cells 243,260, 323, 337-338 transport 108, 177, 323, 396-397 Biomatrix 276, 288 see also Extracellular matrix; Matrigel Biopsies of human liver 7CL72, 343 Blebbing of plasma membrane and Ca2+ 102 drug-induced 101, 102, 111, 119, 196195 in response to cold 100. 101 in response to hypoxia 42, 101, 102 morphology of 102, 117 Bovine serum albumin see Albumin Buffers bicarbonatekarbon dioxide 23-25, 41, 155, 157, 164, 249, 295, 297 HEPES 32, 42, 155, 242, 249, 297. 328

SUBJECT INDEX

in cell isolation 24. 3 2 , 4 1 4 3 . 78,249 in metabolic studies 89. 155, 157-158. 164-166. 205. 212 in monolayer culture 295, 297, 300 in suspension culture 257. 258 MOPS 13@-l31, 358 phosphate 23-25. 155, 157 TES 205 Tris 198 Butyrate 172, 316, 318, 338 Ca2+ activation of collagenase 8. 1 I. 33 and A23187 218. 219 and aequorin 224 and arsenazo Ill 218 and atomic absorption spectroscopy 216-218 and EGTA 212. 217. 226 and FCCP 218-219 and fluorescent indicators 224-229, 236235 and glycogen phosphorylase 230-233 and LaCI, 217, 221. 223 and Mn2+ inflow 233-235 and MnCI, 226, 234, 235 and spectrofluorometry 225-228. 234 cytoplasmic free 203, 224-229, 316 extracellular concentration and cell damage 6, I I. 34.96.97. 141, 142, I55 fluxes 219-223, 235 function in cell adhesion 3.4, I@-I I , 12, 16. 35. 62 hormone action on 202,203,215,216. 220. 226. 229 in cell attachment to substrata 282, 287 in culture media 257, 277. 303 in incubation media 141, 142. 155. 205

443

in mitochondria and endoplasmic reticulum 218-220 inflow 2 15, 220-223, 229-235 intracellular 6, 96, 132, 141 measurement of see Chapter 9 movement using 45Ca2+(non-steadystate) 219-220 movement using 4sCa2+(steadystate) 21 9-223 perfusate concentration I I , 31, 33, 34, 38, 62. 63 pools 215, 220 total cellular, measurement of 2 15-21 8 (Ca” + Mg2+) ATPase 215,218, 219. 220 Carbogen 24-25, 41, 152, 259, 297 Carbohydrates in culture media 256, 302-303 Carbon dioxide as buffering agent 24, 42, 155 I4CO2measurement of 1 6 6167 physiological role 4 2 4 3 , 164, 168 Carbon tetrachloride 77, 188, 191. 197 Carcinoma, hepatocellular 270 Catalase 193-194 Cell adhesion molecules (CAMS) 4, 269-270, 323 Cell counting in monolayers 277, 348 in suspensions 46, 47, 4 8 - 4 9 with coulter counter 49 Cell cycle 2 7 6 2 7 7 . 315 Cell-cell contacts heterotypic 260, 266. 2 7 6 2 7 5 , 32 1-325 homotypic 260, 266, 269-270, 319-321, 341 Cell-lines epithelial 237, 322-325 fibroblast 278. 324 hepatoma 237 Cell-matrix contacts 272-274. 278-280,287,3 13-3 14,322-324

444


Centrifugal elutriation 81,245,396,399 [3H]Choline 208, 209 CI. histolyticum 6, 8, 9, 38. 39 Co-culture of hepatocytes characteristics 274-275, 292-293, 320, 321-323 effects on hepatocyte function 269. 274-275, 276, 323-325 with endothelial cells 274, 323-325 with fibroblasts 324 with liver epithelial cells 274-275. 322-325. 344, 345 with reticulo-endothelial cells 276, 325 Collagen receptors 274, 280, 287 Collagen, type I cell attachment to 267,280,282-284 coating of culture dishes 279, 283, 285-286 effects in culture 267-269, 272, 276 in gels 269, 272, 280, 286-287 preparation (from rat tail) 284-285 synthesis and secretion 267,275,322 Collagen, type IV attachment to 280, 282 effects on cultured cells 272. 276 in extracellularmatrix 279,288-289 preparation 284 Collagenase and Ca2+ 8. I I , 33. 38 and digitonin perfusion 397-398 and Mg2+ 41 concentration in medium 25.37.249 crude 8, 9, 11, 37-38, 61 damage to receptors 61, 204, 240 mode of action 8, 242-244 purified 38 residual effects 61, 62,204,240,241. 252 sources 8, 37 specific 38-39 sterilization 248 Conditioned media 321, 325 Conjugation reactions 184-187

Connective tissue in liver 2. 243 Contents of isolated hepatocytes adenine nucleotides 95, 144-147 enzymes 5.78,90-91, 144,361,398 glycogen 24, 52, 77, 103-104, 109, 110, 147-148. 169-170 labile metabolites 40, 62, 131-132, 139-140 lipids 129, 144 Na+ and K+ ions 134, 137-138, 141-143 stablemetabolites 13&131. 144, 148, 149 water 50, 51-52, 97, 142, 387 Coulter counter 49 Covalent binding of proteins to drug metabolites 188, 198-199, 200 to glass coverslips 279, 283 to plastic 279. 283 Cryopreservation 395 Cu2+ 40, 58, 193, 271, 295, 303 Culture dishes bacteriological 261, 282, 283 plastic 261. 279. 282, 291 Primaria 261, 282, 345 Cyclic AMP assay 207 effect of hormones on 203,207-208 effect on glycogen phosphorylase 233 phosphodiesterase 204 Cytochrome P-450 binding of drugs to 180, 186 drug-induced spectral changes 180. I86 induction by pretreatment in vivo 76, 187 induction in culture 181-182, 273, 275, 308. 315-317 isozymes 181-182, 275, 317 levels in cell suspensions 181, 254 levels in cells in culture 62, 181-182, 240. 267. 317, 343. 346 Cytokinesis 277

SUBJECT INDEX Cytoskeletal proteins 102, 105. 107, 195, 267, 273 Cytoskeleton 267, 270, 274, 307, 316 Cytosol 357 Cytospheres biochemical characterization 365.37 I isolation reagents 365-3615, method of preparation 365-370 properties 369, 371 Damage, indicators of cellular see also Toxic damage to hepatocytes blebbing 99-100. 101, 102, 119, 19G195 choice of 85,83-88,93-98,189,381 enzyme leakage 88, 90-91, 92-93, 94, 189, 198, 361, 374 maximum percentage acceptable 47, 86 microscopic appearance 78, 84, 100-102, 106, 108, 111, 119 succinate oxidation 8 6 8 8 , 89-90, 94 trypan blue staining 46, 47, 85, 91, 93-94. 100 Damaged hepatocytes, preparation of 4, 77, 7 8 4 0 Damaged hepatocytes, removal of via selective attachment 246, 281 with dibutyl phthalate 190 with differential centrifugation 245 with Metrizamide 55-56 with Percoll 55, 56, 57.64, 240, 245, 246, 255, 258-259 with trypsin 55. 76 Density (number per ml) of hepatocytes for spheroid formation 261 in suspension incubation 49, 125-126, 151--154,206,256,259 Density (number per plate) of hepatocytes effects on function 266, 268-270, 3 19-32 I , 342, 343

445

mechanism of density effects 269-270, 321 plating methods 280-281.329-330. 332, 334-335, 343, 345 Density ( p ) of hepatocytes 55, 56, 245 of nonhepatocytes 245 Desmosomes see also Junctional complexes adhesive properties 10, 11 cleavage 10, I I , 16, 35 morphology 3, 10, 105 Development, hepatic 321-322, 337-342 Dexamethasone 210-21 I , 267, 280, 298, 306-308, 312, 326, 328, 330 see also Glucocorticoids; Hydrocortisone Diabetic animals, hepatocyte isolation from 77, 140-141, 147, 151 Diacylglycerol 20 I , 203, 208 Dietary treatment prior to hepatocyte isolation 7 6 7 7 Differentiation of hepatocytes 253, 266-275, 292, 299-309, 315-323, 337-342, 343-344 Digitonin action on cell membranes 87, 358 and collagenase perfusion 397-398 fractionation of hepatocytes 213, 358-361, 362-363 instability 88 permeabilization of hepatocytes 87-88, 89-93, 373, 375-376, 387, 390 Digitonin fractionation of hepatocytes alternatives to 213, 362-364 and autophagy 377 and intramitochondrial pH 390 cross-contamination of fractions 360, 361 nature of fractions 357 recovery of metabolites 360 technique 358-361

446


temperature-dependence 361, 362 Dimensions of hepatocytes 100, 244 of nonhepatocytes 244 of spheroids 260 Dimethylsulphoxide and cryopreservation 395 as a solvent 159, 328 as an antifreeze. 362 in media 271,277,316316,338,344 Dispase 8, 9, 38, 75, 239, 243 Disruption of hepatocytes by freezing with liquid nitrogen 136, 351-352 by sonication 351, 365 in culture 351-352 in suspension 351, 364-365 with detergents 90,92, 140,351-352, 362 with Dounce homogenizer 212, 351-352, 364 with Ultraturrax homogenizer 351-352, 365 DMSO see Dimethyl sulphoxide DNA damage and repair 241, 252 hepatocyte content 125, 127, 353 synthesis 275-277, 278, 3 1 6 3 1 3 , 315-317, 341, 344 Drug metabolism by cultured hepatocytes 181-182. 253-254, 267, 3 3 6 3 3 7 by human liver 187-188 by microsomal preparations 184-186 by perfused liver 183, 184-185 by periportal and perivenous (centrilobular) hepatocytes 180-181, 3 3 6 3 3 7 induction of enzymes of 185, 273, 275, 317 interspecies differences 187 NADPH requirement 186 phase I reactions 184-186

phase I1 reactions 184-186 rates of 184-187 variation with age and sex 187 Drug toxicity, indicators of see also Toxic damage to hepatocytes metabolic 188-190 morphological 194-195 Drug toxicity, mechanisms and measurement of and cell culture 266 covalent binding to macromolecules 198-199 ethanol 199-200 free-radical formation 192-194, 195 glutathione depletion 195-196 lipid peroxidation 1 9 6 1 9 8 solvent damage to membranes 188. 191-192 Drug uptake measurement of 182-1 83 mechanisms of transport 182-183 Drug-treated animals 76, 77, 181, 183. 187 Dual wavelength spectroscopy 21 8 EDTA [slC,]EDTA and Ca2+measurement 223 in hepatocyte isolation 4 , 3 9 4 0 , 6 3 . 240 in incubation media 210 EGF see Epidermal growth factor EGTA in hepatocyte isolation 35, 3 9 4 0 , 249 quenching of quin2 fluorescence 226 EHS (Engelbreth-Holm-Swarm) turnour 273, 287-289 Electron microscopic techniques critical point drying 120 cryo-immunoelectron microscopy 113-I 14 deep-etching 1 16, I I7

SUBJECT INDEX freeze-fracture I 1 4 - 1 15, 116, 369 glutaraldehyde fixation 112-1 I3 immuno- and lectin-labelling 108, 113-Il4, 117 osmium tetroxide I12 thin-sectioning I12 Electropermeabilization of hepatocytes membrane characteristics induced 377-378 method 376-377 Elutriation, centrifugal 8 I , 245, 396, 399 Embryonic liver 52.7k76.321-322, 337-340 Endocytosis 105. 114, 117 Endoplasmic reticulum and cell damage 6, 108, 184, 375 Ca2+ stores in 218-219, 373 in isolated hepatocyte 103, 108 microscopic appearance 104, 106, 109, I l l microsome preparation 185 Endothelial cells 2, 81, 242-245, 323-325 Entrained water in hepatocyte pellets markers for 133-135, 223 occurrence and measurement of 50. 123-1 24, 133- 134, 135-f37. 140, 360, 385 Enzyme induction 187.210-21 I , 254, 308 Enzyme leakage as an indicator of cellular damage see also Damage, indicators of cellular alanine aminotransferase 93 lactate dehydrogenase91, 92-93, 98. 189, 348, 361, 374, 375 Enzymes for cell isolation clostripain 8, 9 collagenase see separate entry dispase 9, 239, 243 enzyme mixtures 8. 37, 38, 243, 337, 344 hyaluronidase 6, 7. 8, 37. 204

447

lysozyme 7. 9. 76 pronase 7, 9. 80 trypsin 8. 9, 37. 76, 239. 243 Epidermal growth Factor cell response 202,203,261, 276, 309. 310, 344 in media 298. 300, 309. 314, 328, 330, 333 Epithelial cells. liver (clear. clonigenic) 81,240,243-245. 314. 323-325, 338 Erythrocytes, use of in perfusion 43. 204, 379 Erythrosin B 94, 378 Ethane 197-198 Ethanol as dehydrating agent 112. I19 as solubilizing agent 159 as sterilizing agent 23 as substrate 159. 199 in assay of phosphorylase 232-233 metabolism and toxicity 199-200 Evaporation in incubation vessel 153, 199. 216 Extracellular matrix 223, 260, 266, 272-274. 279. 307. 322 Extracellular volume see Entrained water in hepatocyte pellets Extracts of hepatocytes 129. /35--140. 178,207,217.23I-232,350-352 Fasting, effects of 95.97. 100, 125. 127. 141, 147, 151. 167. 169 Fat storing cells (lipocytes) 2, 81. 242-245 Fatty acids binding to albumin 156. 172-173 metabolism 171, 172. 378 removal from albumin 156 FCCP and Ca2+ measurement 218-219 Fibroblasts contamination with, in culture 243-245. 314, 338, 345

448


in co-culture 324 Fibronectin 272, 274, 276, 280, 287-289 Filters cellulose ester for cell culture 334-335 for measurement of covalent binding I98 for medium sterilization 248. 328 for perifusion 3 8 6 3 8 I, 382-383 gas-permeable, for cell culture 335 used during hepatocyte isolation 17. 31, 248 Fixation of hepatocytes 112--115, 119, 353-354 see also lmmunohistochemistry, fixation for Flasks, for suspension culture 255, 259 Foetal hepatocytes culture 337-340. 3-346 growth 301-302, 339-340, 345-356 isolation 7 6 7 6 , 337-338, 344-345 relative number 52, 75. 76 Foetalization 268 a-Foetoprotein 267, 273. 339, 345 Fractionation, subcellular 2 13, 352-353, 355-358, 359-360. 361-363, 365 Free-radicals and redox cycling 192-193 from quinones 192 glutathione depletion 195-196 hydrogen peroxide, formation of 193- I94 in monolayer culture 306 scavengers of 58. 198 spin traps for 192 superoxide 192- I93 Freeze-clamping 128, 363 Freeze-fracture 1 1 4 - 1 15, 1 16, 117, 369 Fructose metabolism 145. 147 Fungicides 305

Fura2 228-229,

233-235

Gap junctions see Junctional complexes Gas phase for cell incubations carbogen 152, 164-165, 167, 197, 206. 259, 297 oxygen 167, 197. 305-306 Gassing of perfusion medium 25, 4 1 4 3 , 247 Gassing manifold 31, 152 Gelatin 1 5 6 1 5 7 , 205, 213 Genetic disorders in hepatocyte donor animals 77 Glisson’s capsule 245, 323 Glucagon in culture media 262. 271, 298, 308, 3 12, 328, 330 receptors 95 responses to 95, 96, 175, 203, 204, 207-208, 276, 312, 336 Glucocorticoids (steroids) during attachment 280 in media 2 1 b 2 1 I, 257, 275, 306-308.312.3 14, 326,328,330, 338, 343, 345 receptors 307-308 responses to 174,203,210-211,276, 306-308, 312 Gluconeogenesis 95-96, 153, 155, 168-169, 203, 238, 267, 337 Glucose in culture media 302-303 in incubation media 24, 147, 154, 169. 170, 206, 256, 302-303 in perfusion media 204 metabolism see Gluconeogenesisand Glycolysis Glucose-I-phosphate 231, 232, 233 Glucose-6-phosphatase 5, 125, 267. 268. 317 Glutamine synthetase 337, 398 y-Glutamyl transpeptidase 62.240.267, 303, 308

SUBJECT INDEX Glutathione conjugation with 184, 187 drug-induced depletion 188, 193. 195-196 hepatocyte content 35.40, 102, 182. 188, 195, 240, 252 measurement of 195-1 96 oxidized and reduced 195-196 Glutathione-S-transferase 278. 3 I5 Glyceraldehyde 3-phosphate dehydrogenase 363 Glycerol metabolism 145, 147 labelled 208 Glycogen breakdown 24.52,96, 147, 169, 186, 203, 204 content in hepatocyte 147 microscopic 103, 104, 108, 109. 110, I12 phosphorylase 230-233 source 231 storage disease 77 synthesis 96, 169-170, 189,203,232, 238, 322 Glycogenolysis 96, 169, 203 Glycolysis 150, 154, 169 Glycosaminoglycans 8, 9. 269, 272, 313-314 Growth factors 276, 298, 309-3 11, 3 4 k 3 4 1 see also Epidermal growth factor; Growth hormone: Hepatopoietins; Insulin; Oestrogens; Prolactin; Transferrin; Transforming growth factors Growth hormone (somatotropin) 262, 298, 309. 312 Growth of hepatocytes in culture characteristics 276-277, 309-310, 339-342. 344 effects of cell density 268-270, 320-32 I , 340, 345 effects of media 276-277.299-300,

449

301-304, 309-313, 3 1 6 3 1 8 , 327, 339-342, 344, 345-346 effects of substrata 261,276,278,321 other requirements 241, 321. 325 procedures for studies of 331-333, 339-342, 344-346 Growth of liver nonhepatocytes in culture 242-244, 3 1 6 3 1 5 , 338-339, 345 Haematopoietic cells in liver 52, 75-76 Haemocytometer 46, 47. 4 8 4 9 , 125, 369 Haloalkanes 188, 191, 197 Heparan sulphate proteoglycan 3,283, 313 Heparin 39, 272, 311, 313-314 Hepatoblasts. mouse embryonic 338 Hepatocyte dimensions 100, 133, 244 Hepatocyte doublets chemical studies 396-397 electrophysiological studies 396 morphology 99, 100 Hepatocyte extracts see Extracts of hepatocytes Hepatocyte isolation, enzymatic methods development of 1-2, &8 for extended culture 239, 241-251, 248-251,337-338,340,3&345 from abnormal animals 7 6 7 7 from avian species 68-69. 347 from biopsy specimens 60, 70, 71 from cattle 60, 68 from caudate lobe 60, 67 from fish 69. 347 from foetal animals 7676,337-338 from guinea pigs 67 from humans 60, 70-72, 342-345 from mice 64-66, 74, 346 from monkeys 73 from neonatal animals 74-75, 340 from pigs 73, 74, 347

450

lSOLATED HEPATOCYTES

from rabbits 346 from rats 6 8 , 16-33, 74, 75, 7 6 7 7 , 204, 241-251 from sheep 60,67 from various species 60, 72-73 one-step procedure see One-step perfusion procedure two-step procedure see Two-step perfusion procedure Hepatocyte isolation, mechanical methods apparatus for 4, 5, 7 8 4 0 damaged hepatocytes, preparation of 6 6 , 12, 77, 78-80, 108, 1 I 1, Hepatocyte isolation, non-enzymatic methods perfusion using EDTA 6 1 4 2 , 63-44. 239-240 Hepatoma cell-lines 237 Hepatopoietins 276, 310 Hexobarbital 18I, 184 Hexokinase D (glucokinase) 169 Histology of liver cell numbers 2, 52, 76 cell types 2, 52, 75-76, 105 desmosomes 3, 10, 105 Abroconnective tissue 2 junctional complexes 2, 3, 103, 105, 106, 107 Histone acetylation 318 Homogenization of hepatocytes see Disruption of hepatocytes Hormone action and effects on enzymes 203. 210-211, 254, 308, 336, 340 hepatocyte sensitivity 204, 252, 253, 310, 311-312, 336 intracellular Ca2+203,216, 219,224, 226, 229 intracellular messengers 202,203,207 metabolism 96, 171, 174, 175, 203 mitochondria 211-213 Mn2+ inflow 234-235 Hormones in culture media see

individual hormones Human, adult hepatocytes culture 342-344 drug studies 187-188 isolation 60, 70-72, 342-343 Human, foetal hepatocytes culture 344-346 isolation 344-345 Hyaluronidase 6, 7, 8, 37. 204 Hydrocortisone 298, 3 0 6 3 0 8 , 325, 338,345 see also Dexamethasone; Glucocorticoids Hydrogen peroxide measurement 193- 194 Hypoxia consequences of 95, 100, 109, 130, 141, 144, 152, 167. 204 Immunohistochemistry, fixation for 113-114, 354 Incubation conditions for hepatocyte suspensions see also Protocols albumin, requirement for 40,58. 155-156, 205, 257, 258 antibiotics 210, 246, 258 buffering 155,157-158,205,257,258 Ca2+ 141, 142, 155, 205, 210 carbon dioxide 152, 155 gas mixture 152, 155, 167-168.206 gelatin, requirement for 156-157, 205 length of incubation 148, 149-150, 162,206, 207, 21 I, 252-255, 258 media 155, 205, 2 5 6 2 5 8 see also Media for metabolic studies with hepatocytes optimal quantity of cells SO, 51-52, 152-154, 206, 256, 259 osmolality 157-158, 205, 256 oxygenation of medium 57, I5 I , 152, 161, 162, 168, 206, 2 5 6 2 5 6 pH 155, 205 substrate concentration 154, 169, 172 substrates 148, 159, 168-176, 206, 256

SUBJECT INDEX temperature of incubation 151. 259 use of carbogen 152. 255 vessel dimensions I5 I. 152, 206, 207. 255 see u/so Incubation vessels volume 150. 151. 153--154.206,255 water bath, shaking 152, 206, 255 Incubation conditions for monolayer culture see Buffers; Co-culture of hepatocytes; Density (number per plate) of hepatocytes; Media for hepatocyte culture; Oxygen tension; Substrata for monolayer culture Incubation conditions for suspension culture buffers 257 incubation vessels 2 5 6 2 5 6 . 258 media 2 5 6 2 5 8 oxygenation 255. 259 shaking conditions 2 5 6 2 5 6 . 259 Incubation vessels chamber for perifusion 335-336, 380-382 chambers for multiple sampling 207, 216, 221 for measurement of oxygen uptake 161, 163, 164. 165 for metabolic studies 151-153. 206, 207. 210 for suspension culture 2 5 G 2 5 6 Inhibitors of gluconeogenesis 86. 143. 168-169 of glycogenphosphorylase phosphate phosphatase 230 of proteases 37, 175. 176, 202, 210 of respiration 97, 149, 390 use of 149, 156, 158-159, 387, 390 Inner mitochondria1 membrane potential measurement of 213, 386-388, 389 significance of 384. 389-390 [3H]lnositol 208-209 lnositol 1.43-trisphosphate 203, 373 Insulin

45 1

during cell attachment 280, 324, 329, 332 in cell isolation media 242 in culture media 257.261-262.267, 271. 298, 308, 311, 315, 317, 328, 330, 333, 343 receptors 202 responses to I7 I , 174. 175,202, 203, 209, 210, 276, 308, 311, 340, 343 Integrity of hepatocytes, metabolic measures of see also Damage, indicators of cellular ATP content I I , 44,58, 95, 98, 16147 glucagon, response to 95-96, 204, 205, 207-208 gluconeogenesis I I , 42. 58, 86, 95-96, 204 glycogen synthesis 96 maintenance of ion gradients 34,44, 9 6 9 7 , 141-143 protein synthesis 96, 148. 252, 255-256 respiration 5, 6, 57. 96 urea synthesis 95 Intracellular (cytoplasmic) free Ca’+ 203, 224-229, 3 I6 lntracellular messengers 203, 207 lntracellular water in hepatocyte pellets, measurement with tritiated water 133. 135-137. 140, 385 Inulin, radiolabelled and Ca2+ measurement 217. 222 decomposition of 134 in the measurement of intra- and extra-cellular water 131, 132, 133--134,135-137. 140, 178,385, 217 Ion gradients 34. 44, 9 6 9 7 . 141-143 transport 143. 1 7 6 1 7 7 . 219-223. 229-235, 3 9 6 3 9 7 Ion concentrations in culture media see individual ions

452


in isolated hepatocytes 34. 134, 137--138,141, 142,215,22&229, 252,385 lsozymes cytochrome P-450 181-182. 273, 275, 317 pyruvate kinase 267, 302, 318 Ito cells see Lipocytes Janus Green B 85 Junctional complexes cleavage of 10, 1 I , 12. 13. 16, 35.62, lOGI07 components of 3, 103, 104, 105, 269 re-establishment during culture 269-270. 313, 324

K+ concentration in hepatocytes 97, 134, 137-138. 142 loss from hepatocytes 34,44,9&97, 141, 143 Ketogenesis 171-172 Kupffer cells 2.8,52,80-8 I , 105, 174, 242-245. 302 LaCI, and Ca2+measurement 2 17,221, 223 Lactate dehydrogenase (LDH) activity in hepatocytes 92-93. 94, 267, 361 leakage 88.91.92-93, 189,207,348, 374, 375, 382 Lactate metabolism 148, 150, 153. 154. 168. 169 Lactate/pyruvate ratio 152. 153 Laminin 272, 274, 276, 279. 280, 287 Lineages, liver cell 244, 337-338 Lipid extraction 129 metabolism 171- I72 Lipid peroxidation detection of 196-198

effects of drugs 188, 191. 193, 196-198, 200 in culture 316 Lipocytes 2, 81, 242-245 Lipogenesis 168, 171, 308 Lipoproteins 108 Liver epithelial cells (clear, clonigenic) 240,243-245,314,317.323-325, 338-339 Liver perfusion see Perfused liver Liver regeneration 270, 336 Malate 144 Malate-aspartate shuttle 35, 148 Matrigel characteristics 273, 288-289, 353 effects in monolayer culture 272-274, 276, 292-294. 321 preparation 288-292 use as culture substratum 281, 288-289,291-292,327,32%-331 Mechanical methods of hepatocyte isolation see Hepatocyte isolation, mechanical methods Media for hepatocyte culture (commercially available) Chee’s medium 271, 294-298. 300-301, 326 Dulbecco-modified MEM:F-I2 293, 300, 303 Eagle‘s minimal essential medium (MEM) 256, 293, 299 F-12 medium 299 Leibovitz L-15 256, 293, 300 Medium 199 293, 299 RPMl 1640 293, 299 Waymouth media 256, 293, 299 Williams’ Medium E 293, 299, 304. 328, 333, 339 Media for hepatocyte culture (supplemented) lsoms RPCD 294-298, 299-300, 303. 304, 314, 315

SUBJECT INDEX Jefferson/DMEM:F- 12 294-298, 299, 300, 901 LanfordNME 294-298, 299, 327, 343 MX-13 294-298, 300, 304 Reid HDM 271,294-298,299-300. 303-304, 314 Media for hepatocyte culture, constituents albumin 257, 258, 295. 304, 329 amino acids 256, 294, 301-302 antibiotics 246, 261, 297, 305, 330. 333 arginine 294, 301-302. 314 ascorbate 296, 305 buffers 257, 297. 328 butyrate 316, 318, 338 Ca2+ 257, 277, 295, 303 carbohydrates 256, 302-303 Cu2+ ions 271, 295, 303 dexamethasone see separate entry DMSO 271,277,31+316, 338,344 EGF see separate entry ethanolamine 296, 304, 328, 330 glucagon see separate entry glucocorticoids see separate entry glucose 256, 2%. 302-303 growth hormone (somatotropin) 262, 298, 309, 312 heparin 272, 311. 313-314 hormones see individual hormones hydrocortisone see separate entry insulin see separate entry ion concentrations see individual ions iron salts 295, 304 linoleic acid 271, 296, 304, 329. 330 nicotinamide 297, 3 17-3 I8 oestrogens 276, 31 1-312 ornithine 294, 302, 314, 338 pH 256, 328 phenobarbital 3 1 6 3 1 7

453

prolactin 262, 298, 309, 312 proline 294, 301, 339, 342 protease inhibitors 293 pyruvate 256. 296, 302, 312 selenium see separate entry serum see separate entry thyroid hormones 309, 342 trace elements 295, 303-304 transferrin 295, 304, 328, 330 vitamins 296-297, 305 Zn2+ ions 271, 295, 303-304 Media for metabolic studies with hepatocytes see also Incubation conditions for hepatocyte suspensions Eurocollins 72 Hanks 41, 59 Krebs bicarbonate saline 4 1 4 2 , 155. 256 phosphate buffered saline 24-25,347 Waymouth 205, 210 Membrane filtration of media see Filters Menadione 101, 193, 1 9 L 1 9 5 Metabolic activity of hepatocytes. basis for expressing cell dry weight 122-124, 127 cell number 125-126, 127, 353 choice of 126 conversion factors 127 DNA content 125, 127, 353 in terms of whole liver 123-124 PCA or TCA dry weight 122--123, 124, 126, 127 protein content 125, 127, 353 Metabolism see individual metabolites Metabolite-indicatormethods 363-364 Metabolites extracellular 40, 130, 131-132, 144 intraceilular 128, 130, 131-132,

454


137-140. 144-145, 3 5 6 3 5 8 , 358-360 leakage of 144, 372, 376 Metrizamide 55-56 Metyrapone 18 1 Microcarrier culture (cytodex beads) 333-334 Microcentrifuges I30 Microencapsulation of hepatocytes 262-263. 393 Microscopy of hepatocytes interference contrast 100, 101, 102 light 84, 99, 100 scanning electron 117-120 transmission electron 103-1 17 Microsomes 184-186. 353 Mitochondria and Ca2+ 6, 218-219 and respiration 6, 86. 87, 96 cholesterol content of membranes 358 enzymes of 90, 93, 214, 361 membrane potential 213, 386-390 morphology of 106, 108-1 I I pH of 390, 391 preparation from hepatocytes 21 1-214, 352-353 succinate oxidation by 6, 8 6 8 8 . 88-90, 94, 212 swelling 6, 108-109, I 1 1 Mitosis 275-277, 303, 341 Mn2+ inflow 233-235 MnCl2, quenching of quin2 fluorescence 226, 234-235 Monolayer culture of hepatocytes see Chapter I 1 see UINJ Attachment. of hepatocytes for monolayer culture; Co-culture of hepatocytes; Density of hepatocytes, Differentiation of hepatocytes; Growth of hepatocytes in culture; Media for hepatocyte culture; Substrata for monolayer culture Morphological indications of damage see Damage, indicators of cellular

Morphology of cultured hepatocytes after isolqtion with EDTA 240 effect of media 3 0 6 3 0 7 . 313, 315, 316318 in co-culture 322-323 in media with DMSO 315 in spheroids 260 on collagen films 267, 336 on collagen gels 269 on matrigel 273, 288, 331 Morphology of intact hepatocytes in suspension 99-108, 109, 110, 117-1 19, 253,260 Morphology of intact liver 103-108, I I7 see also Histology of liver Mouse hepatocytes culture 300, 346 gene transcription 308 growth 340, 346 isolation 5, -66. 74 use in drug studies 187 mRNA albumin 271,272,275,307.313.315, 339 cytoskeletal proteins 253, 267, 268, 272, 273, 313 turnover 271, 272, 307, 315 Na+ concentration in hepatocytes 96. 134, 137-138, 142 effects of ouabain 143 entry into liver cell 34, 44,96-97, 141, 143 NAD/NADH ratio 364 NADH 187, 192-193 NADPH 186 NaF 230 Neonatal hepatocytes culture 26-262. 266, 340-342 growth 277, 299, 309, 321 isolation 74-76, 340 Neutralization of acidified hepatocyte extracts 129-131 Nicotinamide 297, 3 17-3 I8

SUBJECT INDEX

455

NMR 364 Non-adherent culture of hepatocytes microencapsulated 262-263 spheroids 256. 260-262 suspensions 252-260 Non-hepatocytes (non-parenchymal cells) co-culture of hepatocytes with 260. 323-325 contamination of cultures with 242-245, 260, 3 14-3 15. 338-339. 340. 345 density ( p ) 245 dimensions 244 in liver 2, 52, 123. 242-243 isolation 80-81, 242-245 removal during cell isolation 31-32. 56, 243-245 Nuclei counting 277 isolation 352 Number of hepdtocytes in liver in relation to age 52-53 in relation to nutritional state 50. 52. I25 per gram liver 52-53, 123. 126, 127

reagents 23-25. 36-43 surgery 25-30, 43 swelling of lider 30, 32. 65-66 temperature control 17. 20-2 I. 23. 41145. 204 water quality 24. 40. 54 Osmolality 157-1 58,205,206,256.37 I Ouabain 143. 183. 381 Oxygen in gas phase during incubation 152. 197. 305-306 rate of consumption 151. 161. 203. 378 requirement in gas phase during perfusion 25. 4 1 4 3 Oxygen tension in hepatocyte suspensions 152. 162. 167-168. 206, 255. 305-306 in liver lobule 167. 306 in monolayer cultures 305-306.333 in perfusion media 25 Oxygen uptake, measurement of manometric 162. 164-166 polarographic 161-162, 163. 383, 389 with an oxystat 167-168

Oestrogens 276, 3 I 1-3 13 One-step perfusion procedure and collagenase 25. 30 apparatus 17-23 buffers for 4 1 4 3 Ca2+ion replacement 31, 34.36, 141 cannulation of vessels 26-30 cleansing of apparatus 22-23 hepatocyte washing 31-32, -5 in cell isolation for extended culture 24 I modifications to 36-45. 75, 204 oxygenation 17. 20, 21, 204 perfusion, duration of 3 0 . 3 3 . 4 3 A . 204 perfusion, flow-rate 25. 27, 29 perfusion pressure 44 post-perfusion incubation 31

Palmitate metabolism 378 PCA dry weight 122 extraction of hepatocytes 130. I3 I , 217. 222. 359-360 Percoll 55-57, 61. 64. 98. 190, 240. 246. 255. 258-259. 353. 366370, 372. 398 Perfused liver comparison with hepatocytes 143. 151. 171. 174. 183. 184-185.201. 209. 301, 390 Perfusion see One- o r Two-step perfusion procedure Perfusion medium, composition albumin 40. 249 buffers 24. 32. 4 1 4 3 . 78. 249

456


EDTA 4, 3 9 4 0 , 63, 240 EGTA 35, 39-40, 249 erythrocytes 43, 204, 379 ions 23-25, 41 metabolites 40, 204 Mg2+ ions 41 nutrients in 242. 248-249 pH 24, 4 1 4 2 Perifusion of hepatocytes in monolayer 335-336 in suspension 377-38 I , 380, 382-383. 384 Periportal and perivenous (centrilobular) hepatocytes collagenase digitonin perfusion 336. 397-398 culture 3 3 6 3 3 7 differential drug metabolism 18&181, 3 3 6 3 3 7 localization of glutamine synthase 337, 398 preparation of 336, 397-399 Permeabilization of hepatocytes chemical 87,88,354,373-374,374. 375-376 electrical 176, 373, 3 7 6 3 7 8 hepatocyte damage during 87. 373. 375. 376 pH of media for hepatocyte isolation 5.24.41-42, 249 for hepatocyte suspensions 155, 174. 205. 256, 257, 259-260 for monolayer culture 328, 333 pH, intracellular measurement 390-391 values for 390 Phalloidin 183, 195 Phenobarbital enzyme induction 76, 185. 187, 273, 3 15-3 17 in media 3 I 6 3 I7 induction of cytochrome P-450 181, 187, 273. 315-317

Phenylephrine 216, 229 Phosphatidylcholine 208' Phosphoenolpyruvate carboxykinase 203, 208, 210 Phosphorylation potential 363-364, 390 Pinocytosis 135, 228, 376 Plasma membrane blebbing of 42, 100, 101, 102, 111, 117, 119, 1 9 6 1 % Ca2+ movement across 215, 229-235 cholesterol content of 358 enzymatic, oligosaccharide and immunologic markers 113-1 14 junctions see Junctional complexes microvilli 107, 117-1 19. 253 permeabilization, chemical 87, 88, 373-374. 374. 375-376 permeabilization. electro- 176, 373, 376378 potential 85. 86, 384-385 preparation of 352. 372 receptors 61, 95-96, 202, 204, 209. 223, 252 transport functions 9 6 9 7 . 108, 176178 Plastic culture dishes 261, 279, 282, 291 vials 206, 207, 208, 225, 255 Plating conditions for monolayer culture 281, 319-321, 324-325. 329-330, 332, 343 see also Attachment of hepatocytes for monolayer culture Ploidy 52. 125, 126,244,276,341-342 Pluronic acid 228 Polarity of hepatocytes 105, 107, 108. 180, 253 Polysomes 148, 256 Post-transcriptional contro? 271. 272, 302, 313 Potassium ions see K + Primaria dishes 261. 282, 345

SUBJECT INDEX Primate hepatocytes culture 342-346 isolation 60,70-72, 73, 342-345 Progenitors of hepatocytes 244. 337-339

Prolactin 262, 298, 309, 312 Pronase 7, 9, 80 Prostaglandins 31 I Protease effects on hormone receptors 204 inhibitors 37, 293 Protein see also Cytoskeletal proteins; Covalent binding of proteins assay 127, 353 content of hepatocytes 122, 125, 127 precipitation 122, 129, 174, 207 Protein degradation hormone action on 175,202,203,210 in hepatocytes 17G176, 377. 256 Protein kinase C 274 Protein synthesis as an indicator of cell damage 189 hormone action on 174, 203, 210-211, 301, 307-309

in hepatocyte suspensions 125, 173-174.

255-256, 279

in monolayer cultures 279,301-302. 307-309,

320

Protein-tyrosine kinase 202, 210 Proteoglycans 3, 269, 272. 279, 288, 3 13-3 I 4

Protooncogene expression 253, 303 Pyruvate carboxylation 203 in media 204, 256, 302, 312 Quality of hepatocytes, causes of poor I I , 42, 54-55, 77, 204 Quin2 2 16, 224-228. 233-235 Quinones 181, 192-195 Rat liver cellular composition of 2, 52

457

hepatocyte number 50, 52-53, 125- I26

hepatocyte ploidy 52, 125, 126 hepatocyte volume 50, 52, 123 Rats as hepatocyte donors age 52, 125-126. 241, 341, 342 weight 16, 53, 121, 125-126, 241 Receptors a and B adrenergic 202, 203, 31 I collagens 274, 279, 287 damage to 202, 204, 209, 252, 253 EGF 202, 209, 318 fibronectin 274, 279, 287 glucagon 95, 96, 204 glucocorticoid 307-308 insulin 202, 209 laminin 274, 279, 287 thyroid hormones 309 Redox cycling 192-193 Release of hepatocytes from culture substrata 348-350 Respiration see Integrity of hepatocytes, metabolic measures of Respiratory control ratio 212, 213 Reticulo-endothelial cells 84 see also Endothelial cells; Kupffer cells Retrograde perfusion 65-69, 75, 397-398

RNA 173, 176, 199 Sandwich configuration for culture 272 Saponin 354, 373-375 Scanning microscopy 117-120 Selective media 302, 307,3 1 6 315,338 Selenium in culture media 271,295,304. 330, 333

Serum effects during attachment 261, 279, 345

effects on differentiation 293, 338 effects on growth 293, 310 growth factors in 293

458


in culture media 257, 261, 293. 310, 314. 336, 338. 345 Serum protein see also Albumin mRNA levels 271 synthesis and secretion 254. 263,271 Silicone oils, use in separation procedures 131, 139-140, 359. 190, 221-223 Sodium ions see Na+ Solvent damage 159, 191-192 Soybean trypsin inhibitor 37 Spectrofluorometry 225, 234 Spheroids 256, 260-262. 275, 282 Spin traps 192 Spinner cultures 2 5 6 2 5 6 , 334 Stem cells in liver 244, 302 Sterility 22-23, 2 4 6 2 4 8 , 284-285, 290-291, 328 Steroid hormone action 210. 306308. 311-312 Storage of hepatocytes cryopreservation 395 long-term 58 metabolism after 57-58, 395 short-term 57-58 Subcellular fractionation of hepatocytes 359-360, 352-353, 355-358, 361-363, 365 Subcellular structure see Morphology of intact liver or Histology of liver Substrata for monolayer culture 243, 267,272-274.278-293,321,329, 332,333-335,337-338,340,345 Succinate oxidation and iodonitrotetrazolium 88 by hepatocytes 5,8688,89-9&94 mitochondria1 212 Sucrose hepatocyte permeability to 134-135, 375, 376, 377 in fractionation media 140, 358, 364 in isolation media 5, 78-80, 365-369

Superoxide 192- I93 Surgery, for isolated hepatocyte preparation from cattle, goats or sheep 67-68 from chickens 68-69 from foetal rats 7 6 7 5 from humans 70-72 from mice -66 from rats 25-30. 43, 247, 250 Suspension culture of hepatocytes cell density 256 comparison with monolayers 252-254 hepatocyte isolation 241-251. 250-25 I media 2 5 6 2 5 8 procedures for 257-260 shaking conditions 254-256 spinner culture 2 5 6 2 5 6 SV-40 transformed hepatocytes 299. 300 TCA dry weight 122. 124 Temperature for cell incubation 151 for cell storage 57-58. 97 of perfusion medium 5. 17, 54, 249 of washing medium 3 1 . 4 4 - 4 5 . 147, 249, 251 Thyroid disorders in hepatocyte donor animals 77 Thyroid hormones 309, 342 Tight junctions see Junctional complexes Tissue slices from liver as a source of isolated hepatocytes 4, 7. 59-60, 61, 67, 75 as an experimental system 14. 143, 151 gene expression 272 Toluene in cell permeabilization 191 toxic effect 188 Toxic damage to hepatocytes 156.2 19,

SUBJECT INDEX 224 SPP also Drug toxicity, indicators of; Drug toxicity. mechanisms and measurement of Trace elements in media 295.303-304 Transcription 268,271-275.307-308, 313, 316, 322 Transferrin in culture medium 295, 304, 328, 330. 333 receptors 304 Transforming growth factors 276. 310-31 1, 340 Transplantation of hepatocytes as treatment for hepatic failure 394 cryopreservation in 395 hepatocyte survival after 392, 394 measurement of function after 393-394 microencapsulation for 262 procedure 392-393 recognition of transplanted cells 393-394 Triphenylmethylphosphonium ion (TPMP') 384. 386-389 Triton X-100 81, 90, 92. 140, 226, 361 Trypan blue acceptable percentage of cells staining with 47, 86, 97, 246 desired concentration 46 effect of pH 46 haemocytometer, use of 4 6 4 7 preparation of solution 46 technique for using 46. 47, 93-94, 348 Trypsin 6. 8. 9, 37, 55, 76, 239. 243 Two-step perfusion procedure duration and flow rate 32-33,35,44, 250 HEPES buffering 32. 42, 249 in cell isolation for extended culture 241-251, 248-251 in standard cell isolation 32-33, 33-36, 40, 42 Mg2+ omission from medium 41

459

non-recirculating perfusion 32-33, 35. 36 oxygenation of media 42, 247 pre-perfusion 10. 32, 35. 36, 40 role of CaZf 1&11, 33. 34, 35, 36, 38 surgery 43 under sterile conditions 246-251 Tyrosine amino transferase induction 210-211, 241, 261, 307, 340 Urea synthesis 95, 168, 170-171, 176, 336. 337 Valinomycin 143, 151, 389 Vasopressin 203, 226, 229-232, 234, 312 Viability 84, 245-246, 255, 347-348 see ulso Integrity of hepatocytes, metabolic measures of Washing of glass-, plastic-ware 54, 258 hepatocytes 25, 31-32. 44-45, 56-57, 6 3 4 4 , 80, 91, 147. 204, 217, 245, 249, 25I, 395 Water content of hepatocytes 52, 133, 135-137, 14&142 entrained in hepatocyte pellet 50, 135-137. 123-124, 133-134, 140, 360, 385 purification and purity 24,40,54,328 quality 54, 328 Water, tritiated, use of for measurement of cell water 133, 135-137, 385 for measurement of lipogenesis 171 Xenobiotics 254. 266 see Drug metabolism; Drug toxicity, indicators of; Drug toxicity, mechanisms and measurement; Drug uptake

460


Yield of hepatocytes as a percentage of liver weight 52-53 comparison of one- and two-step procedures 33, 38 for determining desired quantity of cells in a suspension 50, 51-52, 151

from foetal or neonatal animals 74-76 from human liver 71

from livers of abnormal animals 77 from mouse liver 65-66 from rat liver 39, 52-53, 64, 258 measured by PCA or TCA dry weight 122-123, 123-124 measured by wet weight of hepatocyte pellet 50, 51-52, 123-124 using EDTA perfusion 64, 240 Zn2+ 295, 303-304

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