Biological Specimen Preparation for Transmission Electron Microscopy [Core Textbook ed.] 9781400865024

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Practical Methods in Electron Microscopy

Volume I 7

Titles in t h e Practical M e t h o d s i n Electron Microscopy series

Volume 15

Vacuum Methods in Electron Microscopy by Wilbur C. Bigelow

1994

In this volume the characteristics of common vacuum devices are described and instructions are given for operating the many different types of vacuum systems found in electron microscope and other scientific laboratories. This book is essential reading for all scientists, engineers, and technicians who operate and maintain vacuum systems on electron microscopes and similar scientific equipment.

Volume 16

X-ray Microanalysis for Biologists by Alice Warley

1997

In this volume the theory of X-ray microanalysis with the electron microscope and the instruments used are fully described. There is also a wealth of practical advice on methods for the preparation of biological specimens and their qualitative and quantitative analysis. This book is essential reading for all biologists who are using, or who wish to use, the techniques of X-ray microanalysis.

Volume 17

Biological Specimen Preparation for Transmission Electron Microscopy by Audrey M . Glauert and Peter R, Lewis

1998

Practical M e t h o d s in Electron Microscopy Volume I 7

Series editor:

Audrey M. Glauert Clare Hall University of Cambridge U.K.

Princeton University Press Princeton, NewJersey

Biological Specimen Preparation for Transmission Electron Microscopy

Audrey M. Glauert Clare Hall University o f Cambridge U.K.

Peter R. Lewis Physiological Laboratory University o f Cambridge U.K.

Princeton University Press Princeton, NewJersey

Published in North America by Princeton University Press, 41 William Street, Princeton, NewJersey 08540, U.S.A. Published in the United Kingdom by Portland Press Ltd, 59 Portland Place, London WIN 3AJ, U.K. Tel: (+44) 171 580 5530; e-mail: [email protected] Copyright © 1998 Portland Press Ltd, London

Library of Congress Catalog Card Number ISBN

98-88115

0-691-00749-7 (cloth) 0-691-00900-7 (pbk)

All rights reserved

Although, at the time of going to press, the information contained in this publication is believed to be correct, neither the authors, the editor nor the publisher assumes any responsibility for any errors or omissions herein contained. Opinions expressed in this book are those of the authors and are not necessarily held by the editor or the publishers.

Typeset by Portland Press Ltd Printed in Great Britain by the University Press, Cambridge, U.K. http://pup.princeton.edu 10 9 8 7 6 5 4 3 2 1

Editor's preface to the series

Electron microscopy is a fundamental technique with wide applications in all branches of science and technology. Every year large numbers of students, technicians and research workers start to use the electron microscope and need an introduction to the instrument, and to the many and varied methods for the preparation of specimens. Many books have been published which describe the techniques of electron microscopy in general terms, but when this series was originally conceived, there was an urgent need for a set of practical laboratory handbooks in which these techniques were described in sufficient detail to enable the isolated worker to carry them out successfully. Now, over 25 years later, Practical Methods in Electron Microscopy has gained an international reputation as the unique source of practical information for all electron microscopists and is still fulfilling its original purpose. This achievement largely results from the fact that all the authors are recognized experts in their particular fields and are thus able to describe the techniques of electron microscopy in sufficient detail for them to be understood and applied accurately by beginners and experienced microscopists alike. In contrast, certain single author texts cease to be of any practical value once the authors move outside the field of their own practical experience. Perhaps the best compliment that Practical Methods in Electron Microscopy has received is a statement in a review of one book that "The book leaves the reader with the impression of not having read a textbook, but having talked to a practising microscopist". Each book in the series starts from first principles, assuming no specialist knowledge, and is complete in itself. The books are designed to be guides through the often bewildering choice of methods that are available, and every attempt is made to simplify this choice. Consequently, in general, only well-established methods which have been used successfully outside their laboratory of origin are described in detail. We do not provide descriptions of the latest exciting 'break through', although each book does indicate the most promising of developments for the future.

W/

viii

Editor's preface

The present volume is the 17th in the series, and the high standards attained by the previous volumes have been maintained. I look forward to a future in which authors, editor and publisher continue to contribute to the science and art of electron microscopy. Audrey M. Glauert, Sc.D. Series Editor, 1998 Clare Hall University of Cambridge

Authors' preface and acknowledgements

Preface

Most biological material can be examined in the transmission electron microscope only in the form of ultrathin resin sections. There is a series of well-defined steps in the method for the production of embedded specimens suitable for sectioning, from initial fixation, through dehydration and infiltration with resin, to hardening of the final block. The aims of this book are to describe these procedures in detail and to provide practical advice for both the novice and the expert in transmission electron microscopy to enable them to obtain the optimum preservation of ultrastructure. Methods for cutting and staining ultrathin sections are fully described by Norma Reid and Julian Beesley, and by Peter Lewis and David Knight in companion books in this series. There is a bewildering variety of published methods for processing biological material and every attempt is made here to enable an informed choice to be made when examining a particular type of specimen. Consequently a detailed analysis of the molecular events accompanying fixation and embedding is included for the first time in a textbook of this kind. We have brought together data from a wide variety of sources and have been particularly reliant on the advice of polymer chemists for the analysis of the properties of embedding resins. This information provides the necessary basis for explaining why particular formulations and conditions for specimen preparation are specifically recommended. It is also aimed at helping readers to amend established techniques to suit their individual requirements. The references to published work have been selected with the same objectives in mind. In 1975 one of us (AMG) published a volume in this series entitled 'Fixation, dehydration and embedding of biological specimens'. The present book is much more than merely a second edition. The whole book has been completely rewritten and many new fixatives and

χ

Authors' preface a n d acknowledgements

embedding resins are described, together with methods for processing specimens at low temperatures. The overall format of the book follows that of the Practical Methods in Electron Microscopy series, with the emphasis on providing full practical details of dependable fixation and embedding techniques for the beginner and the experienced microscopist alike.

Acknowledgements

We are extremely grateful for the valuable and friendly advice we have received during the preparation of this book from electron microscopists throughout the world. They are too many to mention individually here, but we particularly wish to thank Eric Carlemalm (of the Biozentrum, Basel), Roy Gillett (of the London Resin Company), John Gray (formerly with Ted Pella), Peter Gerrits (of the University of Groningen) and Dick Martin (of Ciba-Geigy at Duxford) for providing the basic information on the molecular structure and properties of embedding resins, which makes such a valuable contribution to this book. We are especially grateful to Karen Klomparens (of Michigan State University) for her valuable comments on the manuscript at an early stage, and also to Jeremy Skepper (of the University of Cambridge) for all his advice, criticism and encouragement over many years, and for his willing and practical help in testing new fixatives and embedding media. Many people have very kindly provided electron micrographs and given permission for their inclusion in this book and we wish to thank the following: Joan Bain, John Bozzola, Ludwig Edelmann, Peter Eggli, Bruce Felgenhauer, J. Friihling, Peter Gerrits, Frederick Grinnell, C.G. Groot, Brian Gunning, C.E. Hulstaert, Ernst Hunziker, D. Kalicharan, Andrew Kent, Herbert Kuhn, Fiilton Mollenhauer, Jorge Moreira, W.F. Neiss, A.M. Page, Mary Reedy, Jeremy Skepper, Paul Strausbauch, P.J. Warren, Christel Westphal-Frosch, Roderick Woods and Werner Villiger. We are also grateful to Siegfried Bohler (of Bal-Tec), Steve Cham (of Leica), Lynne Joyce (of Agar Scientific), Christel Pella (of Ted Pella), Jeremy Skepper (of the University of Cambridge) and Robert Young (of the University of Wales at Cardiff) for providing photographs of apparatus and equipment.

Authors' preface and acknowledgements

We thank the following publishers for permission to use copyright material: the Company of Biologists (Figs. 1.1, 3.11 and 3.12), Elsevier Science (Figs. 9.1, 9.2 and 9.3), Gustav Fischer Verlag (Fig. 10.2), the Histochemical Society (Fig. 5.6), Jones & BartIett (Figs. 1.4, 3.2 and 3.9), Rolston Gordon Communications (Figs. 7.4 and 10.5), the Rockefeller University Press (Fig. 5.9), the Royal Microscopical Society (Figs. 3.14, 8.5, 8.8, 8.9 and 10.4), Scanning Microscopy International (Fig. 8.7), the Societe Francaise des Microscopies (Fig. 4.3), SpringerVerlag (Figs. 3.16 and 7.5) and John Wiley (Figs. 3.13, 4.2, 6.5, 6.7 and 10.3). We wish to thank Ron Parker for preparing some of the line drawings, Austin Hockaday and Peter Starling for help with the photography, Jo Leeds for assistance with some of the typing and Joyce Lewis for all her help and encouragement.

Audrey M. Glauert and Peter R. Lewis Clare Hall and the Physiological Laboratory University 7 of Cambridge

xi

Contents

xvii

Contents

1

Editor's preface to the series

vii

Authors' preface and acknowledgements

ix

An introduction to fixation and embedding procedures

I

and their safe use in the laboratory 1.1

The scope of this book

I

1.2

Criteria for the good preservation of ultrastructure

4

1.3

Artefacts in electron micrographs

6

1.4

Safety precautions in the electron microscope laboratory

9

1.4.1

General safety precautions

1.4.2

Hazards from electrical equipment and fire

10

9

1.4.3

Safe procedures for handling animals

11

1.4.4

Dangers from sharp implements

12

1.4.5

Safe procedures with chemicals

12

1.4.5a Labelling for toxicity and flammability

12

1.4.5b The safe handling of chemicals

15

1.4.5c

16

Storage and disposal of chemicals

1.4.6

Hazards with liquid nitrogen

17

1.4.7

First-aid in the laboratory

18

2

Fixatives

21

2.1

An introduction to fixation

21

2.1.1

The development of fixatives for electron microscopy

22

2.1.2

The chemistry and physics of fixation

24

2.1.3

The penetration of fixatives

25

2.1.4

The choice of aldehyde fixative

2.2

27

The safe preparation and disposal of fixatives

29

2.2.1

Preparation of a fixative

30

2.2.2

Disposal of fixatives

32

2.2.3

Spillages

33

Xiv

Contents

2.3

2.4

Vehicles for fixatives

33

2.3.1

The choice of buffer

34

2.3.2

Osmolarity

35

2.3.3

The preparation of buffers

37

Glutaraldehyde fixatives

38

2.4.1

The effects of glutaraldehyde on cells and tissues

40

2.4.1a Reactionsofglutaraldehydewithproteins

40

2.4.1b Reactions of glutaraldehyde with other cell constituents

44

2.4. Ic 2.4.2 2.5

The effects of the osmolarity of glutaraldehyde fixatives

The preparation of glutaraldehyde fixatives

44 45

Formaldehyde fixatives

46

2.5.1

The effects of formaldehyde on cells and tissues

41

2.5.2

The preparation of formaldehyde fixatives

50

2.5.2a A phosphate-buffered formaldehyde fixative

51

2.5.2b A cacodylate-buffered formaldehyde fixative

51

2.5.2c A PIPES-buffered formaldehyde fixative

52

Glutaraldehyde—formaldehyde fixatives

52

2.5.3

2.5.3a The preparation of a glutaraldehyde-formaldehyde fixative 2.5.4 2.6

Additives to aldehyde fixatives

53 53

Osmium tetroxide fixatives

55

2.6.1

The effects of osmium tetroxide fixation

56

2.6.2

Routine post-fixation with osmium tetroxide

58

2.6.3

Osmium tetroxide and complex cyanides

60

2.6.4

Amine complexes with osmium

61

2.6.5

Other osmiumC = C< double bond. Note that the aldol condensate formed possesses a free aldehyde group which can form a bridge to a second protein chain. The condensate is able to combine with another molecule of glutaraldehvde to form a pyridine derivative with the uptake of molecular oxygen via a complex series of reactions which are not shown here, but are discussed in detail bv Johnson (1987). Note that the pyridine derivative possesses additional aldehyde groups available for cross-linkages.

Chapter 2:

Fixatives

inhibitory effect on this aspect of fixation, and the use of hydrogen peroxide could be very important as a source of oxygen. The uptake of oxygen makes these reactions irreversible, which may explain the very stable fixation produced by glutaraldehyde as opposed to formaldehyde. Even if these reactions do not go as far as the oxidation to the pyridine derivatives, copious cross-linking can occur as a result of the aldol condensation reaction referred to above, which adds further aldehyde groups to the cyclic immine structure first formed. The net effect of all these possible reactions is that during glutar­ aldehyde fixation, extensive, multiple, stable cross-bridges are formed between amino groups with about two molecules of glutaraldehyde being consumed for each reactive amino group. The more oxygen available, the more cyclic immines will be converted to the more stable pyridine derivatives. The uptake of oxygen can produce anoxic conditions in the depths of large blocks of tissue, which could explain the beneficial effects of hydrogen peroxide reported by Peracchia and Mittler (1972). It is therefore prudent to use only thin slices of tissue when fixing by immersion with glutaraldehyde alone. The problem does not arise with a mixed fixative, since the formaldehyde diffuses in more rapidly and also largely suppresses the uptake of oxygen, although this does not mean that pyridine derivatives are not formed since the formaldehyde may act as an oxidizing agent. Precise data are missing on this point, however, so one must accept the possibility that the course of fixation is different and may be less complete when formaldehyde is included with the glutaraldehyde, although ultrastructural preservation appears not to be impaired. Since all the reactions between an aldehyde and a free amino group lead to the net production of acid, adequate buffering power is essential. Phosphate, cacodylate and PIPES all have ρK values below 7.0 and are therefore able to take up adequate amounts of acid without the pH falling too much. The first two are also small enough to diffuse through the extracellular spaces of a tissue faster than glutaraldehyde. All three buffers, however, carry a net negative charge at physiological pH values and probably diffuse through cell membranes more slowly than the uncharged molecules of the fixative. Hence the cell interior may become acid during fixation, and we do not know what effect this may have on the morphology of cytoplasmic organelles and inclusions. In order to minimize the possible adverse effects of a fall in pH during fixation it is desirable that the fixing

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Biological specimen preparation for TEM

solution should contain as high a concentration of buffer as is consistent with other considerations, such as the overall osmolarity. In summary, therefore, glutaraldehyde undergoes a series of reactions with free amino groups in proteins which results in rapid cross-linking to form a three-dimensional network throughout the cell cytoplasm. A substantial network is formed within a few minutes, even with a low concentration of glutaraldehyde, but the process probably continues for much longer. Uptake of oxygen and release of hydrogen ions both occur during the formation of the cross-links and should be taken into account in deciding upon the composition of the fixing solution and the conditions of fixation. 2.4. Ib

Reactions o f glutaraldehyde with other cell constituents

The role of glutaraldehyde in the fixation of other cell components is more problematical. DNA is certainly well fixed, but this is probably due to the fact that the aldehyde reacts avidly with the histones associated with the DNA. Direct reaction with the DNA molecules themselves appears not to occur at room temperature and the same is probably true for RNA. A major proportion of glycogen is retained by glutaraldehyde fixation, possibly as a result of the glycogen granules becoming enmeshed in a cross-linked matrix of protein. Glutaraldehyde reacts with some lipids, such as phosphatidylethanolamine, which carry a free amino or imino group, but there is little direct evidence for this from electron microscopy. Glutaraldehyde also reacts with the protein components of the lipoproteins which form important constituents of the overall structure of cell membranes, but this can cause the release of the lipid components. Glutaraldehyde fixation alone appears not to protect most lipids from extraction during the standard dehydration and embedding procedure. To preserve lipids secondary fixation with osmium tetroxide followed by uranyl acetate is necessary. Only then is good preservation of membrane ultrastructure obtained. 2.4.1c

T h e effects o f t h e osmolarity o f glutaraldehyde fixatives

It is important to stress that glutaraldehyde does not render cell membranes fully permeable to all the constituents of the cell cytoplasm or of external media. The overall composition and osmolarity of both the fixing and the washing solutions are therefore of prime importance to

Chapter 2:

Fixatives

avoid swelling and shrinkage artefacts, as already discussed in Sect. 2.3.2. Unfortunately, there is no definitive information on the contribution made by glutaraldehyde to the effective osmolarity of fixatives during their initial reaction with cells. In fact, the contribution will depend upon the membrane properties of the type of cell being fixed and is likely to vary considerably from one cell type to another. The balance of evidence in the literature suggests that glutaraldehyde often makes a significant contribution but one appreciably less than that predicted from its molarity in the fixing solution. A figure of about 50% appears to be appropriate for many tissues. Furthermore, according to Lee et al. (1982) the osmotic effect of glutaraldehyde depends upon the type of buffer present and is therefore difficult to predict. Small variations in the effective osmolarity of the initial fixative do not appear to cause major artefacts in the ultrastructural appearance of tissues, apart perhaps from some overall swelling or shrinking of the individual cells. In conclusion no single formulation for 'perfect' fixation can be proposed. For routine ultrastructural studies it is sufficient to assume that glutaraldehyde makes some contribution to the osmotic pressure experienced by cells during fixation. In practice a fixative consisting of 1.0 to 2.5% glutaraldehyde in a 0.1 M cacodylate, phosphate or PIPES buffer gives satisfactory ultrastructural fixation of cells in tissues and in culture without obvious signs of osmotic damage. 2.4.2

T h e p r e p a r a t i o n o f g l u t a r a l d e h y d e fixatives

Glutaraldehyde is available from general suppliers (see Appendix) as an aqueous solution in the range 8 to 50%, although 25% (EM grade) is the strength usually used. It should be stored at —20°C. If the solution turns yellow or has a pH below 4, it should be discarded. The safety pre­ cautions to be observed in its use and disposal, described in Sect. 2.2.1, must be strictly followed by everyone in the laboratory. To make up a standard primary glutaraldehyde fixative the pro­ cedure is as follows: 1.

Mix the alkaline and acid forms of the chosen buffer in the proportions given in Tables 2.1, 2.2 and 2.3 (see Sect. 2.3.3) to obtain a solution at the desired pH.

45

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Glauert and Lewis:

2. 3. 4.

Biological specimen preparation for TEM

Add sufficient calcium chloride (unless phosphate is being used) to give a final concentration of 2 to 3 mM and mix well. Add the necessary volume of a stock glutaraldehyde solution and make up to the appropriate final volume with distilled water. Check the pH of the fixative and adjust it, if necessary. An accuracy of 0.1 units is quite sufficient for electron microscopy.

To take a specific example, the most popular fixative for ultrastructural studies is prepared as follows: 1. To 100 ml of 0.2 M sodium cacodylate add 10 ml of 0.1 N HCl. 2. Add 5 ml of 0.1 M calcium chloride and mix well. 3. Add 20 ml of 25% glutaraldehyde from an automatic pipette or from a syringe, preferably of the disposable variety. 4. Make up to a total volume of 200 ml with distilled water and mix well. 5. Check the pH of the fixative, which should be near to 7.4. If it is below 7.2, add sufficient 0.1 N NaOH to bring it back into the range 7.2 to 7.4. This fixative contains 2.5% glutaraldehyde and 2.5 mM calcium chloride in 0.1 M cacodylate buffer at a pH of 7.2 to 7.4. If the pH initially measured is below about 6.8, the stock glutaraldehyde solution has probably deteriorated and should be replaced. Small quantities of other constituents, such as potassium chloride or a vasodilator, which may be necessary for fixation by perfusion, should be added after the calcium chloride and before the glutaraldehyde. Glutaraldehyde undergoes a progressive series of reactions in dilute solution and so the complete fixative should be stored at 4°C. If it is kept for more than a week, the pH should be re-checked to ensure that the glutaraldehyde has not decomposed. Glutaraldehyde fixatives buffered with phosphate or PIPES should be prepared freshly each day.

2.5

Formaldehyde fixatives

As mentioned in Sect. 2.1.1, formaldehyde in the form of commercial formalin was found to be unacceptable as a fixative for electron micro­ scopy, because it contains traces of formic acid and 10% or more of

Chapter 2:

Fixatives

methanol. A purer form of formaldehyde can be produced, however, by the depolymerization of paraformaldehyde, and this has been found to be suitable as a primary fixative for electron microscopy. To avoid a clumsy use of words, in all that follows, the word 'formaldehyde' will be used to mean the chemical prepared by depolymerizing paraformaldehyde by heating in a suitable aqueous solution to about 60°C. The method of depolymerization is described in detail later (Sect. 2.5.2). Formaldehyde, which has the empirical formula CH2O, reacts with water in aqueous solution to form the monohydrate with the structure HO —0¾-OH. At concentrations above about 1 to 2%, this hydrated formaldehyde begins to polymerize to form a series of polyoxymethylene glycols having the general formula HO — (CH—OH)„ — 0¾-OH, the percentage of these polymers and the average value of V rising as the overall concentration is increased. Interconversion of the polymers is a slow reaction and when a concentrated solution is diluted to 1 to 2% it takes many hours for depolymerization to become complete. This fact may be important in preparing formaldehyde fixatives, since the poly­ mers react more effectively with proteins than the monomer (Baschong et al. 1983, 1984). At a strength of 4%, a formaldehyde fixative will contain about 85% of the aldehyde in the monomeric form when an equilibrium has been reached, but only 66% if prepared immediately beforehand from a 10% stock solution (Walker 1964). It is probably wise not to use stronger stock solutions than this, since inconsistent results may be obtained because of the high proportion of polymeric forms present in more concentrated solutions. Formaldehyde is a gentler fixative than glutaraldehyde and many of its reactions with tissues are at least partially reversible. Formaldehyde is used as a primary fixative for many cytochemical techniques, but glutaraldehyde is preferred for purely ultrastructural studies. Form­ aldehyde, however, penetrates tissues more rapidly and may modify the semi-permeability of cell membranes more than glutaraldehyde. A com­ bined glutaraldehyde-formaldehyde mixture is therefore often the primary fixative of choice for many tissues. 2.5.1

The effects of formaldehyde on cells and tissues

At the concentrations of formaldehyde normally used for fixation (1 to 4%) the main effect on tissues is produced by the formation of

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Glauert and Lewis:

Biological specimen preparation for TEM

methylene cross-bridges between the free amino groups on proteins. As with glutaraldehyde the most important amino group is thought to be the terminal group on lysine residues. The initial reaction involves the simple loss of one molecule of water: R-NH 2 + HO-CH 2 -OH -» R-NH-CH 2 -OH + H 2 O The initial product formed will react with a second amino group with the loss of another molecule of water and the formation of a methylene bridge: R-NH-CH 2 OH+R'-NH 2 -> R-NH-CH 2 -NH-R'+H 2 O There is reason to believe that other groups besides the amino group of lysine can participate in the formation of these protein cross-bridges. Also implicated are amide and aminido groups, sulphydryl groups and the imidazole ring of histidine. There is some disagreement in the literature about the physical and chemical stability of these methylene cross-bridges. They are, in general, regarded as less stable than those formed by glutaraldehyde. The available experimental evidence in the literature (see Puchtler and Meloan 1985 for references), however, suggests that they are stable enough under the conditions to which tissues are subjected during normal processing for electron microscopy. It is possible that some effects of formaldehyde fixation can be reversed by prolonged washing in buffer (although not necessarily the methylene cross-bridges) and it is advisable to incorporate a small amount of formaldehyde (0.2 to 0.5%) into any solution used for prolonged storage of tissues. There is conflicting evidence about the speed of formation of the cross-bridges. Some changes in tissues occur within minutes of contact with formaldehyde, although many hours are required for these various changes to be complete (Fox et al. 1985), but it should be noted that perfectly adequate fixation may be achieved in a much shorter time. None the less there is general agreement that formaldehyde reacts with tissues more slowly than glutaraldehyde and forms fewer, less stable cross-bridges. A concentration of about 4% formaldehyde for several hours is probably required for adequate fixation of thin slices of tissue, and even then the preservation of ultrastructure is normally considered

Chapter 2:

Fixatives

to be inferior to that which can be achieved with glutaraldehyde in a much shorter time. Overall background staining of the general cell cytoplasm is also often less obvious when formaldehyde alone is used as the primary fixative. Formaldehyde reacts both with pure DNA (unlike glutaraldehyde) and with the histones normally associated with it, but again many of the reactions may be partially reversible. Formaldehyde appears not to react very strongly with either lipids or carbohydrates in their pure form, but does react strongly with any associated proteins. Formaldehyde, like glutaraldehyde, partially modifies, but does not completely destroy, the semi-permeability properties of most cell membranes. As a result, distortion of cells and cell organelles may occur if the subsequent washing solutions are not iso-osmotic. This is an important con­ sideration when cytochemical reactions on tissue slices are necessary before secondary fixation. As with glutaraldehyde the inclusion of calcium ions in the fixing, washing and storage solutions leads to better preservation of membrane structure. Formaldehyde, a mono-aldehyde, is less likely than glutaraldehyde, a di-aldehyde, to introduce extra free aldehyde groups into the tissue. For some cytochemical studies, therefore, formaldehyde is the preferred fixative. One very important difference in the actions of glutaraldehyde and formaldehyde is in the degree of destruction of the specific biological activity of the individual protein molecules. Formaldehyde, particularly if used at 4°C, retains much of the activity of many enzymes, certainly sufficient for cytochemical reactions to give satisfactory results in tissue slices incubated before post-fixation with osmium tetroxide. Similarly the immunoreactivity of many proteins is largely retained after formaldehyde fixation but not after glutaraldehyde. There are exceptions to these general statements, of course, but formaldehyde is often the primary fixative of choice for cytochemical studies and should be used first in preliminary investigations when retention of biological activity is important. Formaldehyde penetrates tissues rapidly and appears to contribute very little to the effective osmolarity of the complete fixative. It is therefore preferable to have the vehicle for the fixative only slightly hypoosmotic to body fluids (see Sect. 2.3.2). Three fixatives are proposed in the next section and all of them are suitable for perfusion as well as immersion. Since the initial reactions of formaldehyde with proteins are

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Glauert and Lewis:

Biological specimen preparation for TEM

largely reversible, whereas the slower formation of methylene bridges is irreversible, optimum fixation times for formaldehyde are usually longer than for glutaraldehyde. Fixation of thin slices of tissue for 4 to 6 hours at room temperature is typical for formaldehyde alone and fixation overnight appears not to be deleterious for ultrastructural studies, provided 2 to 3 mM calcium ions are included in the fixative. It should perhaps be emphasized here that because many of the reactions of formaldehyde can be reversed, prolonged storage of tissue is not recommended and calcium ions must be included even for short periods of storage to aid the retention of lipids and to improve membrane preservation. 2.5.2 The preparation of formaldehyde fixatives Formaldehyde fixatives are most often used at a final strength of 2 to 4% in an aqueous vehicle at a pH of 6.8 to 7.4. Formaldehyde, prepared from paraformaldehyde, is so nearly neutral that the pH of the final fixative should not need to be adjusted, although it should be checked before use. To prepare an aqueous solution the paraformaldehyde must be warmed to about 60°C. This operation must be conducted in a fume cupboard and the heated solution should be of the minimum concentration necessary to prepare the final fixative, say 8 to 10% formaldehyde as a maximum. When phosphate is used, dissolve the paraformaldehyde in the alkaline form of the buffer, that is, in the Na2HPC>4 solution. The paraformaldehyde then forms a clear solution when the temperature reaches 50 to 55°C, when the heating should be turned off and the solution allowed to cool. If the paraformadehyde is dissolved in water or in a saline mixture, heat to 60°C and, if necessary, add a drop or two of N NaOH to produce a clear solution. Opinions differ about the stability of formaldehyde solutions. For critical studies, both ultrastructural and cvtochemical, it is probably safest to make up fresh solutions each week, but some research workers have stored dilute buffered formaldehyde solutions for several months. A solution should be rejected if any opalescence appears.

Chapter 2:

2.5.2a

Fixatives

A phosphate-buffered formaldehyde fixative

1. To 80 ml of 0.1 M Na2HP04 in a conical flask add 2.0 to 4.0 g of paraformaldehyde. Place the flask on a hot plate or in a water bath and warm gently, preferably with a magnetic stirrer included. This operation should be carried out in a fume cupboard or on a ventilated bench. 2. When this solution has cleared, usually when a temperature of about 50 to 55°C has been reached, allow it to cool to room temperature and add 20 ml of 0.1 M NaH2P04. This fixative contains 2 to 4% formaldehyde in 0.1 M sodium phosphate buffer at a pH of 7.4 or just below, but no calcium ions. To prepare a phosphate-buffered formaldehyde fixative at other pH values, use the volumes of the two solutions given in Table 2.1 in Sect. 2.3.3 in exactly the same way. These phosphate-buffered fixatives keep well, so that large quantities can be made up and stored in a brown bottle at room temperature ready for use, although it may be better to use freshly madeup fixative for some critical studies. 2.5.2b A cacodylate-buffered formaldehyde fixative

1. To 50 ml of distilled water in a conical flask add 4.0 to 8.0 g of paraformaldehyde and warm gradually to 60°C. If the solution does not clear, add one or two drops of N NaOH. 2. When this solution is cool add: 0.2 M sodium cacodylate 100 ml 0.1 M calcium chloride 5 ml 3. Mix well and add the volume of N HCl sufficient to bring the pH to 7.0 to 7.4, as given in Table 2.2 in Sect. 2.3.3. 4. Make up to a total volume of 200 ml with distilled water. This fixative contains 2 to 4% formaldehyde and 2.5 mM calcium chloride in 0.1 M cacodylate buffer. It can be stored in a brown bottle for several weeks at 4°C.

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2.5.2c

Biological specimen preparation for TEM

A PIPES-buffered formaldehyde fixative

To 40 to 80 ml of distilled water in a conical flask add 4.0 to 8.0 g of paraformaldehyde. Warm gently to 60°C, and if the solution does not clear, add one or two drops of N NaOH. 2. Prepare a 0.2 M solution of PIPES by dissolving 6.1 g of PIPES in 100 ml of distilled water, add 5 ml of 0.1 M calcium chloride, followed by the formaldehyde solution when it is cool. 3. Mix well and add the volume of N NaOH necessary for a pH of 7.0 to 7.4, as given in Table 2.3 in Sect. 2.3.3. 4. Make up to a total volume of 200 ml with distilled water. 1.

This fixative contains 2 to 4% formaldehyde and 2.5 mM calcium chloride in 0.1 M PIPES buffer. It does not keep well and should be made up freshly each day. 2.5.3

Glutaraldehyde-formaldehyde fixatives

Both glutaraldehyde and formaldehyde have their advantages as fixatives and in some tissues a mixture of the two can give better preservation of ultrastructure than either alone. These mixtures are very widely used as primary fixatives at the present time for many types of tissue (see Chapter 3). Formaldehyde penetrates tissues much more rapidly than glutaraldehyde and it is thought that the formaldehyde temporarily stabilizes structures which are subsequently fixed more permanently by the glutaraldehyde. In consequence pieces of tissue are well fixed to a much greater depth than with glutaraldehyde alone. It is still advisable to add calcium chloride to the fixative whenever possible, and fixation must be followed by a second fixation with osmium tetroxide, and preferably also with uranyl acetate. It should be noted that the chemistry of fixation by glutaraldehydeformaldehyde mixtures is not fully understood. With glutaraldehyde alone oxygen is taken up and pyridine derivatives are formed, as explained in Sect. 2.4.1a. In the presence of formaldehyde, however, oxygen uptake is largely suppressed. Whether formaldehyde takes over the role of an oxidizing agent is not known, but the preservation of ultra­ structure of some tissues seems better with the mixture.

Chapter 2:

Fixatives

The fixative originally suggested by Karnovsky (1965) consisted of 4% paraformaldehyde, 5% glutaraldehyde and 0.05% calcium chloride in 0.08 M cacodylate buffer, pH 7.2. Lower concentrations of the aldehydes are now normally used, and most recent practice has been to use 0.5 to 2.0% formaldehyde and 0.5 to 4.0% glutaraldehyde, together with the addition of 2 to 3 mM calcium ions when possible, in 0.10 M buffer. To make up a mixed fixative add the appropriate volume of concentrated glutaraldehyde solution from a disposable syringe or pipette (see Sect. 2.4.2) to one of the formaldehyde fixatives given in Sect. 2.5.2. 2.5.3a The preparation of a glutaraldehyde-formaldehyde fixative

1.

2.

3.

4.

To 50 ml of distilled water in a conical flask add 2.0 to 4.0 g of paraformaldehyde and warm gradually to 60°C in a fume cupboard. If the solution has not cleared, add one or two drops of N NaOH. When this solution is cool add: 0.2 M sodium cacodylate 100 ml 0.1 M calcium chloride 5 ml Add 4 to 32 ml of 25% aqueous glutaraldehyde to this solution. Check the pH, with the solution still in a fume cupboard, and add sufficient N HCl to bring the pH within the range of 7.0 to 7.4. Make up to a total volume of 200 ml with distilled water.

This fixative contains 1.0 to 2.0% formaldehyde, 0.5 to 4.0% glutaraldehyde, and 2.5 mM calcium chloride in 0.1 M cacodylate buffer. It should be freshly prepared just before use. 2.5.4

Additives to aldehyde fixatives

Many substances have been advocated as possible additives to aldehyde fixatives in order to improve some aspect or other of the appearance of the specimen in the final electron micrograph. Some specific additives are dealt with in detail later, such as dichromate for the demonstration of biogenic amines (Sect. 2.8.3) and tannic acid for the preservation of a range of ultrastructural components (Sect. 2.8.2) The remaining additives which appear to merit a mention are included in this section.

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Fig. 2.5 Ruthenium red staining of the cell surface of a mouse lymphocyte, fixed in cacodvlate-buffered 2.5% glutaraldehvde for 1 hour, followed bv a mixture of 1.5% osmium tetroxide and 0.1% Ruthenium red in cacodylate buffer, pH 7.2, for 3 hours at room temperature. Scale bar = 0.2 μιη.

Scallen and Dietert (1969) added digitonin to improve the retention of free cholesterol, which is an essential component of many membranes, but distortion of ultrastructure by digitonin-cholesterol complexes occurs. Similarly, Malachite green has been advocated as an additive, at an overall concentration of 0.1%, to give better preservation of some cellular lipids. Alcian blue or Ruthenium red, at a concentration of 0.5 to 2.0%, can be used to enhance the staining of cell surfaces (Fig. 2.5) and various extracellular substances. Picric acid (2,4,6-trinitrophenol) and other trinitro compounds have also been proposed as additives to aldehyde fixatives. Ito and Karnovsky (1968) tested a range of compounds and it would appear that 2,4,6trinitrocresol in the concentration range of 0.02 to 1.0% gave the most satisfactory results when added to a mixed formaldehyde-glutaraldehyde fixative. In particular, membrane systems were well fixed, myelin figures were rare and excellent preservation of smooth endoplasmic reticulum in steroid-secreting cells was obtained. One such fixative,

which is

Chapter 2:

Fixatives

particularly suitable for some immunocytochemical studies, consists of a mixture of 0.05% glutaraldehyde, 4% formaldehyde and 0.2% picric acid. For ultrastructural studies a higher concentration of glutaraldehyde, say 2.0%, is to be preferred. All these trinitro additives probably work by destroying the impermeability of membranes, allowing rapid penetration of the aldehyde fixatives into cells and organelles, as well as by acting as protein precipitants. Ruthenium hexammine trichloride (at a concentration of 0.4 to 0.7%) has been recommended as an additive to a standard glutar­ aldehyde fixative to preserve the proteoglycans in cartilage (Hunziker et al. 1982). The same concentration is added to the osmium tetroxide post-fixative. If an optimum osmolarity is used, there are no artefactual spaces around the chondrocytes of cartilage, but intracellular detail is poorly preserved. Ruthenium hexammine trichloride therefore has only a very specialized role as a fixative.

2.6

O s m i u m t e t r o x i d e fixatives

The value of osmium tetroxide for the preservation of fine detail was made apparent long ago when Strangeways and Canti (1927) studied the effects of various fixatives on the appearance of living cells by dark-field microscopy (see Fig. 1.1 in Sect. 1.2). They found that osmium tetroxide was the only fixative of those tested that preserved delicate cytoplasmic processes and caused no alteration in mitochondria and fat droplets. When the first studies on ultrathin sections were made in the late 1940s and early 1950s it was soon found that osmium tetroxide gave better preservation of cellular fine structure than other fixatives developed for light microscopy. The main disadvantage of osmium tetroxide as a primary fixative is that it penetrates tissues so slowly that considerable changes in structure can occur before fixation is complete, particularly in the centre of even quite small specimens. In consequence osmium tetroxide has been completely superseded by aldehydes as a primary fixative and the appearance of specimens fixed with osmium tetroxide alone will not be considered in any detail here. It still plays an essential role, however, as a second fixative since it reacts with components of tissues, particularly

55

Glauert and Lewis:

Biological specimen preparation for TEM

lipids, that are not fixed by aldehydes, and also acts as a stain. During this second fixation the slow rate of penetration of osmium tetroxide is no longer such a disadvantage since the structure of the specimen has already been partially stabilized by the primary aldehyde fixation. 2.6. I

T h e effects o f osmium tetroxide fixation

The chemical reactions by which tissues are fixed by osmium tetroxide are only poorly understood, but the tetroxide is known to react with lipids, carbohydrates and proteins (for reviews see Hayat 1981 and Behrman 1984). Osmium can exist in a series of oxidation states, the most important for fixation

being eight as in the tetroxide, six as in

potassium osmate and four as in osmium dioxide. In fixed tissues much of the osmium is initially covalently bound in the hexavalent and tetravalent states, but after dehydration and embedding most of it probably exists as the black dioxide. Osmium tetroxide reacts avidly with the olefinic double bond of unsaturated lipids to form a five-membered

ring containing two oxygen

atoms separated by an osmium atom which is now in the hexavalent state (see Fig. 2.6). Cross-linkage between two lipid molecules can occur by the

CH (a)

HC"^ + OsO 4

(J

•

HC VIII

.

-OH

I CH

+

VI

.

\/ CH (b)

Os /

J

CH

\

\ -OK _ HC"^ \ ^ . ° OsOj 1 —•

^ ^OH

I HC

Os /

/ VI

O

VI

Fig. 2.6 Illustrations of two important initial reactions in osmium fixation. The \alencv of the osmium is given as a roman numeral, (a) Osmium tetroxide reacts extremely rapidly with the ethylenic double bonds in unsaturated lipids to form a five-membered ring compound, (b) The same ring structure is formed by the reaction of an osmate ion with a ns-diol group in a carbohydrate.

Chapter 2:

Fixatives

formation of a dimeric complex of two of the hexavalent osmium atoms in five-membered rings joined by two oxygen atoms in a four-membered ring (Fig. 2.7). Hexavalent osmium formed by reduction forms a similar bridging five-membered ring with carbohydrate chains containing two adjacent, as-orientated, hydroxyl groups (Fig. 2.6). The formation of these five-membered rings is a very characteristic feature of the reactions of osmium with organic compounds, and the rings are highly stable (both physically and chemically) in aqueous solution. Osmium readily forms a total of six links to other atoms and any of the six positions not occupied by oxygen are avidly filled by nitrogen atoms. Thus the osmium compounds formed with lipids and carbohydrates form stable addition products with proteins through free amino, imino and aminido groups (Fig. 2.7). Osmium tetroxide is reduced by the free sulphydryl group of cysteine residues and the osmium remains linked to the sulphur atom. All these reactions are rapid, and go virtually to completion under the conditions normally used for fixation with osmium tetroxide, that is, a 1% aqueous solution at 4 to 22°C. The most important fact, however, is that the osmium which becomes bound to lipid or carbohydrate readily binds to nitrogen atoms in proteins (as in histidine, tryptophan, arginine and lysine side chains), thereby forming cross-links between the different tissue components which help to stabilize the cytoplasmic structures and surface membranes. Many of these compounds probably react with

ο HC-"" °\ Il ^NH2 (a)

+ Amino groups

•

| ^C\

\ (b)

Dimerization

„

R.

Os 0 /|^

nh

0

Il

R2

2

0

n

Il

4 fixative which selectively contrasts glycogen. Journal of Ultrastructural Research 42, 29-50 De Bruijn, W.C. and Van Buitenen, J.M.H. (1980) X-ray microanalysis of aldehyde-fixed glycogen contrast stained by Os vl -Fe" and Os vi -Ru vl complexes. Journal of Histochemistry and Cytochemistry 28, 1242-1250 De Bruijn, W.C., Memelink, A.A. and Riemersma, J.C. (1984) Cellular membrane contrast and contrast differentiation with osmium triazole and tetrazole complexes. HistochemicalJournal 16, 37-50 Fairen, A. and Smith-Fernandez, A. (1992) Electron microscopy of Golgi-impregnated interneurons: notes on the intrinsic connectivity of the cerebral cortex. Microscopy Research and Technique 23, 289-305 Fox, C.H., Johnson, F.B., Whiting, J. and Roller, P.P. (1985) Formaldehyde fixation. Journal of Histochemistry and Cytochemistry 33, 845-853 Gibbins, I.L. (1982) Lack of correlation between ultrastructural and pharmacological types of non-adrenergic autonomic nerves. Cell and Tissue Research 221, 551-581 Glauert, A.M. (1975) Fixation, dehydration and embedding of biological specimens. In Practical Methods in Electron Microscopy, Vol. 3, Part I, Glauert, A.M. (ed.), NorthHolland, Amsterdam Good N.E., Winget, G.D., Winter, W., Connolly, T.N., Izawa, S. and Singh, R.M.M. (1966) Hydrogen ion buffers for biological research. Biochemistry 5,467-477 Griffiths, G. (1993) Fine Structure Immunocytochemistry. Springer-Verlag, Berlin and Heidelberg Hayat, M.A. (1981) Fixation for Electron Microscopy. Academic Press, San Diego and London Hopwood, D. (1970) The reactions between formaldehyde, glutaraldehyde and osmium tetroxide, and their fixation effects on bovine serum albumin and on tissue blocks. Histochemie 24, 56-64 Hopwood, D. (1972) A review: theoretical and practical aspects of glutaraldehyde fixation. HistochemicalJournal 4, 267-303 Hopwood, D. (1985) Cell and tissue fixation, 1972-1982. Histochemical Journal 17, 389-142 Hunziker, E.B., Herrmann, W. and Schenk, R.K. (1982). Improved cartilage fixation by ruthenium hexammine trichloride. Journal of Ultrastructure Research 81, 1-12 Ito, S. and Karnovsky, M.J. (1968) Formaldehyde-glutaraldehyde fixatives containing trinitro compounds. Journal of Cell Biology 39,168a only Johnson, T.J.A. (1986) Glutaraldehyde fixation chemistry. A scheme for rapid crosslinking and evidence for rapid oxygen consumption. In The Science of Biological Specimen Preparation, Muller, M., Becker, R.P., Boyde, A. and Wolosewick, J.J. (eds.), pp. 51-62, SEM Inc., AMF O'Hare, Chicago Johnson T.J.A. (1987) Glutaraldehyde fixation chemistry: oxygen-consuming reactions. EuropeanJournal of Cell Biology 45, 160-169

Glauert and Lewis:

Biological specimen preparation for TEM

Karnovskv, M.J. (1965) A formaldehyde-glutaraldehyde fixative of high osmolarity for use in electron microscopy.Journal of Cell Biology 27,137A only Kiernan, J.A. (1978) Recover)' of osmium tetroxide from used fixative solutions. Journal of Microscopy 113, 77-82 Kobs, S.F. and Behrman, E.J. (1987) Reactions of osmium tetroxide with imidazole. Inorganica Chemica Acta - Bioinorganic Chemistry 138, 113-120 Lee, R.M.K.W. (1984) A critical appraisal of the effects of fixation, dehydration and embedding on cell volume. In The Science of Biological Specimen Preparation, Revel, J.-P., Barnard, T. and Haggis, G.H. (eds.), pp. 61-70, SEM Inc., AMF O'Hare, Chicago Lee, R.M.K.W., McKenzie, R., Kobayashi, K., Garfield, R.E., Forrest, J.B. and Daniel, E.E. (1982) Effects of glutaraldehyde fixative osmolarities on smooth muscle cell volume, and osmotic reactivity of the cells after fixation. Journal of Microscopy 125, 77-88 Lewis, P.R. (1983) Fixatives: hazards and safe handling. Proceedings of the Royal Microscopical Society 18, 164—167 Lewis, P.R. and Knight, D.P. (1992) Cytochemical staining methods for electron microscopy. In Practical Methods in Electron Microscopy, Vol. 14, Glauert, A.M. (ed.), Elsevier, Amsterdam Locke, M. (1994) Preservation and contrast without osmification or section staining. Microscopy Research and Technique 29,1-10 Maupin, P. and Pollard, T.D. (1983) Improved preservation and staining of HeLa-cell actin-filaments, clathrin-coated membranes and other cytoplasmic structures by tannic-acid glutaraldehyde-saponin fixation. Journal of Cell Biology 96, 51-62 Mersev, B. and McCully, M.E. (1978) Monitoring of the course of fixation of plant cells. Journal of Microscopy 114, 49-76 Peracchia, C. and Mittler, B.S. (1972) Fixation by means of glutaraldehyde-hydrogen peroxide reaction products. Journal of Cell Biology 53, 234—238 Puchtler, H. and Meloan, S.N. (1985) On the chemistry of fomaldehyde fixation and its effects on immunohistochemical reactions. Histochemistry 82,201-204 Reedy, M.K. and Reedy, M.C. (1985) Rigor crossbridge structure in tilted single filament layers and flared-X formations from insect flight muscle. Journal of Molecular Biology 185,145-176 Riemersma, J.C. (1968) Osmium tetroxide fixation of lipids for electron microscopy. A possible reaction mechanism. Biochimica et Biophysica Acta 152, 718-727 Robards, A.W. and Sleytr, U.B. (1985) Low temperature methods in biological electron microscopy. In Practical Methods in Electron Microscopy, Vol. 10, Glauert, A.M. (ed.), Elsevier, Amsterdam Sabatini, D.D., Bensch, K. and Barrnett, R.J. (1963) Cytochemistry and electron microscopy. The preservation of cellular ultrastructure and enzymatic activity by aldehyde fixation. Journal of Cell Biology 17, 19-58 Sabatini, D.D., Miller, F. and Barrnett, R.J. (1964) Aldehyde fixation for morphological and enzyme histochemical studies with the electron microscope. Journal of Histochemistry and Cytochemistry 12, 57-71 Scallen, T.J. and Dietert, S.E. (1969) The quantitative retention of cholesterol in the mouse liver prepared for electron microscopy by fixation in digitonin-containing aldehyde solution. Journal of Cell Biology 40, 802-813 Strangeways, T.S.P. and Canti, R.G. (1927) The living cell in vitro as shown by darkground illumination and the changes induced in such cells by fixing reagents. Quarterly Journal of Microscopical Science 71, 1-14

Chapter 2:

Fixatives

Tashima, T., Kawakami, U., Harada, M., Sakata, T., Satoh, N., Nakagawa, T. and Tanaka, H. (1987) Isolation and identification of new oligomers in aqueous solution of glutaraldehyde. Chemical Pharmacology Bulletin 35, 4169—4180 Tranzer, J.-P. and Richards, J.G. (1976) Ultrastructural cytochemistry of biogenic amines in nervous tissue: methodologic improvements. Journal of Histochemistry and Cytochemistry 24, 1178-1193 Walker, J.F. (1964) Formaldehyde, 3rd edn. Van Nostrand-Reinhold, New York Weibel, E.R. (1979) Stereological Methods: Practical Methods for Biological Morphometry. Academic Press, London Whipple, E.B. and Ruta, M. (1974) The structure of aqueous glutaraldehyde. Journal of Organic Chemistry 39,1666-1668 White, D.L., Andrews, S.B., Faller, J.W. and Barrnett, R.J. (1976) The chemical nature of osmium tetroxide fixation and staining of membranes by X-ray photoelectron spectroscopy. Biochimica et Biophysica Acta 436, 577-592 Wouterlood, F.G. (1992) Techniques for converting Golgi precipitates in CNS neurons into stable electron microscopic markers. Microscopy Research and Technique 23, 275-288

3 Fixation methods

There are many ways of fixing specimens for electron microscopy and they are as varied as the types of specimen to be fixed. It is possible, therefore, to lay down only very general guide-lines for the best method or methods to be followed. In most investigations it is advisable to first try a method that has been used by previous workers with satisfactory results on similar specimens and then to make such empirical modifi­ cations as seem appropriate in the light of the results. This chapter covers a wide range of methods, tissues and specimens. It should normally be possible to find here a technique that can form the basis for a preliminary investigation, and pilot studies are always worthwhile. The fixation techniques available range from simple immersion of the specimen in a solution of the fixative to highly localized, intra­ arterial perfusion of a sophisticated fixing solution into an animal under anaesthesia. Methods for the fixation of animal tissues for ultrastructural studies are considered first and are dealt with in approximate order of increasing complexity of the equipment required. Methods for other types of tissue are considered next, again in approximate order of in­ creasing experimental complexity, followed by methods for organ and cell cultures, isolated cells and pellets.

3.1

The choice of fixation method

It is sometimes easy to decide upon the method of fixation to be used. For small specimens, such as tissue cultures, cell suspensions and pellets of iso­ lated organelles, fixation in situ is the obvious choice. Similarly, biopsy specimens are usually fixed by simple immersion. The best method of fixing many invertebrates, particularly insects, is by injection. For verte­ brates the decision on which method to use can be difficult and should be

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Biological specimen preparation for TEM

governed mainly by the time it will take for the fixing agent to reach all the cells of interest. In all electron microscope investigations it is vitally important to limit post-mortem changes to a minimum. This fact is dramatically illustrated in Fig. 3.1. For animal tissues the interval between death and exposure to the fixative should always be as short as possible. This is particularly important in birds and mammals which have a high body temperature. The less the normal blood supply to the tissue the slower in general will be the post-mortem changes. Even in tissues with little or no blood supply, however, such as subcutaneous connective tissue or articular cartilage, it is advisable to begin fixation as soon as possible. The diffusion of a fixative into a tissue is a slow and complex process (see Sect. 2.1.3). Consequently fixation by immersion of larger pieces of animal tissues or organs has many disadvantages. Because diffusion is so slow, post-mortem changes can occur in the deeper layers of a thick piece of tissue before the fixative arrives. The buffer and other constituents may penetrate tissues more rapidly than the fixative itself and so cause morphological changes before the fixation process starts. Furthermore, one of the initial effects of the fixative, which is more marked with glutaraldehyde than with formaldehyde, is to reduce the inward rate of diffusion of the fixative into the deeper layers of the tissue so that the rate of fixation differs with depth more profoundly than might be expected from simple diffusion theory. Hence cells in the deeper layers are exposed to very different fixation conditions from those lying more superficially. An added problem with glutaraldehyde as the sole fixative is that the deeper tissue layers may become anoxic before this slowly penetrating agent arrives, and this can alter the interaction of the glutaraldehyde with proteins (see Sect. 2.4.1a). Anoxia also alters the appearance of some cellular organelles, such as mitochondria, and so too does autolysis, which continues until it is ultimately stopped by the action of the fixative. For several reasons, therefore, it is necessary to keep the diffusion distance to a minimum by using sufficiently small specimens. Immersion is technically much the simplest method of fixation, but perfusion fixation has many advantages and should always be considered as a possibility. With perfusion the blood is first displaced by a suitable saline solution and then immediately by the fixative. Hence fixation starts before any autolytic or other degradative changes have a chance to occur. It is in the preservation of the overall ultrastructure, and especially

Chapter 3:

Fixation methods

Fig. 3.1 Isolated rat hepatocytes fixed in 4% glutaraldehyde (a) immediately, and (b) after suspension for 1 hour in unbuffered 0.9% sodium chloride. The onset of necrosis is very obvious in the lower illustration. Scale bar -2 μηι. (Unpublished micrographs from a study by DrJeremy Skepper, reproduced with permission.)

Glauert and Lewis:

Biological specimen preparation for TEM

of the integrity of intracellular organelles, that fixation by perfusion offers the greatest benefits (Fig. 3.2). When perfusion is a practical possibility the choice of fixation method is probably best governed by the degree of vascularization of the

Fig. 3.2 A comparison of fixation by (a) immersion and (b) perfusion of a rat Leydig cell. The smooth endoplasmic reticulum is very poorly preserved after immersion fixation. (Reproduced from Bozzola and Russell 1992, with permission.)

Chapter 3:

Fixation methods

tissue to be fixed. In a rapidly metabolizing tissue, such as mammalian skeletal muscle, the maximum distance between adjacent capillaries is likely to be considerably less than 50 μπι, and well under 20 μιη in mam­ malian cardiac muscle. Clearly in such specimens diffusion of fixing agents will be far more rapid if perfusion is used. In connective tissue or articular cartilage, fixation by immersion will be better, especially if the regions of interest have been exposed by rapid preliminary dissection. Sometimes these tissues are best fixed by injection of the fixative immediately post-mortem or under deep anaesthesia. Another important factor in deciding the method of fixation is the percentage of extracellular space and its degree of interconnection. Where these are high, as in many tissues containing a lot of connective tissue, im­ mersion fixation is quite adequate and may be preferred. In tightly packed tissues with little extracellular space, such as mammalian or avian brain or many endocrine tissues which are of epithelial origin, perfusion fixation is the only way to obtain good preservation of ultrastructure. This conclusion is strongly reinforced in a paper by Wild and Setoguti (1995) who summarize a long series of studies of the parathyroid gland and its ultrastructure. These studies showed that the most consistent, homo­ geneous picture of ultrastructure was obtained by initial perfusion with a suitable aldehyde fixative. In whole-animal perfusion, there is the added bonus that several tissues can be fixed simultaneously from the same animal. Minor disadvantages are that large quantities of the fixing reagents have to be used and that more care has to be taken to avoid chemical hazards from the fixative. These disadvantages are outweighed by the much better and more uniform fixation which can be obtained by perfusion (Fig. 3.2).

3.2

The choice of fixatives

The composition and properties of the various fixatives used in electron microscopy have already been described in Chapter 2. It is important to emphasize here that the precise composition of the fixatives chosen will depend critically on the needs of the experimenter. When the prime need is for accurate preservation of ultrastructure, a relatively strong mixed aldehyde fixative is indicated (say 2 to 4% glutaraldehyde plus 1 to 2%

81

82

Glauert and Lewis:

Biological specimen preparation for TEM

formaldehyde), followed by post-fixation in 1% osmium tetroxide and treatment 'en bloc' with 1% aqueous uranyl acetate. Such a protocol ensures that most chemical constituents of the specimen, such as proteins, nucleic acids, carbohydrates and lipids, react chemically in such a way as to ensure their continued immobilization throughout subsequent processing of the material. When some cytochemical or immunocytochemical techniques are also to be applied, a different choice of fixatives may be necessary to prevent unwanted chemical changes to particular cellular components. Thus a lower concentration (say 0.5%) of glutaraldehyde is sometimes preferable and the uranyl acetate treatment is often omitted. For enzyme cyto­ chemistry, the primary aldehyde has to be at 4°C and tailored to the particular enzyme under study, and the cytochemical technique is applied before the post-fixation in osmium tetroxide. For immunocytochemical studies, the primary fixation becomes even more critical. Low concen­ trations of aldehydes (say 0.05 to 0.20%) are sometimes required, without post-fixation, to preserve the epitopes under study, with inevitable loss of some ultrastructural detail, and aldehyde fixation may need to be followed by dehydration at low temperatures. Thus every fixation procedure is a compromise and it is necessary to determine empirically the most appropriate method of fixation for each tissue, enzyme and antigen. The adjective 'routine' is often applied to the noun 'fixation' but no one method of fixation should be regarded as immutable. When satisfactory results have been obtained with some established fixation procedure, thought should be given to possible ways of improving the overall result. Concentration, duration and temperature are variables which may produce improvements, but exhaustive study of all the permutations and combinations is obviously out of the question. Common sense and an elementary knowledge of the chemistry involved should be a sufficient guide. More important still is the use of possible additives, such as tannic acid to the aldehyde fixative or potassium ferroor ferri-cyanide to the osmium tetroxide. Some consideration of the rationale for these other additives and fixing agents was given in Chapter 2. Details of their practical application to specific problems are given either in this Chapter or in Chapter 10.

Chapter 3:

3.3

Standard fixation methods

3.3. 1

Primary fixation

Fixation methods

The optimum temperature for primary fixation is very dependent on the type of specimen. For animal tissue it is still very much a matter for debate, and there has been some controversy in the literature. Some specimens, particularly mammalian cells in culture or in suspension, show rapid surface changes when exposed to a sudden drop in temperature, in view of which it could be argued that the initial contact between tissue and fixative should be at the environmental temperature of the tissue. In fixation by perfusion body temperature is probably maintained, at least for a brief period, which should be sufficient to prevent changes from occurring while the tissue later cools. In fixation of biopsies by immediate immersion some fixative probably penetrates before the temperature has fallen significantly. Most workers do not pre-warm the fixative when fixing tissue from warm-blooded animals, but use the fixative at ambient temperature, placing tissues in the refrigerator later if fixation at 4°C is recommended for some specialized techniques. Ideally monolayer cultures should be fixed at the temperature of incubation, although some workers allow them to cool for a few minutes to room temperature before applying the fixative. For botanical specimens and tissue from cold-blooded animals, of course, the ambient temperature is used. After a short period, primary fixation can be continued at room temperature or at 4°C; the lower temperature may possibly reduce the extraction of lipids and other cell constituents, but a longer period of fixation is required. For purely ultrastructural studies, room temperature is usually best. The total time of fixation depends on the type of fixative, the type of specimen and on the needs of the investigation and may vary from a few minutes to a few hours. For cell suspensions or monolayer cultures 10 to 30 min are sufficient. After perfusion, half an hour of immersion fixation is quite adequate, and may not be necessary at all. Tissues fixed by immersion only need a longer period of fixation for samples more than about 0.5 mm thick. 3.3.2

Washing and storage

It is customary to wash the specimen in a buffer solution after primary aldehyde fixation and before any subsequent treatment such as post-

83

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Glauert and Lewis:

Biological specimen preparation for TEM

fixation with osmium tetroxide. The objective is to remove traces of aldehyde which might interfere with later fixation steps. Ideally the washing solution should be made up with calcium ions in the same buffer and with the same osmolarity (see Sect. 2.3) as the primary fixative in order to reduce the changes in the environment of the specimen to a minimum. Usually two or three changes of the washing solution over a period of 1 to 2 hours is considered sufficient for slices of tissue 250 μηι thick. During this washing period is the ideal time to dissect out selected areas of a tissue to form pieces for eventual embedding. After treatment with osmium tetroxide most tissues are too brittle to dissect and individual structures are too difficult to identify because of the general blackening. It is at this stage that many cytochemical and immunocytochemical techniques are performed; that is after primary fixation and before secondary fixation (Lewis and Knight 1992; Griffiths 1993). These techniques usually require the tissue to be sectioned thin enough for adequate penetration of the reagents, and it is often important to maintain the tissues in a known orientation during processing. These problems are discussed in more detail in Sect. 3.5.1. It is often convenient to store aldehyde-fixed specimens after washing. Again it is ideal if the same buffer can be used. Phosphate buffers are not suitable, however, except perhaps for overnight storage, mainly because calcium ions cannot be added to help preserve lipids. Cacodylate is often the ideal buffer for storage. It does not form precipitates with calcium ions, which are normally added to a final concentration of 2 to 3 mM, and it is a good buffer to counteract any acid that may be released by the specimen during storage. It depresses any autolysis which might occur, and inhibits growth of moulds and other micro-organisms. PIPES, HEPES and MOPS buffers can also be used, but then a crystal of thymol should be added to inhibit the growth of micro-organisms. Storage should be at 4°C. If specimens are to be stored for more than a few days, some workers add a small amount (5 to 10%) of the original fixing solution to the washing solution to help preserve the tissue and prevent any reversal of the formation of cross-bridges which otherwise might occur, particularly with formaldehyde (Sect. 2.5.1). The effect of adding a fixative during storage should be tested in any particular investigation before adopting it as a routine. Excess fixative should be removed by washing with buffer before proceeding. It is certainly better not to store tissues for very long periods for cytochemical and immunocytochemical studies if this can be avoided.

Chapter 3:

3.3.3

Fixation methods

Secondary a n d tertiary fixation

After primary aldehyde fixation and washing in buffer, samples are postfixed with osmium tetroxide (Sect. 2.6) for ultrastructural studies. The time and temperature for this fixation are much less critical than for the first. Specimens are normally fixed in 1% osmium tetroxide, either unbuffered or with the same buffer as for the primary fixation, at room temperature for 1 to 2 hours. When it is particularly important to preserve glycogen and to obtain strong staining of lipid membranes, some research workers incorporate potassium ferricyanide into the osmium fixative. A typical formulation is 1.0% osmium tetroxide plus 1.5% potassium ferricyanide, either unbuffered or in 0.1 M PIPES buffer at pH 7.2 to 7.4. For all ultra­ structural studies treatment with uranyl acetate is strongly recommended as a third fixative, since it gives particularly good preservation and staining of membrane structure (Sect. 2.7). After osmium tetroxide fixation, samples are washed briefly in distilled water and then immersed in 0.5 to 1.0% aqueous uranyl acetate for 1 to 2 hours at room temperature in the dark. Details for the safe use and disposal of osmium tetroxide are given in Sect. 2.2 and Sect. 2.6.2, and for uranyl acetate in Sect. 2.2 and Sect. 2.7.

3.4

Fixation by i m m e r s i o n

The simplest method of fixation is by immersion directly in the primary aldehyde fixative. It is necessary to use small specimens, because of diffusion problems. Cubes of tissue should be no more than 1 mm thick in each dimension, and it is preferable to use slices no more than 0.5 mm thick, and ideally only about 250 μηι thick. Cutting tissue so thinly inevitably causes mechanical damage even when especially sharp equipment is used. Suitable methods for cutting slices are described in Sect. 3.5.1a. The fixative is contained in a glass vial, preferably about 20 mm in diameter and from 20 to 30 mm high, with a close-fitting polyethylene cap (Fig. 3.3). Vials of this size are convenient, since they are easy to handle without risk of accidents, are economical in the use of reagents, and stand firmly on a flat surface. In many procedures the specimens can remain in these vials throughout fixation, dehydration and infiltration with the embedding medium, and need only be removed when they are placed in capsules or moulds for the final embedding. Unfixed specimens are easily damaged and they are transferred to pre-labelled vials containing the

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primary fixative as gently as possible with a Pasteur pipette having a mouth just wide enough to take the specimen or on the tip of a folded piece of glazed paper or filter paper. The relative volumes of the specimen and the fixative are important, because if there is too little fixative, the effective concentration of the fixing agent is reduced by dilution with soluble components of the specimen and by combination of specimen components, such as proteins, with aldehydes during the fixation process. In general, the fixative should be at least 10 to 20 times greater in volume than the specimen. Fixation is usually at room temperature, but if it needs to be carried out at 0 to 4°C the vials containing the fixative are placed in a bath of crushed ice some time before fixation is due to start. Typically specimens are fixed for 30 min to 1 hour at room temperature or for 1 to 4 hours at 0 to 4°C. After primary fixation, the fixative is removed with a fine pipette and transferred to a 500 ml bottle reserved specifically for fixation residues, ready for safe disposal as described in Sect. 2.2.2. The fixative is immediately replaced with a similar or larger volume of a suitable buffered washing solution. After washing and possible storage overnight in buffer,

Fig. 3.3 The equipment required for fixation by immersion. Pieces of tissue are placed in a small drop of fixative (d) on a sheet of dental wax, and are then cut into small fragments with a sharp razor blade. These fragments are transferred to a small glass vial (v) containing the fixative using a wide-mouthed pipette or a pair of tweezers.

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Fixation methods

the specimens are fixed with osmium tetroxide, washed, fixed with uranyl acetate and washed again, following a schedule of the type outlined below, all in the same vial. They are then ready for immediate dehydration as described in Chapter 4. The procedure is carried out in a fume cupboard and the solutions are added and removed with a fine pipette, and arc disposed of with the necessary safety precautions (see Sect. 2.2). During this series of solution changes the vials are kept on a tray to minimize the effects of any spillage. It is also possible to use a specimen rotator (Fig. 3.4) or an automatic processor, such as the typical unit shown in Fig. 3.5 (see Appendix), but these are not essential at this stage. The choice depends upon the type and number of specimens being handled, on the availability of the equipment, and on the personal preferences of the operator.

Fig. 3.4 A specimen rotator. Vials or tubes are accommodated on a head which is tilted at an angle of 45°. The head rotates at a variable speed and provides slow and gentle agitation of the specimens during fixation, dehydration and infiltration of embedding media. The Agar rotator holds up to 12 glass specimen vials and is small enough to be placed in a refrigerator. The rotating speed of the head is 2 to 26 rpm and alternative heads are available with clips to hold tubes of different sizes. (Photograph by DrJeremy Skepper.)

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Fig. 3.5 The Leica Lvnx EM Tissue Processor. This automatic processor is a selfcontained instrument for the fixation and dehydration of biological specimens, and for infiltration with the monomers of embedding resins. Specimens are retained throughout processing in tissue baskets, which are available with divisions of various sizes to suit different applications. Up to 20 reagent vials are located around a carousel and hold the sequence of solutions required during processing. The vials are sealed during operation to prevent evaporation of volatile reagents, and an integral fume extractor unit ensures that the operator is protected from harmful vapours when the instrument is loaded and unloaded. Agitation and temperature control are effected through a heater/cooler unit which can be programmed from 0 to 60°C. The Lynx processor is simple to operate and the user can store up to 10 different programs, each with 20 programmable steps. (Courtesy of I.eica, UK.)

3.4.1

A standard schedule for fixation by immersion

The sequence of fixation by immersion with an aldehyde, followed by osmium tetroxide and then by uranyl acetate, has become the standard schedule for ultrastructural studies of a great variety of specimens. A representative schedule for the fixation of tissue slices about 250 μιη thick by immersion is as follows: 1.

Fix for 30 min to 1 h at room temperature in a fixative containing 0.5 to 4.0% glutaraldehyde and 0.5 to 2.0% formaldehyde (prepared

Chapter 3:

Fixation methods

from paraformaldehyde), and 2 to 3 mM calcium chloride, in a 0.1 M cacodylate or PIPES buffer adjusted to a pH of 7.2 to 7.4, as described in Sect. 2.5.3a. 2. Wash for 30 min to 1 h at room temperature in three changes of 0.1 M cacodylate or PIPES buffer (as used for fixation) containing 2 to 3 mM calcium chloride at a pH of 7.2 to 7.4. Store in the same buffer solution overnight at 4°C, if necessary. Carry out any necessary dissection or slicing of the specimen at this stage in the schedule. 3. Post-fix for 1 to 2 h at room temperature in unbuffered 1.0% osmium tetroxide, or in unbuffered 1.0% osmium tetroxide also containing 1.5% potassium ferricyanide, and wash briefly in distilled water. 4. Treat for 1 to 2 h at room temperature in the dark with 0.5 to 1.0% aqueous uranyl acetate and wash briefly in distilled water.

3.5

The primary fixation of animal tissues

The relative merits of fixation of animal tissues by immersion and by perfusion have already been discussed in Sect. 3.1. Although several factors favour perfusion, there will be many occasions when the choice to fix by immersion will be made. It is the original method used and has the advantage of simplicity in the actual manipulations. Even when per­ fusion is used in the initial stage of fixation, the required specimen is often dissected out and the fixation continued by simple immersion, as described in Sect. 3.4. 3.5.1

Immersion fixation of animal tissues

Organs and tissues, such as skin, that can tolerate a temporary interruption to their blood supply and still retain their function and structure, are adequately fixed by immersion. The animal is killed and then the tissue is removed by dissection as rapidly as possible. A relatively large piece of tissue is taken and then cut into smaller pieces, the size of the pieces depending on the nature of the fixative and the density of the tissue. The cutting of the tissue into smaller pieces is done by placing the tissue on a sheet of dental wax in a fume cupboard, adding enough fixative to keep it moist, and then cutting it cleanly with a new single-edged razor blade (Fig. 3.3) or dissecting knife. Great care is

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required during cutting to ensure that the tissue is damaged as little as possible. As soon as the pieces are prepared they are transferred to prelabelled glass vials containing the primary fixative and processed by the method described in Sect. 3.4. 3.5.1 a

Preparation of thin tissue slices

As explained in Sect. 2.1.3, diffusion of fixative into tissues is a slow and complex process, so much so that pieces of tissue for immersion fixation should ideally be thin slices, no more than a few hundred micrometres thick. One simple method of cutting thin slices was described by Lewis and Knight (1992) for fixed tissue and can be adapted for unfixed tissues with a firm texture: 1.

Construct a small hand microtome by cementing two standard coverslips onto a microscope slide with epoxy or cyanoacrylic adhesive, orientated to leave a space between for the tissue, as shown in Fig. 3.6.

Fig. 3.6 A diagram of a simple device for cutting slices of tissue. The upper part of the diagram shows the device in plan view at approximately actual size (scale marker denotes 10 mm). The lower part of the diagram shows a transverse view along the line A-A and is not to scale in the vertical direction. The device consists of a glass microscope slide (1 mm in thickness) onto which are cemented three pieces of glass to form a recessed area for a block of tissue. If the pieces of glass are from thin coverslips, tissue slices can be cut at a thickness of 150 to 250 μηι (scale marker equivalent to 0.6 to 1.0 mm). The method of using the device is explained in the text.

Chapter 3:

Fixationmethods

2.

Place the slide in a Petri dish and cover the slide and coverslips well with buffered saline.

3.

Gently press a clean-cut face of a piece of fresh tissue downwards against the slide, carefully positioning it between the two coverslips. Draw a sharp razor blade across the tissue block in a sawing motion with the ends of the blade held flat against the coverslips.

4.

The thickness of the coverslips determines the thickness of the slices which can be cut. With practice 3 to 6 slices can be cut from fresh blocks of many tissues at a thickness of 150 to 250 μπι. Firm tissues, such as kidney, muscle or adrenal, give the most satisfactory slices. A more convenient, but much more expensive, option is to use one of the several types of mechanical devices which have been produced for cutting slices of tissue. Probably the best choice is a Vibratome (see Appendix), which is essentially a vibrating microtome and is illustrated in Fig. 3.7. To use the Vibratome the underside of the specimen is blotted dry

r'.'/i

Fig. 3.7 The Vibratome® Series 1000 Sectioning System. An arm carries a vibrating razor blade. The rate of forward movement of the arm is regulated by the potentiometer control at the top left-hand corner and the amplitude of the transverse vibration by the right-hand control. The height of the specimen stage is controlled by the knob at the bottom of the picture. (Photo courtesy of Ted Pella, Inc.) (Vibratome is a registered trademark of Technical Products, International.)

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and placed on a chuck coated with a thin smear of cyanoacrylate adhesive. The chuck is then clamped in the jaws of the specimen stage and the bath is filled with chilled buffer solution to cover the specimen. A razor blade is held horizontally on an arm which is vibrated rapidly from side-to-side across the line of advance so that sections are cut with a sawing motion. The specimen stage is raised mechanically after each return stroke of the blade by an amount which determines the thickness of the next section. The Vibratome has many advantages over other machines; a disadvantage, apart from its cost, is that sections have to be cut individually and therefore rather slowly. It is, however, highly versatile. Fixed and fresh tissue can be cut at a wide range of thicknesses and the block face can be as large as 20 mm square, which is an enormous advantage when selecting areas for electron microscopy from heterogeneous tissues, such as brain or kidney. Other instruments, all working on this same general principle, are also available. 3.5. Ib

Orientation of tissue pieces and slices

After primary fixation of tissues the fixative is removed and is im­ mediately replaced with a similar volume of a suitable washing solution (Sect. 3.4). At this stage further dissection or slicing may be conveniently carried out, if necessary. This is also by far the best stage at which to cut the tissue into a shape which makes it easy to orientate the specimen for embedding. This is a particularly important requirement when it is necessary to select a specific region of a heterogeneous tissue for ultramicrotomy. The specimen, preferably in the form of a thin slice, 200 to 250 μηι thick, is viewed by a suitable method of light microscopy and the region of interest identified. If this proves impossible, a very dilute solution in buffer of a dye such as Methylene blue can be applied with a small brush to the upper surface of the specimen, which is then viewed with top lighting. The area of interest is identified and then the slice is cut with a sharp razor blade into a shape that is easy to recognize after em­ bedding. The slice is trimmed so that the chosen area is at the corner opposite the most acute-angled corner, as illustrated in Fig. 3.8. Since the shape of the slice is asymmetrical, it is possible to identify which surface is which of the original slice, whichever way the slice happens to have been embedded. The large size of the slice enables it to be manipulated with curved tweezers during processing.

Chapter 3:

Fixation methods

4 to 5 mm

E E

Identified area

CO

O

CNJ

Fig. 3.8 An example of the type of shape which enables a specific area of tissue slice to be located unambiguously after the slice has been embedded. The tissue slice is trimmed with a razor blade so that the area of interest is at the corner opposite the acute-angled corner. Since the shape is asymmetrical it is possible to know which surface is which of the original slice, whichever way up the slice happens to have been embedded.

3.5.2

Surface application of a fixative

The fine structure of many animal tissues is critically dependent upon a continuous blood supply and it is essential that fixation of such tissues should be started while the animal is still alive. One way of achieving this is to drip the fixative onto the exposed tissue in an anaesthetized animal. Obviously the method is only useful if the superficial layers of the tissue are being studied and then only if there is no well-developed capsule around the tissue to impede diffusion of the fixative. The capsule can be slit open, but this may damage the underlying tissue. The method is per­ haps most useful for the study of the underside of skin, of skeletal muscle, of tendons and many other connective tissues, of the muscle layers in the intestine and similar structures. Some preliminary dissection is usually necessary and this must be carried out very carefully to avoid damaging the tissue and to preserve the vascular and nerve supplies intact. As soon as the tissue is suitably exposed in a fume cupboard, flood it with an ample supply of a fixative cooled to 0 to 4°C in order to reduce to a minimum the continuation of any metabolic activity in the tissue. For the same reason the blood supply to the tissue should be stopped, if possible, as soon as the fixative has been applied. The choice of fixative is the same as for immersion fixation but higher concentrations should be used to aid in­ ward diffusion. A suitable fixative is 4% glutaraldehyde plus 4% form­ aldehyde in 0.1 M cacodylate or PIPES buffer at pH 7.4. The addition of formaldehyde is important because of its more rapid penetration. Higher concentrations of the aldehydes can also be used to improve penetration of the fixative. Keep the tissue continuously bathed with fixative for about

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20 min by adding drops of fixative every few seconds either manually or through a gravity feed. Sometimes the tissue can be covered with a thin layer of cotton wool after a couple of minutes and the rate of adding fixative reduced to a few drops per minute, so long as the tissue surface can still be kept well moistened. As excess fixative accumulates, withdraw it with a pipette or soak it up with a pad of cotton wool. It may be an advantage to tilt the animal slightly to allow excess fixative to drain away from the area being fixed. At the end of 20 min dissect the tissue and cut it into pieces in such a way that the orientation of the well-fixed superficial layer is preserved and can be readily identified. Fix these pieces by immersion in the same fixative in small vials for 1 hour at room temperature or 1 to 3 hours at 4°C and then process them in the same way as described in Sect. 3.4. The whole operation must be conducted in a fume cupboard and great care must be taken to minimize personal exposure to the fumes from the fixative. 3.5.3

Fixation by injection

The internal structure of an organ or tissue can be fixed by the injection of fixative through a hypodermic needle. The method of preparing the tissue for fixation is the same as described in the previous section on superficial fixation, and the same glutaraldehyde-formaldehyde fixative is suitable. Inject the fixative with a hypodermic syringe over a period of 5 to 20 min. Carefully dissect out the region of tissue around the injection site, making sure that only well-fixed tissue is included. Prepare suitably sized slices and fix them in the same fixative for 1 hour at room temperature or for 1 to 3 hours at 0 to 4°C, transfer to a washing solution and then process them in the same way as specimens fixed by immersion (Sect. 3.4). This method is especially suitable for hollow structures such as the stomach, bladder or lengths of intestine. After the injection has been in progress for 2 or 3 min, cut a small hole well away from the injection site to allow the escape of excess fluid. The inner surfaces of hollow struc­ tures are well fixed by this technique.

3.6

Thefixationofinvertebrates

The methods which have been used for the fixation of tissues from invertebrates are many and varied. Typically small insects, such as moths or flies, are killed by decapitation, the body pinned out on a cork board

Chapter 3:

Fixation methods

and the required tissues dissected out for fixation by immersion in a standard aldehyde fixative of the type already recommended in Sect. 3.4.1. The insects can often be cooled beforehand to reduce the rate of tissue metabolism without damage to their ultrastructure. For aquatic creatures the ideal anaesthetic is MS 222 (tricaine methane sulphate), but often no anaesthetic is used. Thus clams can be fixed by immersion in sea water containing a few percent of an aldehyde as the fixative. Small specimens of, say, nematode worms are fixed by direct immersion in a suitable standard fixative. The more direct the application of the fixative the better, and the same fixing solutions as recommended in Sect. 3.4.1 are generally suitable.

3.7

The fixation of botanical specimens

3.7. I

Fixation procedures

It used to be believed, that in comparison with animal tissues, botanical tissues were relatively unaffected by removal from the plant, but it is now well recognized that ultrastructural changes occur quite rapidly, mainly due to water loss. Pieces of leaves, stems, flowers or roots are therefore fixed immediately after removal from the plant. In fact, it is sometimes advisable to start the fixation before the required tissue has been completely removed, or before dissection of a specific group of cells is attempted. The fixation of plant cells presents certain problems not common in animal tissues. These problems arise largely from the impermeabilty of the cell wall and the presence both of the vacuole, containing materials very different from the cell cytoplasm (Fig. 3.9), and of intercellular air spaces. Growing, meristematic tissues, such as root tips, are usually easier to handle since the cell wall is more permeable and the vacuole smaller. For such tissues, the standard immersion fixation procedures advocated in Sect. 3.4 are directly applicable. For older, especially woody, tissues the standard procedure must be modified. The greatest problem is that such plant specimens float in the fixative, and it is generally agreed that for adequate fixation to occur excess air must be removed by vacuum treatment in order to make the individual pieces of tissue sink. The vacuum should be applied gently. It should be increased gradually over a couple of minutes and then held at a

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Fig. 3.9 An electron micrograph of a typical plant cell with a thick cell wall and a large cytoplasmic vacuole. The arrows indicate microtubules. G, Golgi apparatus; N, nucleus. Scale bar = 1 μιτι. (Reproduced from Gunning and Steer 1996, with permission.)

vacuum not exceeding half an atmosphere (or 380 mm Hg) for a few more minutes. A single vacuum treatment should remove sufficient air from most specimens to allow them to sink in the fixative, but if they do not, the treatment should be repeated two more times over a period of half an hour. Any specimen which still will not sink should be rejected. For these more woody tissues, allowance must also be made for slower rates of diffusion of the fixative than in typical animal tissues. The times for the individual steps in the whole fixation procedure may have to be increased by a factor of up to four. Plant tissues must first be cut into slices thin enough to allow adequate penetration of the fixative. Slices should not exceed 1 mm in thickness and ideally much thinner slices should be cut, either free-hand

Chapter 3:

Fixationmethods

with a razor blade or mechanically with a Vibratome or other device, as described in Sect. 3.5.1a. When orientation is important it is best to cut the slices from an asymmetrically shaped block of tissue (Sect. 3.5.1b). If the fresh tissue has a waxy coat or cuticle, which would impede pene­ tration of the fixative, it should be stripped off first. Small specimens of plant material, such as algal unicells, spores and isolated chloroplasts, are fixed in suspension or as a centrifuged pellet as described in Sect. 3.10. 3.7.2

The choice of fixatives

The fixation mixtures favoured by botanists are varied in detail, but have an agreed basic composition. The biggest variations are in the osmolarity and the pH of the primary fixative. The question of the ideal osmolarity is even more difficult to answer than it is with animal tissues since it depends critically on the nature of the specimen and on the aim of the individual investigation. In general, however, the preferred mixtures are not very different from those favoured by electron microscopists working with animal tissues and the fixative of choice consists of glutaraldehyde or a glutaraldehyde-formaldehyde mixture in a buffered solution. Phosphate is usually preferred to cacodylate and PIPES is seldom if ever used. With cacodylate 2 to 3 mM calcium chloride can be incorporated in the fixative, but not with phosphate. Typically, the osmium tetroxide used in post-fixation is also buffered. It is reported that most plant material fixes best at a pH of 6.8, rather than 7.4 (Crang 1997), but it may be worth trying a higher pH for the initial stage of the fixation at least. As with other types of specimen, the best conditions for this initial fixation must to some extent be determined by trial and error. For most purposes fixation should be at room temperature and it can be extended to several hours. After two washes in buffer alone, tissues are normally treated with buffered 1% osmium tetroxide for an hour or more. Plant tissues are often treated with uranyl acetate before dehy­ dration and embedding. Potassium permanganate has been used as a fixative for plant material but has now been largely superseded by fixation with aldehydes, followed by osmium tetroxide and uranyl acetate. The main value of permanganate fixation now is where very waxy materials are involved, but massive extraction of intracellular material, especially nucleoproteins, occurs.

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A standard fixation procedure for plant tissues

In any particular investigation special methods may be necessary, but a basic procedure suitable for preliminary studies is as follows: 1.

Prepare a primary buffered aldehyde fixation as described in Sect. 2.4 and Sect. 2.5. Phosphate and cacodylate are suitable at a pH in the range 6.8 to 7.4 and at a concentration between 0.02 and 0.20 M, as appropriate for the tissue to be fixed. The aldehyde concentration should be in the range 2.0 to 6.0% (possibly up to 10%) glutaraldehvde, with or without the addition of 0.5 to 4.0% formaldehyde. A suitable fixative to begin with is one containing 2.5% glutaraldehyde plus 2% formaldehyde with 2 mM calcium chloride in

2.

0.1 M cacodylate buffer at a pH of 7.0. If it can be achieved without excessive damage, remove any cuticle or waxy coat which might impede penetration of the fixative and cut the tissue into slices, ideally not more than 1 mm thick.

3.

Transfer the slices to a small vial containing the fixative (see Sect. 3.4) and fix them at room temperature by immersion for 1 to 4 h, or even overnight for woody tissues. If the tissue floats, any pockets of air should be removed by applying a slight vacuum for a few minutes. The vacuum should be applied gently, and should not exceed half an atmosphere (or 380 mm Hg).

4.

Wash the slices for 20 min to 1 h in two changes of the same vehicle as used for the fixative, post-fix with buffered 1 % osmium tetroxide for 1 to 4 h, wash twice in distilled water and fix in 1% uranyl acetate for 1 h, all at room temperature.

The specimens are now ready, after two washes in distilled water, for immediate dehydration by the methods described in Chapter 4.

3.8

F i x a t i o n by v a s c u l a r p e r f u s i o n

Immersion fixation suffers from several disadvantages which have already been fully discussed in Sect. 3.1. Sufficient here to emphasize that for many animal studies fixation by perfusion, either of the whole animal or of isolated regions of tissue, such as the kidney, heart or lungs, is much

Chapter 3:

Fixation m e t h o d s

to be preferred. In this section the various perfusion techniques are des­ cribed, beginning with those for small laboratory animals, and followed by some specialized ones for particular tissues. Animals should always be perfused in a fume cupboard or under an extraction hood, and the operator must wear protective gloves and goggles, or preferably a face shield. Following fixation by perfusion, the selected tissue is dissected out and fixation can be continued by immersion (see Sect. 3.4). 3.8.1

An outline o f perfusion fixation

There are essentially two different types of technique available for fixation by perfusion. The one has a reservoir of fixative at a set height above the animal, which produces a non-pulsatile supply at a well-controlled pressure, while the other makes use of a roller-tube pump which produces a pulsatile delivery of fixative to the vascular system but at a pressure which may be difficult to control. Both have their advantages and dis­ advantages. The first method is by far the simplest and is the one most commonly used for routine fixation of tissues for electron microscopy, while the second gives a more realistic type of perfusion. The first is known as the constant pressure method of perfusion and the second can appropriately be called the constant volume method (Wisse et al. 1984). In both methods the animal is quickly, but fully, anaesthetized and the minimum amount of dissection carried out to insert a cannula in an appropriate blood vessel, as described later. The perfusion is begun with a suitable saline solution to wash out the blood, a major blood vessel on the venous side being cut to allow the blood to escape. This is followed by the fixative which is perfused for 10 to 15 min. The required tissues are then dissected out and are sometimes fixed further by immersion. The perfusion techniques in the following sections are basically designed for laboratory mammals. For birds, reptiles, amphibia, etc. much the same procedures can be used, but they may have to be modified to suit the particular species being fixed. 3.8.2

T h e choice o f anaesthetic

Many anaesthetics have been used for perfusion of mammals. For small rodents, such as rats or mice, ether inhalation is often chosen although a non-explosive gas, such as chloroform, is much to be preferred. For larger animals an injected anaesthetic is more convenient. The choice of

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anaesthetic can be very important since significant ultrastructural changes can be induced during the few minutes between administering the anaesthetic and beginning the perfusion. The first important choice is between a volatile and a non-volatile anaesthetic. For practical convenience, if the chest cavity is to be opened, then a non-volatile anaesthetic should be used. For small laboratory animals, however, where the minimum of surgery is necessary before perfusion is begun, a volatile anaesthetic is often quite satisfactory, and easier to administer. Whenever thoracic surgery taking more than a couple of minutes is needed, a non-volatile anaesthetic is essential. Diethyl ether is still the most commonly used volatile anaesthetic, but its high flammability is an added danger, and it has an excitatory effect on much of the autonomic nervous system. Chloroform is a more satisfactory volatile anaesthetic. The animal is more gently sedated and the effects on the peripheral nervous system are much less. Expensive volatile anaesthetics, such as Halothane, offer no worthwhile advantages. The animal should be placed in a suitable chloroform box until anaesthetized and the required level of anaesthesia maintained by a pad of cotton wool moistened with chloroform placed over the nose of the animal. It is most convenient to place the pad at the bottom of a suitably sized glass beaker, and to adjust the position of the beaker to keep the desired level of anaesthesia. A fume cupboard or other extract system is essential to remove the chloroform or ether vapour. It should be remembered that the unwanted carcass still contains much excess anaesthetic and it should be disposed of with care. If ether is used, disposal is particularly hazardous because of the explosion risk. The most commonly used non-volatile anaesthetic is pentobarbitone, sometimes known as Nembutal. Intraperitoneal injection is most popular, but intravenous injection is better, particularly in animals larger than the rat. A suitable dose is in the range of 30 to 50 mg/kg body weight. Fish and amphibia are transferred to water containing the anaesthetic MS 222 at a concentration between 0.02 and 0.05%, the precise con­ centration depending on the species. In general, young animals need a higher concentration, even as high as 0.10%. The animal should be exposed to the anaesthetic for 5 to 10 min or until it is fully anaesthetized and insensitive to touch. If surgery is prolonged beyond a few minutes,

Chapter 3:

Fixationmethods

some method of continuing the exposure to anaesthetic should be arranged, either through the gills or the skin. 3.8.3

The choice of perfusion fluids

In addition to the aldehyde fixative itself, the perfusion fluids for primary fixation must also have an adequate buffering capacity, a suitable osmolarity and an appropriate content of divalent cations. The same attributes must also be possessed by the initial saline solution perfused in advance of the fixative. Buffering capacity is essential because acid is released by the reaction of aldehydes with proteins. Acid production is particularly large when glutaraldehyde is used as the sole fixative because of the production of pyridine derivatives which uses up oxygen and releases extra acid, as explained in Sect. 2.4.1a. It is therefore desirable to have 0.1 M buffer in the perfusion fluid: cacodylate, phosphate and PIPES are all suitable for buffering the acid released by aldehyde fixation. The precise initial pH is not critical, but a value of 7.2 to 7.4 is usually chosen. The question of the total osmolarity of the fixing solution has already been fully discussed in Sect. 2.3.2. For perfusion fixation, however, account must also be taken of the colloid osmotic pressure. A suitable substitute for the plasma proteins must be found that does not react with the aldehyde fixative. A popular substitute is dextran, which is a medium molecular weight (MW) polysaccharide made by the controlled hydrolysis of starch. Even more popular is polyvinylpyrrolidone (PVP), a wholly synthetic polymer which is highly soluble in water and saline solutions. Samples with a mean molecular weight in the range 40,000 to 80,000 daltons are generally used. Divalent cations are important for the preservation of the semi-permeability of cell membranes and for the maintenance of the ultrastructural integrity of lipid membranes. Both requirements are normally satisfied by the inclusion of 2 to 3 mM calcium chloride in all the perfusion fluids. Some workers use a vasodilator substance to prevent premature collapse of blood vessels during perfusion. The simplest, and a very satisfactory, dilator is sodium nitrite, which is added to the initial saline solutions, usually at a strength of 5 mM. Other vasodilators, under various trade names, are available if preferred. It should be borne in mind, however, that the addition of a vasodilator may alter the ultra­ structural appearance of the tissue.

ΙΟΙ

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A typical formulation for the saline used as the pre-fixation perfusion fluid is: 10 mM PIPES buffer at pH 7.4, containing 140 mM sodium chloride, 2.7 mM potassium chloride, 2 mM calcium chloride, 5 mM sodium nitrite, 20 mM glucose and 2.5% PVP (MW = 40,000 daltons). For fixation a suitable perfusion fluid is: 2.0 to 4.0% glutaraldehvde and 0.5 to 4.0% formaldehyde in 0.1 M PIPES buffer at pH 7.2 to 7.4, containing 2 mM calcium chloride and 2.5% PVP. For some purposes much simpler formulations can be used, omitting, for instance, the sodium nitrite, glucose and PVP from the pre-fixation fluid, and the PVP from the fixation fluid. There is no general agreement about how much saline should be used to wash out the blood before introducing the fixative. A volume equal to about 50 to 100% of the total blood volume of the animal should be sufficient, which implies a number of millilitres roughly equal to 5 to 10% of the weight of the animal in grams, perfused over a period of a half to one minute. If a suitable diameter of tubing is chosen, the volume of saline between the tap on the reservoir and the tip of the cannula can be made appropriate for the animal being perfused. Per­ fusion of the fixative should be continued until a volume equivalent to the weight of the animal has been used, which should take about 10 to 15 min. For perfusion of separate organs, the volumes required will need to be adjusted, but the times required should be about the same. There is no consensus about the temperature for perfusion. For best ultrastructural results the initial saline should probably be warmed to about 45°C before being introduced into the tubing so that it enters the animal at about body temperature. The fixing solution itself, however, is usually introduced at room temperature, and for some specialized cytochemical studies may even be pre-cooled to 4°C. 3.8.4

Apparatus for perfusion fixation

For the constant pressure method of fixation the apparatus required is extremely simple. It consists of a suitable reservoir, such as a separating funnel or a storage bottle with a lower outlet, which should be fitted with a tap and be large enough to hold all the fixative required for one perfusion. The outlet of the reservoir is connected bv a long length of plastic tubing to a hypodermic needle or a cannula. The tubing is filled with saline or other blood substitute up to the level of the tap, and the reserv oir is filled with the fixing solution (Fig. 3.10).

Chapter 3:

Fixation m e t h o d s

Gauge (b)

Roller pump

Fig. 3.10 The apparatus for perfusion fixation. For small laboratory animals, such as rats and mice, the reservoir of fixative (a) is connected by plastic tubing to a hypodermic needle (b). The reservoir is suspended 1.2 to 1.5 m (4 to 5 ft) above the needle, and the plastic tubing is filled with a saline solution. For larger animals, such as rabbits, it is more satisfactory to use two reservoirs (c) connected via a three-way tap to the needle (b), again at a height of 1.2 to 1.5 m. For special purposes a roller pump can be connected as shown in (d) between the needle and the pair of reservoirs, which should then be just above bench height.

Some research workers use two reservoirs, one containing the fix­ ation fluid and the other the pre-fixation saline, with the two reservoirs connected to the cannula via a three way tap, as illustrated in Fig. 3.10. This arrangement is very useful for larger animals when the volume of pre-fixation saline needed is greater than can conveniently be accomm­ odated in the tubing supplying the cannula. The perfusion pressure must be high enough to ensure a good flow of fixative but not significantly higher than the arterial pressure in the living animal. With an intra-arterial gravity feed, the level of the reservoir of perfusion fluid should be 1.2 to 1.5 m (4 to 5 ft) above the animal, but when an intravenous route is used, as in perfusion of the liver via the portal vein, the fluid level should be only 200 to 300 mm (8 to 12 in) above the animal. For the roller-pump method of perfusion, a reservoir large enough for all the fixative required should be connected via a tap to an electrically operated roller-tube pump. The pump is connected to a hypodermic

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needle or cannula via an adequate length of plastic tubing. It is a great advantage to include some form of pressure gauge between the pump and the needle. An ordinary mercury manometer is suitable for intra-arterial perfusions, while a water manometer is better for intra-venous ones. As with the constant pressure method, the apparatus should be filled as far back as the reservoir tap with saline or other blood substitute. This apparatus is illustrated in Fig. 3.10. 3.8.5 T h e r o u t i n e perfusion o f m a m m a l s The same basic techniques can be applied to a range of laboratory animals. Following anaesthesia (Sect. 3.8.2) a small quantity of a suitable saline solution is infused, followed by a much larger volume of a fixing solution over a period of about 10 min. For very small animals, such as mice or young rats, perfusion is best made directly into the left ventricle through an ordinary sharp hypodermic needle. For rats, guinea pigs and similarly sized animals, perfusion is best made with a blunt needle, Table 3.1 Perfusiontechniques Tissue

Speaes

Reference

General

Small mammals

Rossi (1975)

Artery

Mammals

Haudenschild et al. (1972)

Aorta

Rabbit

Swinehan et al. (1976)

CNS

Small mammals

Furness et al. (1978)

Embrvo

Mouse

Abrunhosa (1972)

Heart

Rat

Forssmann et al. (1967)

Heart

Fetal lamb

Dae et al. (1982)

Kidney

Cat

Yun and Kenney (1976)

Liver

Rat

Wisse et al. (1984)

Liver

Birds

Bhatnagar et al. (1981)

Lung

Rat

Gil and Weibel (1969)

Lung

Cat

Coalson (1983)

Ovarv

Guinea pig

Paavola (1977)

Spleen

Rabbit

Elgjo (1976)

General

Fish

Hinton (1975)

CNS, central nervous svstem.

Chapter 3 :

Fixationmethods

inserted through a cut made in the wall of the left ventricle, and tied into the root of the aorta. For larger animals perfusion is best limited to only part of the vasculature, say by perfusion into the descending abdominal aorta with ligation of the blood supply to unwanted regions of the body. In addition to perfusion of the whole animal, various specialized perfusion techniques have been proposed for the fixation of particular individual tissues. Complicated surgery is often involved and the level of anaesthesia maintained (usually with Nembutal) is all important. A muscle relaxant such as Flaxadil is sometimes also advantageous. A high degree of surgical skill is often required and this must be gained before attempting to fix tissues for electron microscopy. The original literature should be first consulted and the appropriate references are included in Table 3.1. Further information for a few selected tissues and organs is given in Sect. 3.8.6 to Sect. 3.8.10. 3.8.5a A standard perfusion technique for small mammals

For small laboratory animals, such as rats or guinea pigs, the most satisfactory route for perfusion is via a cannula through the wall of the left ventricle into the base of the ascending aorta. For the perfusion of a mature laboratory rat the following procedure is recommended: 1. Assemble and fill the perfusion apparatus with saline and the chosen fixative (Sect. 3.8.3), preferably warming the saline to about 45°C before introducing it into the tubing, so that the initial flow of fluid will be near body temperature. 2. Immediately anaesthetize the animal as deeply as necessary (see Sect. 3.8.2). Inject a vasodilator beforehand, if desired, or add it to the perfusion fluids. 3. Lay the animal on its back in a shallow tray inside a fume cupboard. Wear protective goggles and gloves. 4. Make a midline incision down the whole length of the thorax and much of the abdomen and fold back the skin. 5. Expose the heart by cutting the ribs and intercostal muscles along each side and raising the flap so formed. 6. Clean away the thymus and excess adipose tissue from the arch of the aorta and draw a ligature beneath it, ready to tie the perfusion cannula in place.

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Make a stitch through the apex of the heart and clip the thread to the side of the animal so that the heart is under slight tension. Make a small perforation in the wall of the left ventricle with a pair of fine scissors and quickly insert the metal cannula into the opening and through the aortic valve so that the cannula projects about 5 mm into the aorta. Tie the cannula in place and as soon as the saline perfusion fluid is

flowing cut the right atrium. 10. Continue the perfusion until a volume of fixative equivalent to the weight of the animal has been used, which should take 10 to 15 min. Hiraoka and Wang (1989) describe a neat device for use in the perfusion of small mammals, such as rats and mice. 3.8.5b Perfusion techniques for larger animals

Whole animal perfusion of larger animals, such as rabbits, cats and dogs, is much less convenient because of the larger volumes of fixatives required. One compromise, when tissues are only needed from the head end, is to clamp off the abdominal aorta as high up as possible once the perfusion has begun. If only the brain needs to be fixed, the two carotid arteries can be cannulated directly and the jugular veins cut to allow perfusate to escape. If tissues from the lower part of the animal are required the abdominal aorta should be cannulated and the perfusion of unwanted structures stopped by clamping off the appropriate arteries. It may be best to carry out a close intra-arterial perfusion of the tissue required, that is, a perfusion via the specific arterial branch supplying the tissue region, but this may require delicate surgery involving a longer period of anaesthesia. A compromise is often necessary. 3.8.6

Perfusion of heart tissues

The perfusion techniques recommended in Sect. 3.8.4 and Sect. 3.8.5 may not deliver sufficient amounts of fixative to the heart tissues to produce optimal fixation. For small mammals it is best to carry out a retrograde perfusion from the abdominal aorta which is a large enough vessel to cannulate easily. For large mammals it is better to insert a catheter into a suitable artery, such as the femoral or a carotid, and advance the catheter, made of fine polyethylene tubing, until it reaches the heart. To prepare

Chapter 3:

Fixationmethods

the rat heart satisfactorily it is necessary to insert the cannula, pointing 'up-stream' towards the heart, into the abdominal aorta at the point between the bifurcations of the renal and iliac arteries. 3.8.7

Perfusionofthekidneys

Two particular problems arise in the fixation of mammalian kidneys for electron microscopy. The first problem is that the blood vessels in the kidney have a pronounced tendency to constrict in response to the slightest interference. A non-irritant anaesthetic (Sect. 3.8.2) must therefore be used plus careful preparatory surgery followed by a period of waiting to allow the circulation to recover and stabilize. The second problem is that some regions of the kidney may not be iso-osmotic with the general body fluids. For fixation of the cortex of the kidney, perfusion with an iso-osmotic fixative appears to give the best results (Yun and Kenney 1976; Elling et al. 1977), but for the medulla hyper­ osmotic solutions can give better preservation of ultrastructure. Trial and error may be the only way of discovering how to obtain the best results. A brief account of the necessary procedure is given by Hayat (1989) and a full description by Yun and Kenney (1976). 3.8.8

F i x a t i o n o f l u n g tissue

Fixation of the lungs presents two special problems. Firstly, opening the thoracic cavity causes the lungs to collapse and secondly, the lungs are supplied with blood from the right rather than the left side of the heart. The animal must be artificially ventilated before any surgery on the thorax starts; and the perfusion cannula must be inserted into the pulmonary artery. To prevent pulmonary oedema, the perfusion fluids, both the initial saline and the fixative, must have an appropriate colloid osmotic pressure, as discussed in Sect. 3.8.3, provided either by dextran or PVP, the appropriate concentration of which is best decided by trial and error (Coalson 1983). Lung tissue can also be fixed by instilling fixative through the trachea; that is, by replacing the air in the lungs by fixative. This method is not as 'physiological' as vascular perfusion, but is simpler and retains the vascular contents intact. It has been used by Weibel (1970) for morphometric studies of the lung vasculature and his paper should be consulted for full experimental details.

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Biological specimen preparation for TEM

Portal perfusion o f liver tissue

Liver holds a special place among tissues which do not lend themselves to whole animal perfusion fixation, because its major blood supply in the intact animal is a low pressure portal supply from the digestive tract. The very unsatisfactory fixation obtained by ordinary intra-arterial perfusion is well shown by the 'blotchy' appearance of the surface of the liver normally obtained when it is fixed by perfusion through the heart or the descending abdominal aorta. A better method of obtaining good, even fixation is by low-pressure perfusion through the main portal vessel, as described by Wisse et al. (1984). Bhatnagar et al. (1981) describe a method of perfusing the liver in turkeys or similar birds. This differs from the method for mammals because in birds the blood vessels to the liver are protected by the breast bone. 3 . 8 . 1 0 The perfusion o f fish

The recommended method of perfusion-fixation of fish is that described by Hinton (1975). The fish should first be placed in a freshly prepared solution of MS 222 at a dilution of 1 in 4000 in fresh water or sea water, as appropriate. Expose the fish to this solution until it is fully anaesthetized and insensitive to touch (usually 5 to 10 min). Soak some patches of gauze in the solution and position them under the gill cover (the opercula) and over the gill arches. Remove the fish from the water and place it on its back or side as appropriate to the anatomy of the species. Make a triangular pattern of incisions and fold back the skin and muscle tissue to expose the heart and proximal aorta. Make an incision into the proximal aorta and insert a length of polythene tubing of a bore suitable for the perfusion. In some species it may be easier to introduce a cannula into the aorta through an incision into the heart ventricle as recommended for small mammals in Sect. 3.8.5a. The volume of fixative needed will be discovered by ex­ perience, but will probably be something approaching the volume of the fish. The same aldehyde fixatives as recommended for mammals are suitable, but the osmotic composition of the fixative will need to match that of the blood of the fish.

Chapter 3:

3.9

Fixationmethods

Fixation of organ and monolayer cell cultures

Organ cultures are very easy to fix, particularly if they have been cultured on a Millipore filter, since all that is necessary is to lift them carefully from the medium in which they have been grown, rinse them in a balanced salt solution to remove excess medium and place them directly in the primary fixative in a small vial for fixation by immersion as described in Sect. 3.4. Since these specimens are very thin, a simple 2 to 4% glutaraldehyde fixative without any added formaldehyde is suitable, but it must contain 2 to 3 mM calcium chloride. Colonies of cells (tissue culture cells, bacteria, fungi, etc.) growing on agar or other solid medium in a culture dish are fixed by the same method. When cells have been cultured on a substrate rich in proteins (such as gelatin or albumin) it is necessary to include a sufficiently high concentration of buffer in the fixative to counteract the production of acid during the reaction between the proteins of the substrate and the glutaraldehyde. After the primary fixation, the solid medium, with the cells attached, is peeled off the culture dish and processed in a small vial as described in Sect. 3.4. Monolayers of cells growing on slides or coverslips or other substrates (Fig. 3.11) are very sensitive to changes in their environment. Consequently the substrate is placed in a small culture dish containing the growth medium so that the cells can be fixed in situ at the tem­ perature at which they were incubated. The procedure is as follows: 1. Warm the primary fixative in the incubator. It is helpful to have a small incubator in a fume cupboard, and special precautions must be taken to avoid inhalation of the fumes of the fixative. A standard glutaraldehyde fixative, containing calcium chloride, is suitable (Sect. 2.4.2). 2. Fix the monolayer of cells by one of the following methods: a. Add sufficient warm 25% (w/v) aqueous glutaraldehyde to the medium in the culture dish to obtain a final concentration of glutaraldehyde of 2.5%, replace the lid of the culture dish, and rock the dish gently to mix the fixative and growth medium; or b. Add a volume of warm medium containing 5% glutaraldehyde equal in volume to the culture medium; or

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for TEM

Chapter 3:

Fixation methods

c.

Remove the culture medium gently with a pipette and replace it with a similar volume of warm buffered fixative, consisting of 2.5% glutaraldehyde in 0.1 M cacodylate or PIPES buffer, pH 7.4, containing 2 to 3 mM calcium chloride. 3. Fix the monolayer for 15 to 30 min at the incubation temperature and then remove the culture dish from the incubator. 4. Leave the substrate and cells in the culture dish during further processing or transfer them to a glass vial containing buffer. After washing, the fixed monolayer can be stored at 4°C in buffer, so long as the buffer contains calcium chloride to stabilize lipids. 5. Fix the cells with osmium tetroxide and then with uranyl acetate (Sect. 3.4). They are then ready for dehydration as described in Chapter 4. Although external cell morphology often changes dramatically fol­ lowing a drop in temperature, intracellular organelles are not necessarily so seriously affected. For some experimental studies therefore it may be convenient to allow the culture to cool for a couple of minutes and then to use the primary fixative at room temperature.

3.10

Fixation o f isolated cells

The fixation methods for isolated cells (cells cultured in suspension, protozoa, bacteria, blood cells, etc.) are very different from the methods used for tissues. There is now no problem in obtaining good penetration of the fixative, but it is sometimes difficult to handle the cells, particularly if only small quantities are available, and the composition of the primary

Fig. 3.11 Monolayers of human monocytes incubated on agar layers and fixed in situ by the addition of 1 ml of warm 25% glutaraldehyde to each 10 ml of the growth medium. The agar layers were supported on thin Araldite coverslips to aid subsequent embedding in Araldite. (a) Light micrograph of a 1 μπι thick section of monocytes incubated on an agar layer (Ag) for 30 min. Stained with Toluidine blue. Scale bar = 20 μπι. (b) Electron micrograph of an ultrathin section of a monocyte adhering to the agar (Ag) surface. Scale bar = 1 μιτι. (Reproduced from Eccles and Glauert 1984, with permission.)

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fixative must be chosen with care. Glutaraldehyde is most commonly used, at a concentration of 0.5 to 4% in a 0.1 M cacodylate or PIPES buffer containing 2 to 3 mM calcium (Sect. 2.4.2). The temperature during fixation should be the same as that of the isolated cells since changes of temperature can, for instance, rapidly affect the external morphology of mammalian cells. Consequently, cells cultured in suspension are fixed at the incubation temperature, while cells which have been obtained by an isolation procedure are fixed at the final temperature of that procedure. This will usually be room temperature or 4°C. Samples of isolated cells are handled in one of two ways. They are fixed in suspension and then centrifuged down to form a pellet, or they are centrifuged gently first and the pellet is fixed by immersion (Sect. 3.4). Cell fractions obtained by ultracentrifugation or other means can be handled in the same two ways. 3.10.1

Fixation o f cells in suspension in medium

For most isolated cells good fixation is best achieved by adding the fixative while the cells are still in suspension in medium (see Fig. 3.12). Any changes to the ionic environment of the cells are minimized and dif­ fusion problems are avoided. The simplest method is to prepare some of the suspending medium and add to it double the amount of aldehyde or aldehydes to give the final required concentration. Then add equal volumes of this double-strength fixative and the cell suspension to a centrifuge tube and mix them by gently swirling and tipping the tube. In this way the correct composition of the fixing medium is automatically achieved. This procedure is usually carried out at room temperature, although for some types of cells a lower or higher temperature may be required. Because diffusion is not a problem, the concentration of the fixing agents can be lower than those appropriate for tissues and the fixation times can be shorter, but neither of these parameters should be reduced by more than a factor of two or three, otherwise the amount of cross-linking and even the type of cross-linking may be affected. Probably the final concentration of glutaraldehyde ought not be reduced below 0.5%, and the fixation time at room temperature should not be shorter than 10 to 20 min. After fixation the cell suspension is centrifuged into a pellet, the fixative is carefully removed with a fine pipette and is replaced with a

Chapter 3:

Fixationmethods

Fig. 3.12 Rabbit platelets fixed in suspension by the addition of 10 volumes of 2.5% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.2, containing 3 mM calcium chloride. After fixation for 30 min at room temperature, the cells were centrifuged into a pellet before post-fixation in osmium tetroxide and uranyl acetate, dehydration in ethanol and embedding in Araldite. Scale bar = 1 μπι. (Reproduced from MacIntyre et al. 1977, with permission.)

buffered washing solution. The cohesion of the fixed pellet depends on the type of cell, the composition of the fixative and the speed of centrifugation, which should be kept to a minimum to avoid distorting the cells. Many pellets which have been fixed in glutaraldehyde are firm and show no tendency to disintegrate in the buffer, and these can be removed from the centrifuge tube at this stage. The fact that the pellet has to be removed in this way should be borne in mind when selecting the centrifuge tube (the standard plastic Eppendorf tubes are ideal). The pellet can usually be freed from the bottom of the tube fairly easily using the end of a fine spatula. If the pellet is difficult to dislodge it is preferable to use a plastic tube which can be cut down to release the pellet. After removal from the centrifuge tube, the pellet can be placed in a small drop of buffer on a sheet of dental wax and cut into small pieces by the method described in Sect. 3.5.1. Alternatively, if the pellet is large

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enough, it can be cut into thin slices to enable rapid penetration of fixatives (Sect. 3.5.1a). Care should be taken to cut out samples from different regions of the pellet so as to check the homogeneity of the original suspension of cells. These samples are placed in small vials where they remain during the subsequent steps of the preparative procedure (Sect. 3.4). Post-fixation in 1% osmium tetroxide at room temperature for 1 hour followed by 1% aqueous uranyl acetate for 1 hour, also at room temperature, is usually adequate. The use of uranyl acetate is strongly recommended in the study of the ultrastructure of isolated cells. 3.10.2

F i x a t i o n o f cells a n d o r g a n e l l e s i n a p e l l e t

Some types of isolated cell, such as bacteria and erythrocytes, and cell organelles, such as mitochondria, can be fixed adequately after they have been centrifuged into a pellet. The supernatant is removed carefully without disturbing the pellet and then an aldehyde fixative is added. Again the speed of centrifugation should be kept to the minimum required to obtain a cohesive pellet. The fixative should be at least twenty times larger in volume than the pellet. It is run slowly down the side of the centrifuge tube keeping the pellet intact. Primary fixation for 30 min to 1 hour in a standard buffered glutaraldehyde-formaldehyde fixative, containing calcium chloride (Sect. 2.5.3a) is suitable for most pellets, while the formaldehyde can be omitted when the pellet is small. Pellets centrifuged at too high a speed become very compact, so that fixatives penetrate much more slowly than into a sample of tissue which may contain considerable extracellular space. The fixation time for these pellets should be increased, possibly up to 4 hours. For the same reason, the contents of larger pellets, more than 1 or 2 mm in depth, may need to be resuspended in the fixative by gentle shaking to ensure that the whole pellet is adequately fixed. After primary fixation the cells or organelles are then centrifuged into a pellet again before the next step in the preparative procedure. After the primary fixation is complete, the fixative is carefully removed from the pellet and replaced with a buffered washing solution and the pellet is processed in the same way as pellets of cells fixed in suspension (Sect. 3.10.1).

Chapter 3:

3.10.3

Fixationmethods

Encapsulating methods for pellets

Only certain favourable pellets are cohesive enough to remain intact after the initial fixation; others tend to disintegrate when the buffer is added for washing and material is easily lost. When this is likely to happen the pellet is 'encapsulated' by surrounding it with a soft gel of agarose or calcium alginate. The method has been used successfully for a wide range of specimens, including mammalian oocytes, plant and animal cells, protozoa, yeasts, bacteria and cell organelles, such as mitochondria. The pellet can be encapsulated at any stage before dehydration has begun, but it is advisable for it to be done as soon as possible to avoid having to centrifuge the cells to a pellet at each stage of the procedure. Agarose. Earlier methods of encapsulating pellets involved the use of agar (Glauert 1975) which solidifies at 55°C. Consequently the procedure has to be carried out as rapidly as possible to avoid premature setting of the agar and the high temperature causes structural damage to many types of cell (Wood and Klomparens 1993). The introduction of the low-melting point agaroses, such as Sea Prep, Sea Plaque and Sigma types VII and IX (see Appendix), which remain fluid at 32 to 37°C, has enabled lower temperatures to be used and simpler methods of encapsulation to be developed. A typical procedure is as follows: 1.

2. 3.

4. 5.

6.

Prepare a 2 to 4% solution of agarose by dissolving the agarose in distilled water or buffer at 37 to 42°C. Pour the solution into a test tube and place the tube in a water bath at a temperature just high enough for the agarose solution to remain fluid. Place a centrifuge tube containing the pellet of aldehyde-fixed cells in the same water bath. Transfer a small drop of agarose (approx. 0.03 ml) to the centrifuge tube with a warm pipette and shake the tube gently to suspend the cells in the agarose. Immediately tilt the tube so that the agarose runs down the side of the tube to form a drop on a cool glass microscope slide. After the agarose has set (1 or 2 min), cut the solidified agarose containing the cells into small cubes, about 1 mm 3 , or preferably into slices about 250 μηι thick, with a sharp, single-edged razor blade. Process these cubes or slices in the same way as pieces of a cohesive pellet (Sect. 3.10.1).

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Fig. 3.13 Sections of a pellet of mouse peritoneal macrophages encapsulated in agarose, showing (a) the cohesion of the cell pellet, and (b) detail of the cytoplasm of a cell. Scale bars: (a) 5 μιη; (b) 0.5 μηι. (Reproduced from Strausbauch et al. 1985, with permission.)

It is important not to add too much agarose or the suspension will be diluted and few cells will be visible in each section. Some workers recommend centrifuging the solution at the end of Step 3. For example, Strausbauch et al. (1985) recommend transferring the solution to conicaltipped BEEM capsules. The capsules are immediately centrifuged for 5 min at 9,000 g, using a micro-centrifuge with a modified head, or an adapter, such as the one described by Sparkman et al. (1987), so that compact pellets are formed. The capsules are then cooled on ice for 15 min, the tip of each capsule is cut off with a razor blade and the cell pellets are removed with a needle and cut into small pieces for further processing. With this method it is possible to observe many cells in a single section, as illustrated in Fig. 3.13. Calcium alginate. A gel of calcium alginate, which is the salt of a polymannuronic acid obtained from seaweed, is formed when sodium alginate comes into contact with calcium ions and this enables pellets to be encapsulated at any temperature between 0 and 25°C. The main diffi­ culty is control of the total osmotic pressure during the gelling process, which could be a problem with unfixed tissue.

Chapter 3:

Fixation methods

Fig. 3.14 A transverse section of a Trypnanosoma brucei cell fixed in suspension in 3% buffered glutaraldehyde and then encapsulated in calcium alginate. Scale bar = 0.2 μτη. (Reproduced from Page et al. 1994, with permission.)

The procedure described by Page et al. (1994) for trypanosomes (see Fig. 3.14) is as follows: 1. Prepare a 5% solution of sodium alginate (see Appendix) by gently sprinkling 1 g of the powder into 20 ml of distilled water whilst stirring, preferably with a magnetic stirrer. The solution quickly becomes very viscous and must be stirred gently for at least 30 min to ensure that all of the powder is dissolved, otherwise the cutting quality of the final block will suffer. The solution can be frozen in 1 ml aliquots and defrosted just before use. For some specimens concentrations of sodium alginate as low as 2% give satisfactory results. 2. Mix the aldehyde-fixed pellet with an equal volume of the 5% sodium alginate solution and then draw up 15 μΐ of this mixture into a micropipette (200 μΐ capacity) and extrude the droplet into a buffer, such as 0.1 M PIPES buffer at pH 7.2, containing 0.1 M calcium chloride, in a small vial. Care must be taken to wipe the tip of the pipette after drawing up the alginate/trypanosome suspension to prevent the drop sticking to the end of the tip on extrusion. Repeat with further droplets.

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3. Once in the buffered calcium chloride the 15 μΐ drops form small beads which gel in approximately 15 min, forming a strong polymer matrix of calcium alginate containing the fixed cells. Post-fix and process the beads in the same way as samples fixed by immersion (Sect. 3.4). 3.10.4

C o l l e c t i n g cells a n d v i r u s e s o n M i l l i p o r e filters

A convenient method for collecting isolated cells and viruses for fixation is to use a Millipore membrane filter. Filters with various pore sizes are available (see Appendix), 0.45 μπι being suitable for mammalian cells and some bacteria, 0.22 μπι for smaller bacteria and 50 nm for viruses. Cells are collected, either by filtering the suspension through the Millipore mem­ brane in the usual way, or a sterile filter is placed in a small culture dish and the concentrated suspension is placed gently in the centre. The filter is laid on several layers of absorbent (blotting) paper to soak up and draw through the medium by surface tension. McCombs et al. (1968) incubated the filter at 37°C for 2 to 3 hours to allow the cells to attach firmly to the filter. The cells on the filter are fixed in a small vial (Sect. 3.4) and dehydrated by standard methods and then the filter is cut into pieces before infiltration with the embedding medium. Propylene oxide may dissolve the filter and so xylene or toluene are used as intermediate solvents.

3.11

Fixation by microwave i r r a d i a t i o n

Microwaves are electromagnetic waves with frequencies between 300 MHz and 300 GHz, corresponding to wavelengths between 1 m and 1 mm, respectively. Domestic microwave ovens operate at 2450 MHz, corresponding to a wavelength of 122 mm, and similar radiation appears to be provided in the ovens used in microscopy. At this frequency microwave irradiation causes dipolar molecules, such as water, to rotate and ions to move back and forth in a liquid medium, but molecules are not ionized and even the weakest chemical bonds are not disrupted (Login and Dvorak 1994b). Microwaves greatly increase the rate at which both glutaraldehvde and formaldehyde diffuse into tissues and consequently some initial ultra-fast fixation can be obtained by irradiating a biological specimen immersed in an ordinary aldehyde fixative for only a few seconds. This initial treatment is then followed bv the standard immersion

Chapter 3:

Fixation methods

fixation schedule given in Sect. 3.4.1. Although this combined technique can give satisfactory preservation of ultrastructure, in practice the results obtained routinely are not always as good as one might hope. One reason may be a lack of uniformity in the local intensity of the radiation within the oven. It is essential to use a microwave oven specifically designed for laboratory use (see Appendix), and the oven has to be calibrated to identify positions of even heating before each fixation procedure. The immersion procedure following the initial microwave-chemical fixation can, of course, be modified when, for instance, the primary consideration is not the pre­ servation of ultrastructure but the application of some cytochemical technique. The microwave oven can then have great advantages, but for ultrastructural studies the advantages are less obvious. In fact, too little is known about the effect of microwaves on tissue to warrant their recom­ mendation for routine use in ultrastructural studies for the present. Anyone contemplating the use of microwaves for electron microscopy should first consult the excellent reviews by Kok and Boon (1990) and Login and Dvorak (1994a). If it is decided to proceed, more detailed information is available in the book by Kok and Boon (1992), and a good source for practical details, including advice on safety precautions, is the book by Login and Dvorak (1994b). Microwave ovens must be checked regularly with a detector for leakage around the door seals.

3.12

Fixation methods for specialized techniques

Cytochemical and immunocytochemical techniques usually require milder fixation procedures than are standard for ultrastructural studies. This is because the object of the exercise is to demonstrate the locali­ zation of some particular chemical entity, which takes precedence over the preservation of ultrastructure. Often the degree of resolution of the techniques is another limiting factor, so that high magnification electron micrographs are not required. This is particularly true of cytochemical techniques for enzymes where some diffusion of the reaction product is inevitable. To take an extreme example, in the study of cholinergic neurones it is often only necessary to localize the staining to the rough endoplasmic reticulum (Fig. 3.15) and this requires a final magnification

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Fig. 3.15 A motor neurone in the hypoglossal nucleus of a rat is easily identified by the intense staining of the Nissl bodies in a tissue slice stained for acetylcholinesterase by the thiocholine method. Scale bar = 2 μπι.

of only a few thousand times. This whole group of techniques is conveniently divided into three categories, which are discussed below in order of increasing specialization of the fixation conditions required. 3.12.1

Fixation for general cytochemical techniques

Only some general remarks on the necessity for special fixation methods and a simplified summary of individual techniques (Table 3.2) are included here. For more detailed information the reader is referred to a companion book in this series by Lewis and Knight (1992). For the majority of cytochemical techniques an aldehyde fixative is used at the ambient temperature and the precise composition of the fixative is chosen to give the best preservation of ultrastructure. That is what is meant by the term 'mixed aldehydes' in Table 3.2. The only general exception to the use of a mixed aldehyde fixative is where the cytochemical technique depends upon the detection of aldehyde groups;

Chapter 3:

Fixationmethods

Table 3.2 Fixation for general cytochemical techniques Technique

Fixative

General methods for nucleic acids

Mixed aldehydes

Specific methods for DNA

4% Formaldehyde

Selective staining for RNA

Mixed aldehydes

Specific staining for sulphydryl groups

Mixed aldehydes

Periodic acid methods for carbohydrates

4% Formaldehyde

Methods for acidic carbohydrates

Mixed aldehydes

Staining with lectins

Mixed aldehydes

Methods for lipids

Mixed aldehydes

typical examples being some methods for DNA and methods involving the use of periodic acid. Here acrolein definitely cannot be used and glutaraldehyde ought not to be used, because the process of fixation can lead to the formation of gratuitous free aldehyde groups which would give a false positive reaction. Formaldehyde is the fixative of choice since it cannot form spurious aldehyde groups. In a few specialized techniques a cytochemical reagent is included in the fixative. In other words, the fixation stage is part of the cytochemical staining procedure. Examples of these fixatives are given in Sect. 2.5.4. and Sect. 2.8.3 With cytochemical techniques the only staining present in the final ultrathin section should be that due to the specific chemical constituent it is wished to demonstrate. Osmium tetroxide has, therefore, to be omitted from most of the schedules, because it introduces chemical groups which give spurious staining. Post-fixation with uranyl acetate is also omitted, because it can obscure the cytochemical staining. The ultrathin sections should be examined 'unstained', although adjacent sections can be stained with lead citrate and/or uranyl acetate to reveal finer details of ultrastructure, if required. In most general cytochemical techniques ultrathin sections are cut and then stained in much the same way that paraffin sections are stained for light microscopy. In a few techniques, however, it is necessary to cut thin slices of the aldehyde-fixed material, say on a Vibratome (Sect. 3.5.1a), and stain or otherwise treat these slices before proceeding with

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Fig. 3.16 The use of colloidal thorium dioxide to stain proteoglycans in cartilage matrix. Note that the cells are not penetrated by the colloidal particles. Scale ΐ33Γ = 2μιτι. (Reproduced from Groot 1981, with permission.)

the rest of the schedule. One very important need for this type of schedule is where treatment with enzymes is necessary, either as a control for the ultimate staining or as a means of distinguishing between different tissue components which would all produce the same type of final staining. There are also some very useful cytochemical stains which do not penetrate ultrathin resin sections. They then have to be applied to thin slices after aldehyde fixation as in Fig. 3.16, which shows the staining of proteoglycans in cartilage by colloidal thorium dioxide applied overnight to aldehyde-fixed slices.

Chapter 3:

Fixationmethods

Iilllllfte Fig. 3.17 Intense staining of the outer cell membrane of a horse chondrocyte stained for alkaline phosphatase. Scale bar = 2 μηι. (Unpublished micrograph from a study by Dr Jeremy Skepper, reproduced with permission.)

3.12.2

Fixation for enzyme cytochemical techniques

Rather more than a hundred enzymes can be demonstrated by histochemical techniques at the tight microscope level and somewhat fewer at the electron microscope level. Not many of them will survive any standard form of embedding procedure, however, so that the staining technique must be applied to thin slices immediately after appropriate aldehyde fixation. A few enzymes, such as alkaline phosphatase (Fig. 3.17), will survive fixation at room temperature, but fixation for enzymes is routinely carried out at 4°C, usually for 1 to 2 hours. The activity of most enzymes is reduced less by formaldehyde than by glutaraldehyde, but a small amount of glutaraldehyde is often added to the fixative to improve the preservation of ultrastructure. A typical fixation schedule is therefore 4% formaldehyde, prepared as usual from paraformaldehyde, plus 1% glutaraldehyde in 0.1 M cacodylate or PIPES buffer, at pH 7.0 to 7.4, for

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Table 3.3 Fixation for enzyme cytochemical techniques Enzyme

Fixative

Acid phosphatase

Mixed aldehydes

5'-Nucleotidase

Mixed aldehydes

Thiamine pyrophosphatase

Mixed aldehydes

Sulphatases

Mixed aldehydes

ATPases

Dilute aldehyde

Alkaline phosphatase

Mixed aldehydes

Glucose-6-phosphatase

Dilute aldehyde

Non-specific esterases

Mixed aldehydes

Cholinesterases

Mixed aldehydes

Peroxidase

Mixed aldehydes

Cytochrome oxidase

4% Formaldehyde

Catalase

Mixed aldehydes

DOPA-oxidase

Mixed aldehydes

Succinic dehydrogenase

None

Precautions

Short fixation

Short fixation Short fixation

Short fixation

α-Hydroxy acid oxidase

2% Formaldehyde

Short fixation

Diaphorase

Mixed aldehydes

Short fixation

Lactic dehydrogenase

Mixed aldehydes

1 to 2 hours at 4°C. This is denoted by the unqualified term 'mixed aldehydes' in Table 3.3. The same apparent enzyme can differ in its properties from species to species and even from tissue to tissue, so that it may be necessary to vary the conditions of the initial aldehyde fixation to suit the particular study being undertaken. If fixation is by perfusion, the perfusion fluid is used at room temperature, but the time is limited to about 10 min and tissues are removed and transferred to fixative at 4°C as soon as possible. If a difficult dissection is involved, such as the brain, the relevant part of the animal, in this case the head and neck, is placed without fixative in the refrigerator in a plastic bag for about an hour after the perfusion to cool down before dissection. Some enzymes will tolerate only formaldehyde and some only short fixation, as noted in Table 3.3. Lewis and Knight (1992) should be consulted for further details.

Chapter 3:

Fixation methods

In a typical schedule following the initial aldehyde fixation, the sample is washed in buffer and cut on a Vibratome or similar instrument (Sect 3.5.1a) into slices 50 to 250 μηι in thickness. The thinnest slices are necessary when the incubation stage of the enzyme technique lasts for 30 min or less. For longer incubation times the thicker slices can be used. Typically the incubation stage is carried out at 0 to 4°C and it is some­ times preceded by a short pre-incubation period, also at 0 to 4°C. A post-incubation treatment is sometimes necessary to stabilize the initial reaction product of the enzyme activity. At this stage the slices are washed again in buffer and post-fixed. Unbuffered osmium tetroxide is routinely used, often without the addition of ferro- or ferri-cyanide. Sometimes, however, osmium tetroxide obscures the enzyme staining and then 4% glutaraldehyde at room temperature for 1 to 2 hours is used instead to improve ultrastructural preservation. Treatment with uranyl acetate is seldom used. 3.12.3

Fixation for immunocytochemical techniques

The reactivity and stability of specific antigens differ so widely from one antigen to another that no one method of fixation can be advocated for their demonstration in the electron microscope by immunological techniques. Few antigens withstand the routine fixation and embedding procedures used for ultrastructural studies and it is usually necessary to Table 3.4 Fixation for immunocytochemical techniques Fixative 1% Glutaraldehyde 4% Formaldehyde plus 0.1% glutaraldehyde

Duration 30-60 mm 60 min

8% Formaldehyde

30—60 min

4% Formaldehyde

60 min

4% Carbodiimide

60 min

1 % Formaldehyde

60-120 min

The fixatives are listed in the order of their decreasing ability to preserve ultrastructure. All the fixatives contain 0.1 M cacodylate or PIPES buffer, at pH 7.0 to 7.4, with 2 to 3 mM calcium chloride, and are used at 0 to 4°C.

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use a much more gentle fixation. The best way of choosing the optimum fixation conditions for electron microscopy is to cut a series of cryostat sections at one particular thickness from an unfixed block of tissue and subject these to a graded set of fixation schedules to assess the strength of the antigenic response obtainable at the light microscope level. Fixation should be at 4°C in an isotonic fixative buffered to a pH of 7.0 to 7.4 with 0.1 M cacodylate or PIPES buffer containing 2 to 3 mM calcium chloride (see Sect. 2.3). Table 3.4 lists such a graded series of fixation schedules which are arranged in order of decreased ability to preserve ultrastructure. For electron microscopy choose the fixative highest on the list which gave an adequate response at the light microscope level. Do not use osmium tetroxide or uranyl acetate as additional fixatives. Low temperature embedding techniques, as discussed in Chapter 8, may have to be used to retain an adequate level of antigenic reactivity. Inevitably there has to be a certain amount of trial and error before the optimum fixation and embedding schedule can be arrived at.

References Abrunhosa, R. (1972) Microperfusion fixation of embryos for ultrastructural studies. Journal of Ultrastructure Research 41,176-188 Bhatnagar, M.K., David, L.L., Vrablic, O., Therien, A. and Blouin, A. (1981) A simple method for perfusion fixation of avian liver for electron microscopy. CanadianJournal of Zoology 59,1179-1183 Bozzola, J.J. and Russell, L.D. (1992) Electron Microscopy. Principles and Techniques for Biologists. Jones and Bartlett, Boston Coalson, J.J. (1983) A simple method of lung perfusion fixation. Anatomical Record 205, 233-238 Crang, R.F.E. (1997) Protocol of plant specimen preparation for transmission electron microscopy. Microscopy Today 97-7,18-20 Dae, M.W., Heymann, M.A. and Jones, A.L. (1982) A new technique for perfusion fixation and contrast enhancement of fetal lamb myocardium for electron microscopy. Journal of Microscopy 127, 301-305 Eccles, M.H. and Glauert, A.M. (1984) The response of human monocytes to interaction with immobilized immune complexes. Journal of Cell Science 71,141-157 Elgjo, R.F. (1976) Platelets, endothelial cells and macrophages in the spleen. An ultrastructural study on perfusion-fixed organs. American Journal of Anatomy 145, 101-120

Elling, F., Hasselager, E. and Friis, C. (1977) Perfusion fixation of kidneys in adult pigs for electron microscopy. Acta Anatomica 98, 340-342

Chapter 3:

Fixationmethods

Forssmann, W.G., Sicgrist, G., Orci, L., Girardier, L., Pictet, R. and Rouiller, C. (1967) Fixation par perfusion pour Ie microscopie electronique essai de generalisation. Journal de Microscopie 6, 279-304 Furness, J.B., Heath, J.W. and Costa, M. (1978) Aqueous aldehyde (Faglu) methods for the fluorescence histochemical localization of catecholamines and for ultrastrucutural studies of central nervous tissue. Histochemistry 57, 285-295 Gil, J. and Weibel, E.R. (1969) Improvements in demonstration of lining layer of lung alveoli by electron microscopy. Respiratory Physiology 8, 13-36 Glauert, A.M. (1975) Fixation, dehydration and embedding of biological specimens. In Practical Methods in Electron Microscopy, Vol. 3, Part I, Glauert, A.M. (ed.), NorthHolland, Amsterdam Griffiths, G. (1993) Fine Structure Immunocytochemistry. Springer-Verlag, Berlin and Heidelberg Groot, C.G. (1981) Positive colloidal thorium dioxide as an electron microscopical contrasting agent for glycosaminoglycans, compared with ruthenium red and positive colloidal iron. Histochemistry 71, 617-627 Gunning, B.E.S. and Steer, M.W. (1996) Plant Cell Biology. Structure and Function. Jones and Bartlett, Boston Haudenschild, C., Baumgartner, H.R. and Studer, A. (1972) Significance of fixation procedure for preservation of arteries. Experientia 28, 828-831 Hayat, M.A. (1989) Principles and Techniques of Electron Microscopy; Biological Applications, 3rd edn. Macmillan Press Ltd., Basingstoke Hinton, D.E. (1975) Perfusion fixation of whole fish for electron microscopy. Journal of the Fisheries Research Board of Canada 32, 416-422 Hiraoka, J.-I. and Wang, W. (1989) A new infusion needle for perfusion fixation. Stain Technology 64, 102-103 Kok, L.P.and Boon, M.E. (1990) Microwaves for microscopy. Journal of Microscopy 158, 291-322 Kok, L.P. and Boon, M.E. (1992) Microwave Cookbook for Microscopists, 3rd edn. Coulomb Press Leyden, Leiden Lewis, P.R. and Knight, D.P. (1992) Cytochemical staining methods for electron microscopy. In Practical Methods in Electron Microscopy, Vol. 14, Glauert, A.M. (ed.), Elsevier, Amsterdam Login, G.R. and Dvorak, A.M. (1994a) Methods of microwave fixation for microscopy. Progress in Histochemistry and Cytochemistry 27, 1-127 Login, G.R. and Dvorak, A.M. (1994b) The Microwave Tool Book. Beth Israel Hospital, Boston Maclntyre, D.E., Allen, A.P., Thorne, K.J.I., Glauert, A.M. and Gordon, J.L. (1977) Endotoxin-induced platelet aggregation and secretion. Journal of Cell Science 28, 211-223 McCombs, R.M., Benyesh-Melnick, M. and Brunschwig, J.P. (1968) The use of Millipore filters in ultrastructural studies of cell cultures and viruses. Journal of Cell Biology 36, 231-243 Paavola, L.G. (1977) The corpus luteum of the guinea pig. Fine structure at the time of maximum progesterone secretion and during regression. AmericanJournal of Anatomy 150,565-604 Page, A.M., Lagnado, J.R., Ford, T.W. and Place, G. (1994) Calcium alginate encapsulation of small specimens for transmission electron microscopy. Journal of Microscopy 175,166-170

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Rossi, G.L. (1975) Simple apparatus for perfusion fixation for electron microscopy. Experientia 31, 998-1000 Sparkman, D.R., Hammon, K.M. and White, C.L. (1987) A small adaptor for the ultracentrifugation of small sample volumes of subcellular particles from Alzheimer disease brain. Journal of Electron MicroscopyTechnique 5, 115-116 Strausbauch, P., Roberson, L. and Sehgal, N. (1985) Embedding of cell suspensions in ultra-low gelling temperature agarose: improved specimen preparation for TEM. Journal of Electron Microscopy Technique 2,261-262 Swinehart, P.A., Bentley, D.L. and Kardong, K.V. (1976) Scanning electron microscopic study of the effects of pressure on the luminal surface of the rabbit aorta. American Journal of Anatomy 145, 137-142 Weibel, E.R. (1970) Morphometric estimation of pulmonary diffusion capacity. I. Model and method. Respiratory Physiology 11, 54-75 Wild, P. and Setoguti, T. (1995) Mammalian parathyroids: morphological and functional implications. Microscopy Research and Technique 32, 120-128 Wisse, E., De Wilde, A. and De Zanger, R. (1984) Perfusion fixation of human and rat liver for light and electron microscopy: a review and assessment of existing methods with special emphasis on sinusoidal cells and microcirculation. In The Science of Biological Specimen Preparation, Revel, J.-P., Barnard, T. and Haggis, G.H. (eds.), pp. 31-38, SEM Inc., AMF O'Hare, Chicago Wood, J.I. and Klomparens, K.L. (1993) Characterization of agarose as an encapsulation medium for particulate specimens for transmission electron microscopy. Microscopy Research and Technique 25, 267-275 Yun, J. and Kenney, R.A. (1976) Preparation of cat kidney tissue for ultrastructural studies. Journal of Electron Microscopy 25, 11-23

4 Dehydration methods

Since most embedding media are not miscible with water, it is necessary to 'dehydrate' fixed specimens by taking them through a sequence of solutions leading up to one which is fully miscible with the embedding medium. Ethanol and acetone are the two dehydrating agents most commonly used in ultrastructural studies, and are usually followed by an intermediate solvent, such as propylene oxide, before embedding in an epoxy resin. In the standard method well-fixed specimens are dehy­ drated at room temperature. Alternatively, excellent preservation of ultrastructure is obtained after dehydration of cryofixed specimens by freeze-substitution and embedding in an epoxy resin. The polar acrylic resins and some epoxy resins are miscible with water and can act as dehydrating agents or can be used in partial dehydration schedules in which dehydration is completed in mixtures of the dehydrating agent, the resin and water. When the preservation of antigens is paramount, specimens are dehydrated at low temperatures by the progressive lowering of tem­ perature method or by freeze-substitution. Similarly, dehydration by freeze-drying is recommended for X-ray microanalysis, to minimize the redistribution of soluble ions.

4. I

Chemical and morphological effects of dehydration

The major chemical effect of dehydration is the extraction of lipids from the specimen by dehydrating agents and intermediate solvents, resulting in a shrinkage of cells and their components. The physical state of proteins, mucopolysaccharides and nucleic acids is also affected by dehydration, but the extraction losses are lower, occurring mainly in dilute solvent solutions.

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Dimensional changes during dehydration

Overall, the specimen shrinks as a consequence of osmotic effects and as water and lipids are extracted. The effect of ethanol dehydration on the volume of specimens has been extensively studied by Boyde and his collaborators (Boyde et al. 1977; Boyde and Boyde 1980; Boyde and Maconnachie 1981) for scanning electron microscopy, and many of their results are also applicable to transmission electron microscopy. There are considerable variations in the response of different types of specimen to dehydration (Lee 1984), but the general pattern of dimensional changes is similar throughout and is illustrated in Fig. 4.1, which shows the effects of taking a glutaraldehyde-fixed mouse embryo limb through successive ethanolic solutions (Boyde 1980). There are two stages in the process. A significant swelling of 15 to 20% by volume in 30% and 50% ethanol is followed by shrinkage. The initial swelling is

80% Ethanol

Time (minutes) Fig. 4.1 Dimensional changes during ethanol dehydration of a 15-day mouse embryo limb, which had been fixed in 3% glutaraldehvde in cacodvlate buffer. The specimen volume in buffer at the start of the experiment is taken as 100%. Swelling occurs in 30% and 50% ethanol, and shrinkage in the stages up to 100% ethanol. The most rapid shrinkage occurs in 80% ethanol. (Data taken from Bovde 1980.)

Chapter 4:

Dehydration methods

nearly reversed in 70% ethanol and then there is massive shrinkage in 80% ethanol, the overall reduction in volume reaching 30% in absolute ethanol. Dehydration with more gradual steps in ethanol concentration has little effect on these volume changes; the most rapid shrinkage still occurs in 80% ethanol and the overall shrinkage is only reduced to 25% by volume. If the specimen is passed directly from water to 70% ethanol, there is only a transient period of swelling, but the subsequent shrinkage is not affected. The extent of swelling in 30% and 50% ethanol, which may well cause irreversible structural damage to the specimen, depends on the fixation procedure and is greater after fixation with osmium tetroxide alone than with glutaraldehyde or with glutaraldehyde followed by osmium. It can be prevented completely by the addition of divalent cations (for example 1 mM to 0.1 M Ca ++ ) at any stage before or during dehydration. However, this inhibition of swelling is accompanied by an increased overall shrinkage of specimens fixed only with glutaraldehyde (that is without post-fixation with osmium tetroxide), giving them a shrivelled-up appearance. Dilute ethanolic solutions extract proteins and other tissue constituents, as well as inducing swelling, and so there are strong arguments for proceeding directly to 70% ethanol in the dehydration sequence (Lee 1984). The overall shrinkage of specimens during dehydration can be as high as 30 to 40% by volume for some specimens, such as isolated cells, and the possibility of differential shrinkage must always be borne in mind. For some specimens shrinkage can be reduced by continuous dehydration (Sect. 4.4.2), but the most important factor is the fixation procedure. For example, Boyde et al. (1977) found that the volume shrinkage of mouse embryo limbs could be reduced to 4% by tertiary fixation with uranyl acetate. 4.1.2

The retention o f lipids

It cannot be emphasized enough that for good preservation of ultrastructure it is necessary to reduce to a minimum the degree of extraction of membrane lipids during dehydration, since membranes are such an important feature of the final image seen in the electron microscope. As much as 95% of the total lipids may be lost during the dehydration of an aldehyde-fixed specimen, while secondary fixation with osmium tetroxide reduces this loss considerably (Korn and

13 I

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Weisman 1966) and further fixation with uranyl acetate still more (Weibull et al. 1983). Aqueous solutions of ethanol of 70% or less dissolve relatively little lipid from specimens fixed with glutaraldehyde and osmium tetroxide, the majority of the lipid loss occurring in the higher ethanol concentrations. Thus the partial dehydration schedule recommended by Idelman (1964), in which the specimen is passed from 70% ethanol to a mixture of 70% ethanol and an Epon embedding medium (Sect. 4.4.4) reduces the loss of lipids. Stein and Stein (1967) used a modified version of this schedule and found that lipid extraction can be reduced even further by using shorter times for the steps in the dehydration procedure and by reducing the temperature to 0°C. These methods are not effective for specimens fixed only with glutaraldehyde, where temperatures of -50°C or lower are required to reduce lipid extraction, as described in the discussion of low temperature embedding in Sect. 8.1. Acetone is usually a more gentle lipid solvent than ethanol, while propylene oxide has been described as a very 'energetic' solvent for lipids by Idelman (1964). Acetonitrile has a low solubility for phos­ pholipids in unfixed liver and is a promising replacement for ethanol and propylene oxide (Edwards et al. 1992). Acrolein is a lipid fixative, but glutaraldehyde and formaldehyde are not. If they are used, some form of post-fixation is essential. Osmium tetroxide is the most widely used secondary fixative and is the best lipid fixative available. Both potassium ferri- and ferro-cyanide enhance the lipid staining produced by osmium tetroxide, and presumably cause more lipid to be retained. For ultrastructural studies many electron microscopists now use an unbuffered solution containing 1.0% osmium tetroxide and 1.5% potassium ferricyanide as their standard secondary fixative. The increased membrane staining produced is illustrated in Fig. 4.2. Further retention of lipids is obtained by the use of aqueous uranyl acetate as a tertiary fixative. Locke (1994) advocates the use of uranyl acetate instead of osmium tetroxide, particularly for studies in which osmium tetroxide is best avoided, as in some immunocytochemical studies. In general, however, the optimum preservation of lipids is obtained with a primary aldehyde fixative containing calcium, an osmium tetroxide secondary fixative and uranyl acetate as a tertiary fixative (Sect. 3.4.1). The use of these three fixatives in sequence gives the best preservation of lipid membranes and consequently the best ultrastructural results.

Chapter 4:

Dehydration methods

w m m m .

Fig. 4.2 A comparison showing the improvement in membrane fixation and contrast when potassium ferricyanide is added to the osmium tetroxide solution used for postfixation. Insect Malphigian tubules fixed without (a,c) or with (b,d) potassium ferricyanide. The apical part of a typical cell is shown in (a) and (b), and the basal part in (c) and (d). Note the much more distinct membranes of the brush border and of the underlying vacuoles in (b), compared to (a), and the more intense staining of the basement membrane in (d), compared to (c). There is also better preservation of the mitochondrial cristae in the presence of potassium ferricyanide. bm, basement membrane; m, mitochondria. (Reproduced from Felgenhauer et al. 1996, with permission.)

In conclusion, the following methods can be expected to reduce the loss of lipids during dehydration: i. ii. iii. iv.

Add calcium to the primary fixative and washing solutions. Fix the specimens with osmium tetroxide before dehydration. Include potassium ferricyanide with the osmium tetroxide. Fix the specimens with uranyl acetate before dehydration.

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v. Select a dehydrating agent with minimal solubility for lipids. vi. Dehydrate the specimens as rapidly as possible. vii. Use a low temperature for the dehydration sequence. viii. Use a partial dehydration schedule. ix. Avoid the use of propylene oxide as an intermediate solvent.

4.2

Safety procedures

Dehydrating agents and intermediate solvents must be handled with the safety precautions described in Sect. 1.4.5. The labels on bottles obtained from suppliers should be examined and the solvents must be stored under the correct conditions. An efficient fume cupboard or a ventilated bench should always be used, together with suitable protective clothing. Since all these solvents rapidly extract lipids from skin, it is particularly important to wear solvent-resistant gloves (see Appendix) at all times. "Waste solvents should be poured into a suitable container and stored in a cupboard designated for flammable liquids to await disposal. Propylene oxide is highly flammable, very volatile and carcinogenic. It must be handled with particular care. Electrical equipment and other potential sources of ignition should be kept well away from the working area. Beginners should always read the description of safety procedures in Sect. 1.4 before commencing a dehydration procedure.

4.3

Dehydrating agents and intermediate solvents

Many different dehydrating agents have been tested for electron microscopy, but few of these are widely used. Those recommended here are all available from general suppliers (see Appendix). 4.3.1

The standard dehydrating agents and intermediate solvents

The usual dehydrating agents are ethanol and acetone and they give similar results with the majority of specimens. Acetone has a lower viscosity than ethanol and the dehydration process is more rapid. It is also a less aggressive solvent for most lipids, but it is very hygroscopic and incomplete dehydration may result unless the acetone is carefully

Chapter 4:

Dehydrationmethods

dried and kept dry in a sealed bottle. In addition, acetone is a radical scavenger and can inhibit the polymerization of acrylic resins. There is still some difference of opinion about the relative merits of ethanol and acetone as the main dehydrating agent, but in general ethanol is preferred (Lee 1984). It is easier to handle, since it is not very volatile, is relatively non-toxic and is readily miscible with water. Although most epoxy resins are miscible with ethanol and acetone, they mix much more readily with some other organic solvents, such as propylene oxide, and these intermediate solvents are routinely used following dehydration of well-fixed specimens. Xylene and toluene are also suitable as intermediate solvents for epoxy resins, but propylene oxide is preferred since it is a low viscosity epoxy diluent which reacts with the anhydride hardener in the embedding medium and becomes an integral part of the cross-linked polymer. Consequently no problems arise if a small amount of propylene oxide remains in the tissue. Propylene oxide is miscible in all proportions with ethanol, acetone and epoxy resins. It quickly replaces the dehydrating agent and then diffuses out of the tissue allowing easy infiltration of the epoxy resin. It must be handled with full safety precautions in an efficient fume cupboard (Sect. 1.4.5). 4.3.2

Alternative dehydrating agents and intermediate solvents

Various modifications to the standard dehydration procedure with ethanol or acetone and propylene oxide have been suggested. Thus acidified 2,2dimethoxypropane (DMP) was advocated by Muller and Jacks (1975) for faster dehydration, since it reacts rapidly with water to release methanol and acetone, and should cause much quicker removal of water from the specimen. The overall shrinkage, however, is frequently more rapid and serious with DMP than with other dehydrating agents (Boyde et al. 1977) and there is a greater extraction of lipids than during dehydration in acetone (Beckmann and Dierichs 1982). DMP has an unpleasant odour and appears to offer no significant advantages. It should be avoided, except for specialist applications, such as X-ray microanalysis. Edwards et al. (1992) proposed acetonitrile as a less toxic alternative in place of the ethanol/propylene oxide combination. Although it is less toxic in itself, acetonitrile is a cyanide derivative, specifically, methyl cyanide (CH3CN). Consequently, it must not be exposed to excessive

135

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heat, to strong acids, or to strong reducing agents. Strong oxidizing agents, with which it can react violently, must also be avoided. With these precautions, which are easily taken, it is as safe to use as other dehydrating agents and safer than propylene oxide. Acetonitrile extracts significantly less phospholipid from unfixed liver and has a lower viscosity than ethanol. It is easier to use than propylene oxide, since it is less volatile with a much higher boiling point (over 80°C compared with under 40°C). It is freely miscible with water, alcohols and epoxy resins, and can replace ethanol as the dehydrating agent, as well as propylene oxide as the intermediate solvent. Again, the complete replacement of acetonitrile with the epoxy resin is not essential. Thus acetonitrile appears to have considerable potential as a combined dehydrating agent and intermediate solvent, although it is more expensive than ethanol and propylene oxide. Other organic solvents proposed for dehydration include methanol, which is a useful alternative to acetone for dehydration by freezesubstitution, but it is not suitable for routine dehydration, since it extracts lipids (Weibull and Christiansson 1986) and the cytoplasmic matrix more extensively than ethanol, when it is used with acetone or propylene oxide as the intermediate solvent (Lee 1984). Polyvinyl alcohol and polyethylene glycol have both been advocated for dehydration, because there is some evidence that they extract less lipid, but they are viscous and penetrate tissues slowly. They cannot be recommended.

4.4

Dehydration schedules

In the dehydration procedure the fixed specimen is passed through a graded series of solutions of increasing concentration of the dehydrating agent in water, ending with pure (absolute) dehydrating agent. Stock solutions of 100% dehydrating agents should be kept dry in tightly sealed bottles and these should be opened for as short a time as possible, particularly in humid conditions. It is often convenient to keep each specimen in the vial (Sect. 3.4) or tube in which it was fixed throughout the dehydration procedure. If plastic vessels are used they must be resistant to solvents, such as propylene oxide. Suitable polyethylene and polypropylene containers are available from general

Chapter 4:

Dehydrationmethods

suppliers (see Appendix). During dehydration one solution is carefully removed with a fine Pasteur pipette and the next solution added. This process is carried out as rapidly as possible to prevent the sample drying out and the vials are then capped firmly to prevent absorption of water from the air. It is essential to ensure that there is a large excess of each solution, at least 20 times the volume of the specimen. Solutions containing larger specimens should be agitated by placing the vials on a specimen rotator (Sect. 3.4). As with fixation, better dehydration is obtained with small and thin specimens. Slices of tissue, not more than 250 μηι thick, are preferable to small cubes. 4.4.1

A standard dehydration schedule

Fixed specimens are washed briefly in distilled water before dehydration to remove excess fixative and buffer components (Sect. 3.4.1). In general, concentrations of the dehydrating agent of less than 70% should be avoided to prevent the morphological changes described in Sect. 4.1.1. Dehydration is normally carried out at room temperature and a typical schedule for dehydration of thin slices of tissue is given in Table 4.1. The steps of the dehydration schedule are not critical, and the times given here are probably the maximum required, even for compact, wellfixed tissue, provided that it is no more than 1 to 2 mm thick. Excessive dehydration should be avoided since it can result in greater extraction and degradation of ultrastructure. When specimens are to be embedded in an epoxy resin, the dehydrating agent is now replaced with the intermediate solvent, propylene oxide, within a fume cupboard. Propylene oxide and concentrated dehydrating agents must not be flushed down a sink after use, but should be placed in a waste bottle for disposal (Sect. 1.4.5c).

Table 4.1 The standard schedule for dehydration at room temperature.

70% Ethanol or acetone in water

10-20 min

90 or 95% Ethanol or acetone in water

10-20 min

100% Ethanol or acetone

15-30 min

100% Ethanol or acetone

15-30min

Ethanohpropyleneoxide Propylene oxide

1:1

10 min 10 min

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When specimens have to be stored during processing, this should be done in the buffer wash following the primary fixation (Sect. 3.3.2), whenever possible. Specimens which have to be stored overnight during dehydration in an emergency should be kept in 70% ethanol or acetone in the cold to minimize the loss of lipids. 4.4.2

Alternative dehydration schedules

Modifications to the standard schedule are required depending on the type of specimen. For example, longer dehydration times are necessary for cells with dense walls, such as plants and fungi (see Sect. 3.7.1). More delicate specimens, such as suspensions of cells and tissue cultures, may need to be dehydrated through a more closely spaced series of concen­ trations of the dehydrating agent than in the standard schedule, but with shorter times for each step. Shrinkage occurs during dehydration with all solvents (Sect. 4.1.1) as a consequence of extraction and osmotic effects and is particularly evident following light fixation (Lee 1984). This shrinkage can be reduced significantly with some, but not all, specimens by gradually and continuously changing the concentration of the dehydrating agent to avoid steep osmotic gradients. A simple method is to place the specimen in a vial with a small quantity of water and then to add 3 to 4 volumes of ethanol or acetone, drop by drop, with continuous shaking. This is followed by two short changes of 100% dehydrating agent. More sophisticated methods of continuous dehydration have been developed, such as the exchange apparatus of Peters (1980), and the continuous flow system of Rostgaard and Tranum-Jensen (1980). These Table 4.2 A schedule for dehydration in acetonitrile at room temperature (Edwards et al. 1992)

50% Acetonitrile in water

15 min

70% Acetonitrile in water

20 min

90% Acetonitrile in water

2X10 min

100% Acetonitrile Acetonitrile:epoxy resin Epoxy resin

3 X 20 min 1:1

1 K and 18 h 3X2h

Chapter 4:

Dehydration methods

continuous dehydration methods are only required for specimens that are particularly sensitive to osmotic effects. For the majority of speci­ mens they do not appear to have any advantages over more simple methods and they are rarely used. A modified procedure is required when acetonitrile (AN) replaces ethanol and propylene oxide (Sect. 4.3.2). The schedule proposed by Edwards et al. (1992) for the dehydration of small cubes (0.5 mm sides) of kidney and liver tissue, followed by infiltration with an epoxy resin (the authors used EMbed 812), at room temperature, is given in Table 4.2. Acetonitrile can also be used just as an intermediate solvent following dehydration with a different reagent. For example, Harb (1993) dehydrated human tissues in methanol and then replaced the methanol with AN at room temperature (AN, 3X10 min; AN:epoxy resin, 1:1, 60 min; epoxy resin, 2 X 60 min). A rapid schedule, in which the times were halved, was also used. 4.4.3

Rapid dehydration

When required the dehydration can be made considerably more rapid. Specimens must be 0.1 mm or less in at least one dimension and the specimens should be agitated throughout dehydration. The rapid dehydration schedule of Bencosme and Tsutsumi (1970), given in Table 4.3, is then possible. Dehydration times can also be decreased by continuous dehydration, as described in Sect. 4.4.2. The use of acidified 2,2-dimethoxypropane (DMP) for rapid dehydration is not recommended (Sect. 4.3.2), except under special circumstances, such as pathological diagnosis, when speed is paramount and ultrastructure is not. Table 4.3 A schedule for rapid dehydration at room temperature (Bencosme and Tsutsumi 1970)

70% Ethanol in water

3 min

80% Ethanol in water

3 min

90 % Ethanol in water

3 min

100% Ethanol

2X5min

Propylene oxide

2X5min

139

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4.4.4

Biological specimen preparation for TEM

Partial dehydration

Some epoxy and acrylic resins are miscible with aqueous solutions of ethanol and enable the higher concentrations of ethanol and the propylene oxide to be omitted from the dehydration schedule, thus avoiding the steps during which the majority of lipid extraction occurs. Epon 812 is miscible with 70% ethanol in water and in the schedule proposed by Idelman (1964) the specimen is passed from 70% ethanol to a 1:1 mixture of 70% ethanol and Epon embedding medium. It is infiltrated with this mixture for 30 min at room temperature and then for 30 min at 37°C before infiltration with the embedding medium in the usual way. The potential value of this procedure is illustrated in Fig. 4.3. A modified version of this schedule has been used extensively by Stein and Stein (1971) for a great variety of lipids. They found that the loss of some lipids is reduced if the specimen is first infiltrated with the Epon resin alone, since the acid anhydride hardeners and the tertiary amine accelerator appear to increase the solvent action of the complete embedding medium. Lipid extraction is reduced even further by using shorter steps in the dehydration procedure and by decreasing the temperature. Their schedule (Stein and Stein 1967) for partial dehydration at O 0 C before embedding in Epon is given in Table 4.4. The specimens are then infiltrated with complete Epon embedding medium, overnight at 4°C, before infiltration and embedding in the usual way. A similar schedule can probably be used with the various Epon 812 replacements (see Sect. 6.2.5). Similar partial dehydration schedules have been proposed for polar acrylic resins, although there is no experimental evidence that these pro­ cedures reduce the loss of lipids (Acetarin et al. 1986). In fact, methacrylate resins have been found to extract more lipids than ethanol

Table 4.4 A schedule for partial dehydration at O 0 C (Stein and Stein 1967)

70% Ethanol in water

2X5 min

95% Ethanol in water

2X5 min

Epon 812

3X1h

Chapter 4:

Dehydration methods

Fig. 4.3 The improved lipid retention obtained by omitting a non-polar solvent during infiltration with the embedding medium. Both sections are from rat adrenal cortex fixed in osmium tetroxide. In (a) the lipid droplets (L) are extracted when propylene oxide was included in the schedule. In (b) they appear dense when the tissue was transferred from 70% ethanol directly to a 1:1 (v/v) mixture of 70% ethanol and Epon. Mitochondria (M); lysosome (Li). Scale bars = 1 μηι. (Reproduced from Fruhling et al. 1969, with permission.)

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and Epon (Idelman 1965), unless the dehydration is at a very low temperature. Schedules for partial dehydration before embedding in London or Lowicryl acrylic resins are given in Sect. 7.3.3b and Sect. 8.7.1, respectively. The epoxy resins, Durcupan and Aquembed, and the acrylic resins, glycol methacrylate and hydroxypropyl methacrylate, are fully miscible with water and the schedules for their use as dehydrating agents are given in Sect. 6.4.2 and Sect. 7.2.4

4.5

Dehydration by freeze-substitution

During dehydration by the freeze-substitution method the specimen is rapidly frozen without chemical fixation (or after only light fixation) and then the ice in the specimen is replaced (substituted) by an organic solvent at temperatures of -80 to -90°C, which are low enough to prevent recrystallization of the frozen water. This procedure results in a significant decrease in the extraction of specimen components as com­ pared with standard dehydration methods (for example see Graham and Beveridge 1990a,b). Following freeze-substitution the specimen is infil­ trated with an epoxy resin for ultrastructural studies, or with an acrylic resin at low temperatures for immunocytochemical investigations. Full details of the principles and methods of freeze-substitution are given by Robards and Sleytr (1985) in a companion volume in this series and by Steinbrecht and Muller (1987), and these two publications are the basis for the outline of the technique given here. 4.5.1

Solvents for freeze-substitution

The most widely used solvents for freeze-substitution are methanol and acetone, while other fluids, such as diethyl ether and tetrahydrofuran, have been found to have some advantages for X-ray microanalysis, since they are less likely to cause translocation of ions, but the preservation of ultrastructure is then very poor. The choice of acetone or methanol depends on a number of factors. Acetone is a poor solvent for water at -85°C and so it is essential to pre-dry the acetone with molecular sieve (0.4 nm). Large volumes (10 to 15 ml) of fluid and long substitution times must be used. Substitution in methanol is much faster than with acetone and it substitutes ice in the presence of 10% water. Acetone is

Chapter 4:

Dehydrationmethods

favoured by some workers, however, just because it acts more slowly and so presumably is more gentle in its removal of water. It also extracts considerably less lipid than methanol during freeze-substitution (Weibull et al. 1984). As usual, the choice of fluid depends critically on the type of specimen. For example, Steinbrecht (1993) made a com­ parative study and concluded that methanol cannot be recommended for insect tissues. 4.5.2

Fixatives for freeze-substitution

The process of freeze-substitution in itself does not, of course, produce any chemical fixation of the specimen and introduces no contrast into the final electron micrograph. Fixatives, such as glutaraldehyde, osmium tetroxide (OSO4) and uranyl acetate, are therefore incorporated at appropriate stages into the freeze-substitution procedure. Little funda­ mental research has been done on the chemical reactivity of these standard fixing agents at low temperatures (see review by Steinbrecht and Muller 1987), but it is a matter of common experience that their addition to substitution fluids results in the excellent preservation of ultrastructure of specimens embedded in an epoxy resin. It has been shown that significant cross-linking of proteins by glutaraldehyde in acetone only occurs at temperatures above -45°C and so it is effective only in the later stages of freeze-substitution (Horowitz et al. 1990). There is some evidence that OsC>4 reacts with the double bonds of unsaturated fatty acids at temperatures as low as -70°C. The addition of uranyl acetate to methanol reduces the extraction of phospholipids at -70°C and of the glycolipids in lung alveolar type II cells at -90°C (Voorhout etal. 1991). Various combinations of the three basic fixatives have been suggested, depending on the substitution fluid used and the purposes for which the ultrathin sections are needed. Thus for immunocytochemical studies fixation is kept to a minimum, although structural preservation is then no more than adequate for the identification of organelles. For ultrastructural studies more than one fixative is often used. For example, Steinbrecht and Muller (1987) recommend a fixative consisting of 1% OSO4, 0.4% uranyl acetate, 3% glutaraldehyde and 3 % water in methanol, which is prepared in two parts. In one flask, precooled with liquid nitrogen, 6 ml of 50% aqueous glutaraldehyde are mixed with 40 ml of methanol, and then 2 ml

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of a 20% (w/v) solution of uranyl acetate in methanol are added. In a second precooled flask, 1.0 g of OsO 4 is dissolved in 50 ml of methanol. Both flasks are cooled to about -50°C, their contents are poured together, and they are shaken vigorously. The mixture is highly reactive and is stable for only a few hours, even at -30°C. The most popular fixatives for freeze-substitution in acetone are 1 to 5% OsO 4 , or a mixture of OsO 4 and uranyl acetate. McDonald and Morphew (1993) reported that 1% OsO 4 in acetone consistently gives the best results in comparison with a wide range of other fixatives. Solutions of OsO 4 in organic solvents are unstable at room temperature and should be freshly prepared and then cooled immediately before use. 4.5.3

The freeze-substitution procedure

The procedure of freeze-substitution is carried out at very low temperatures and so all the safety procedures outlined in Sect. 1.4.6 must be taken. Specimens to be dehydrated by freeze-substitution are cryofixed and stored under liquid nitrogen. A stock solution of the substitution fluid is then prepared and is kept in a closed container away from the laboratory atmosphere, since water is rapidly taken up by cold solvents. In a typical procedure, open containers, such as specimen vials, are half-filled with frozen substitution medium and placed in a Dewar flask containing liquid nitrogen. The volume of the fluid must be at least 1000 times greater than that of the specimen. The frozen specimens are transferred under liquid nitrogen to the surface of the frozen medium and screw caps, containing holes, are placed on the vials. The vials are then transferred to a substitution chamber and the temperature is raised to between -80°C and -90°C, so that the medium melts and the samples sink into it. The temperature of the substitution fluid is increased in stages until it reaches the temperature to be used for infiltration with the embedding medium, as outlined in the schedules in Sect. 6.2.9c and Sect. 8.7.2. The vials should be agitated gently throughout the procedure and care must be taken that the specimen does not dry out during changes of fluid. Alternatively, specimens and the substitution fluid are placed in closed containers which are suspended in the coolant in a deep Dewar vessel, as described by Steinbrecht (1993). Substitution at very low temperatures takes many hours with methanol and some days with acetone. Consequently

Chapter 4:

Dehydration methods

it is preferable to use one of the commercial units supplied by Leica or Balzers (Sect. 8.6.2) for freeze-substitution.

References Acetarin, J.-D., Carlemalm, E. and Villiger, W. (1986) Developments of new Lowicryl resins for embedding biological specimens at even lower temperatures. Journal of Microscopy 143, 81-88 Beckmann, H.-J. and Dierichs, R. (1982) Lipid extracting properties of 2,2dimethoxypropane as revealed by electron microscopy and thin layer chromatography. Histochemistry 76, 407—412 Bencosme, S.A. and Tsutsumi, V. (1970) A fast method for processing biologic material for electron microscopy. Laboratory Investigation 23, 447-450 Boyde, A. (1980) Review of basic preparation techniques for biological scanning electron microscopy. Proceedings of the VIIth European Regional Conference on Electron Microscopy, The Hague 2, 768-777 Boyde, A. and Boyde, S. (1980) Further studies of specimen volume changes during processing for SEM; including some plant tissue. Scanning Electron Microscopy /1980/ II, 117-132 Boyde, A. and Maconnachie, E. (1981) Morphological correlations with dimensional change during SEM specimen preparation. Scanning Electron Microscopy /1981/ IV, 27-34 Boyde, A., Bailey, E., Jones, S.J. and Tamarin, A. (1977) Dimensional changes during specimen preparation for scanning electron microscopy. Scanning Electron Microscopy /1977/ I, 507-518 Edwards, H.H., Yeh, Y.-Y., Tarnowski, B.I. and Schonbaum, G.B. (1992) Acetonitrile as a substitute for ethanol/propylene oxide in tissue processing for transmission electron microscopy. Microscopy Research and Technique 21, 39-50 Felgenhauer, B.E., Spring, J.H. and Bordelon, C.M. (1996) Improved fixation and contrast of insect Malpighian tubules using osmium tetroxide-potassium ferricyanide. Microscopy and Research Technique 35, 361-362 Fruhling 1 J., Penasse, W., Sand, G. and Claude, A. (1969) Preservation du cholesterol dans la corticosurrenale du rat au cours de la preparation des tissus pour la microscopie electronique. Journal de Microscopie 8, 957-982 Graham, L.L. and Beveridge, T.J. (1990a) Evaluation of freeze-substitution and conventional embedding protocols for routine electron microscopic processing of eubacteria. Journal of Bacteriology 172, 2141-2149 Graham, L.L. and Beveridge, T.J. (1990b) Effect of chemical fixatives on accurate preservation of Escherichia coli and Baallns subtilis structure in cells prepared by freeze-substitution. Journal of Bacteriology 172,2150-2159 Harb, J.M. (1993) Interpretation of TEM micrographs for human diagnosis. MSA Bulletin 23,206-218 Horowitz, R.A., Giannasca, P.J. and Woodcock, C.L. (1990) Ultrastructural preservation of nuclei and chromatin: improvement with low-temperature methods. Journal of Microscopy 157, 205-224 Idelman, S. (1964) Modification de la technique de Luft en vue de la conservation des lipides en microscopie electronique. Journal de Microscopie 3, 715-718

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Idelman, S. (1965) Conservation des lipides par Ies techniques utilisees en microscopie electronique. Histochemie 5, 18-23 Korn, E.D. and Weisman, R.A. (1966) Loss of lipids during preparation of amoebae for electron microscopy. Biochimica et Biophysics Acta 116, 309-316 Lee, R.M.K.W. (1984) A critical appraisal of the effects of fixation, dehydration and embedding on cell volume. In Science of Biological Specimen Preparation, Revel, J.-P., Barnard, T. and Haggis, G.H. (eds.), pp. 61-70, SEM, Inc., AMF O'Hare, Chicago Locke, M. (1994) Preservation and contrast without osmification or section staining. Microscopy Research and Technique 29,1-10 McDonald, K. and Morphew, M.K. (1993) Improved preservation of ultrastructure in difficult-to-fix organisms by high pressure freezing and freeze substitution. Microscopy Research and Technique 24, 465-473 Muller, L.L. and Jacks, T.J. (1975) Rapid chemical dehydration of samples for electron microscopic examinations. Journal of Histochemistry and Cytochemistry 23,107-110 Peters, K.-R. (1980) Improved handling of structural fragile cell-biological specimens during electron microscopic preparation by the exchange method. Journal of Microscopy 118, 429^141 Robards, A.W. and Sleytr, U.B. (1985) Low temperature methods in biological electron microscopy. In Practical Methods in Electron Microscopy, Vol. 10, Glauert, A.M. (ed.), Elsevier, Amsterdam Rostgaard, J. and Tranum-Jensen, J. (1980) A procedure for minimizing cellular shrinkage in electron microscope preparation: a quantitative study of frog gall bladder. Journal of Microscopy 119,213-232 Stein, O. and Stein, Y. (1967) Lipid synthesis, intracellular transport, storage, and secretion. Journal of Cell Biology 33, 319-339 Stein, O. and Stein, Y. (1971) Light and electron microscopical radioautography of lipids: techniques and biological applications. Advances in Lipid Research 9,1-72 Steinbrecht, R.A. (1993) Freeze-substitution for morphological and immunocytochemical studies in insects. Microscopy Research and Technique 24, 488-504 Steinbrecht, R.A. and Muller, M. (1987) Freeze-substitution and freeze-drying. In Cryotechniques in Biological Electron Microscopy, Steinbrecht, R.A. and Zierold, K. (eds.), pp. 149-172, Springer-Verlag, Berlin and Heidelberg Voorhout, W., Van Genderen, I., Van Meer, G. and Geuze, H. (1991) Preservation and immunogold localization of lipids by freeze-substitution and low temperature embedding. Scanning Microscopy Supplement 5, S17-S25 Weibull, C. and Christiansson, A. (1986) Extraction of proteins and membrane lipids during low temperature embedding of biological material for electron microscopy. Journal of Microscopy 142, 79-86 Weibull, C., Christiansson, A. and Carlemalm, C. (1983) Extraction of membrane lipids during fixation, dehydration and embedding of Acholeplasma laidlawii-cells for electron microscopy. Journal of Microscopy 129, 201-207 Weibull, C., Villiger, W. and Carlemalm, E. (1984) Extraction of lipids during freezesubstitution of Acholeplasma laidlawii-cells for electron microscopy. Journal of Microscopy 134, 213-216

5 Embedding methods

The aim of embedding is to replace the dehydrating agent or intermediate solvent with a liquid resin monomer which can then be cured or polymerized to produce a solid block with good sectioning properties. Two types of embedding media are in widespread use in electron microscopy. The earliest to be developed were the acrylic resins, «-butyl and methyl methacrylate, but these were superseded in the late 1950s by the epoxy resins for ultrastructural studies. More recently, advantage has been taken of the fact that some acrylic resins can be polymerized at low temperatures and consequently they have now become popular for immunocytochemical investigations. Other embedding media, such as the polyester and melamine resins, are not widely used and will only be described briefly.

5.1

T h e a t t r i b u t e s o f embedding m e d i a

The general properties required of an embedding medium are considered here, and the characteristics of the epoxy and acrylic resins are compared (Table 5.1), while the procedures for obtaining satisfactory blocks with all the suitable resins are described in detail in the following chapters. No embedding medium possesses all the desired qualities, since some of the properties required for good embedding are incompatible with others. For example, a medium which hardens with little shrinkage will often have a high initial viscosity. 5.1.1

Toxicity

The embedding medium should be safe to handle. Almost all the com­ ponents of epoxy and acrylic resin embedding media are irritant and can be allergenic to sensitive individuals, and so they should all be considered to be

Problems Not possible Good

Good Not possible Very good or good

Good Polar resins stain well Poor. Aromatic components increase stability

Influenced by impurities and oxygen Large; reduced with some cross-linkers Exothermic; rise can be high Most polymerize at low temperatures Some problems with polar resins

Problems Slight None No Good

Good

Good Lipids extracted, particularly by polar resins

Slight None No Excellent

Good Slight

GMA LR White; Lowicryls K4M and KllM All low, minimum 8 for LR White

Some components have limited shelf-life None

GMA, LR resins, Lowicryls

Acrylic resins

Problems Slight

7.8

Durcupan Epon Araldite, >1800

Epon, 140-180

No problems None

Spurr, ULV

No problems Helpful, but not essential

Araldite, Epon, Durcupan

Epoxy resins

ULV, ultra-low viscosity; GMA, glycol methacrylate; cP, centipoise (IcP = 10 ~ 3 pascal.seconds).

Stability under the electron beam

Infiltration: uniformity of extraction during Hardening: uniformity of volume shrinkage during temperature rise during at low temperatures Sectioning Section staining: for contrast for histology

Storage and transport Requirement for intermediate solvent Miscibilitv with water: full partial Viscosity of resin (cP at 25°C)

Property

Table 5.1 Comparative properties of epoxy and acrylic embedding resins

ChapterS:

Embeddingmethods

potentially hazardous. In general, however, with the exception of a limited number of components that are highly toxic or carcinogenic or both, hazards are not a serious problem, so long as the safety precautions described in Sect. 5.2 are taken. The special hazards associated with particular embedding media are described in detail in later chapters. 5./.2

Availability

The components of the medium should be readily available, be uniform from batch to batch and have a long shelf-life. All the media discussed here are available from general suppliers (see Appendix), either as separate components or in kit form, and so high standards should be assured. There are problems with the transport and storage of some components of acrylic embedding media and particularly with the pre-mixed media, which are becoming popular because of their convenience. A survey of embedding kits has shown that the components selected for a kit and the instructions in the accompanying data sheet are not always correct. Reference should be made to the standard embedding media described here. 5.1.3

Uniformity of infiltration

For ease of infiltration into the specimen, the liquid embedding medium should be freely miscible with dehydrating agents and intermediate solvents and should have a low viscosity. The infiltration of the viscous epoxy resins, Araldite and Epon, is assisted by the use of an intermediate solvent, such as propylene oxide, but this is not essential. Intermediate solvents are not required for any of the other resins. The epoxy resin vinylcyclohexene dioxide has a low viscosity. Araldite and Epon have higher viscosities, but these decrease dramatically with increasing temperature and consequently they should be warmed briefly to between 55°C and 60°C just before use. All the acrylic resins used in electron microscopy have low viscosity and this is maintained by some of them at low temperatures. Embedding media with high viscosity can exert mechanical and osmotic stresses and can damage some very delicate specimens during infiltration, and it is sometimes necessary to minimize these effects by suitable modifications of the procedure, such as the use of continuous infiltration, in addition to warming the resin. It is also important that there should not be differences in the mole­ cular sizes and consequently in the rate of penetration of the various

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components of the embedding medium, since this will lead to variations in the final composition of the embedding medium at different sites within the specimen (Causton 1980), and there will be local regions within the tissue where the final block is of a different consistency, because curing or polymerization is incomplete. Differential swelling of tissues and tissue components can then occur as the sections float on the water surface in the knife trough and the staining of the sections will be affected. These problem arise with the low viscosity epoxy resins, such as Spurr's resin and Quetol 651, where there is a large disparity between the molecular sizes of the epoxy resin or flexibilizer and the anhydride hardener, so that they move through the tissue at different rates (Mollenhauer 1988). Similar problems of differential infiltration of resin monomers have not been reported with Araldite and Epon epoxy resins or with acrylic resins, but the choice of tertiary amine for use as an accelerator is important for epoxy resins. They vary in their rate of penetration into a specimen, and uniform blocks are more easily obtained with benzyldimethylamine (BDMA), than with 2,4,6-m's(dimethylaminomethyl) phenol (DMP-30), which has a higher viscosity. 5. 1 . 4

Interactions with the specimen

There should be minimum modification or extraction of specimen com­ ponents during the infiltration of the liquid embedding medium. There is little direct evidence for the chemical interaction of resins with tissues, but epoxy resins are expected to cross-link proteins and to react with nucleic acids (Causton 1986). The most obvious effect of resin infil­ tration is the extraction of lipids, but it is not easy to compare the results of various studies with different embedding media, because the amount of extraction is critically dependent on the preceding fixation

and

dehydration procedures. In spite of these reservations, some general conclusions can be drawn. Most epoxy resins, including Araldite, Epon and Spurr s resin, have relatively little extractive effect during standard embedding pro­ cedures. In contrast, many acrylic resins are powerful lipid solvents and can easily affect the appearance of membranes in specimens not fixed with osmium tetroxide. For example, Fehrenbach et al. (1991) found that the multilamellar bodies of type II mammalian lung alveolar cells, which

Chapter S:

Embedding methods

are rich in phospholipids, do not show any substructure after fixation with glutaraldehyde and uranyl acetate and embedding in Lowicryl K4M, while they are excellently preserved in Araldite. There are considerable variations in the amount of lipid extracted during infiltration with different acrylic resins. The polar Lowicryl K4M always extracts more lipid than the non-polar HM20. LR White is parti­ cularly aggressive and extracts 36 times more 14C-Iabelled compounds from plant tissues than methacrylate (Coetzee and Van der Merwe 1989). It seems likely that this high value for LR White is partly due to the fact that one of the resin monomers contains benzene rings, which confer greater stability on the sections in the electron beam, but also increase lipid solubility. The extraction of lipids by acrylic resins is reduced as the temperature is decreased and becomes negligible if the temperature is low enough. Thus, very little lipid is extracted during infiltration with Lowicryl HM20 at -50°C or below (Weibull and Christainsson 1986). 5. 1.5 Uniformity of hardening The transition from the liquid monomers to a hardened block should be a uniform process with very little volume change, resulting in blocks with good sectioning properties. The hardening process is different for acrylic and epoxy resins. Acrylic monomers are linked, or polymerized, by radical chain reactions in a process which is controlled by free radical-producing initiators and accelerators. Oxygen is a radical scavenger and must be excluded, since it inhibits this process. Polymerization is accompanied by considerable shrinkage (15 to 20% by volume) and is affected by the presence of impurities. The reaction is exothermic, so that heat is generated at sites within the hardening block. In consequence, polyme­ rization is uneven and can result in structural damage to the specimen, which is known as 'polymerization damage'. More uniform blocks are obtained by the addition of cross-linking agents to reduce shrinkage and by the use of low temperatures and efficient heat 'sinks' to remove the heat evolved during polymerization. In contrast the hardening of epoxy resins is obtained by the addition of polyfunctional agents, such as anhydrides, which react with the epoxide and hydroxyl groups of the epoxy resin monomers. This process is not strictly a polymerization, but is an addition reaction which is more

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accurately described as curing. It is accompanied by very little volume shrinkage (about 2% by volume), is not exothermic and is not affected by impurities. In consequence, uniform hardening is easily obtained with well-infiltrated specimens, so long as the temperature is not raised above the advised value, which is usually 60°C. At higher temperatures the addition reaction becomes exothermic with adverse effects on the properties of the final block. 5.1.6

Ease of sectioning

The final block should have good sectioning properties. Blocks of epoxy and acrylic resins can all be sectioned routinely with the excellent ultramicrotomes that are now available (see Reid and Beesley 1991), so long as the standard formulations and methods described in this book are used. The blocks will be of uniform consistency and will be of sufficient hardness for thin sections to be cut easily with little com­ pression. In general, the sectioning properties of the epoxy resins, Araldite and Epon, and the non-polar acrylic resins, such as Lowicryl HM20, are somewhat better than those of the low viscosity epoxy resins, such as Spurr's resin, and the polar acrylic resins, such as Lowicryl K4M and the methacrylates (Acetarin et al. 1987). Sectioning of epoxy resins is easier when the plasticizer, dibutyl phthalate, is present, as in Araldite embedding media. 5.1.7

Section staining

The staining of sections to increase contrast or for cytochemical studies should be uniform. The polar acrylic resins are readily infiltrated by aqueous stains and consequently good contrast is obtained in electron microscopy. In addition, a wide range of cvtological stains is available for sections of media based on glycol methacrylate (Lewis and Knight 1992). Aqueous stains do not infiltrate as readily into non-polar embedding media, such as most epoxy resins, but adequate contrast is obtained with a combination of pre-embedding staining with osmium tetroxide and uranyl acetate and section staining with lead. Problems arise, however, if the infiltration of the embedding medium is not uniform. This occurs when the embedding medium contains components that penetrate at different rates, as discussed in Sect. 5.1.3, or when viscous, non-polar monomers are physically excluded by dense and/or well-hydrated tissue

Chapter 5:

Embeddingmethods

structures (Horobin 1983). Aqueous stains penetrate more readily into structures which are only poorly infiltrated by non-polar embedding media. Caution must therefore always be exercised in the interpretation of the distribution of stains and labels. 5.1.8

Stability in the electron beam

The damage induced in thin resin sections in the electron microscope should be minimized to ensure the preservation of the ultrastructure of well-fixed and well-embedded specimens. The initial and immediate effect of irradiation with electrons is the fragmentation and disruption of molecules and the removal of components of the embedding medium, with a resultant loss of mass and shrinkage of the section. Resistance to this damage has been assessed by various methods, including electron energy loss spectroscopy (Michael Lamvik and Audrey Glauert, un­ published observations, 1991), and has been found to be highest for the epoxy resins, and particularly for the aromatic Araldites, followed by Spurr's resin and the aliphatic Epons. Epoxy resins and anhydride hardeners form three-dimensional polymers which are very stable and maintain their structure during irradiation with electrons, even when there is some loss of mass. This loss is detectable in Epon sections as a slight decrease in density, or 'clearing' of the section in the area irradiated. The acrylic resins are much the most sensitive, since they consist of long chains of linear polymers which are readily broken. In consequence, thev can lose as much as 70% of their mass in the electron microscope under normal operating conditions. This loss is accompanied by a break­ down of the whole structure of the polymer and a flow of material, with consequent structural distortion which is greater than that occurring during uneven polymerization. It does not depend on the degree of polymerization of the acrylic resin or on the amount of cross-linking, since most cross-links are just as vulnerable to damage as the resin monomers themselves. The resistance of the sections to electron irradia­ tion can be increased by the incorporation of aromatic acrylic resins into the formulation and by the use of divinyl benzene as a cross-linking agent. In general, however, contrast is more variable and membranes are less well defined than in epoxy resin sections. The use of minimal dose recording does not reduce shrinkage (Braunfeld et al. 1994). The only effective way to reduce radiation damage

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significantly is to examine the sections at very low temperatures of -120°C or less, using a cold stage in the electron microscope, but this procedure is not practicable for the day-to-day examination of sections for ultrastructural studies. It is preferable to chose a resin with good resistance to beam damage in the first place. As examination of the sections continues, more breakdown occurs and gross changes are observed, such as increased shrinkage, the for­ mation of holes and the movement of the whole section. The absence of these effects is frequently cited as evidence of 'stability' of the sections, but it does not provide evidence that the initial structural damage has been avoided. At very high electron doses the section becomes very hot and all embedding resins are effectively carbonized in much the same way.

5.2

The safe handling of embedding media

All resins and other components of embedding media should be con­ sidered to be hazardous and all workers must be familiar with the safety procedures for handling chemicals described in Sect. 1.4.5 before starting work. Due attention must be given to appropriate storage, to safe handling of embedding resins within the laboratory and to disposal. Resins can usually be stored at room temperature in the air-tight con­ tainers in which they are received, but the instructions of the supplier should always be followed carefully. All containers must be clearly labelled with a detailed list of the contents and any known hazards. Most resin monomers and other components of embedding media are sensitizing when they are allowed to contact the skin, and so ade­ quate protective clothing, including suitable gloves, should be worn. The toxic, irritating and sensitizing powers of resin monomers are greatest when they are diluted with solvents during the infiltration stage (Causton 1988). Resins must not be removed from the skin with an organic solvent, since this will aid penetration of toxic components. Wipe the skin immediately with disposable paper towels and then cleanse the affected area with a resin-removing cream, followed by washing with warm soapy water. Special non-solvent hand cleaners and barrier creams are available from general suppliers (see Appendix).

Chapter 5:

Embeddingmethods

Work with resins in containers and vials is best carried out in a shallow plastic tray using disposable vessels and pipettes, and the working surface of the bench should be covered with an impermeable layer, such as Benchcote (Whatman), to facilitate the removal of any spillages. These should be taken up immediately using paper towels or other absorbent materials, which should then be transferred to a safe container. For disposal, excess mixtures of resins and solvents should be placed in separate waste bottles, as described in Sect. 1.4.5c. Mixtures containing methacrylate and acrylate resins should be placed in tightly-closed containers and kept in a refrigerator. Excess embedding media should always be cured or polymerized before disposal, since they are then relatively inert. Acrylic resins must be polymerized in small quantities, since some of the reactions are exothermic and a 'runaway' reaction can occur when polymerizing large amounts of waste resin. Some procedures can be carried out in a well-ventilated laboratory, but the vapours of many embedding media are sensitizing when they are inhaled, particularly when they are hot. Embedding ovens should be vented directly to outside the laboratory or should be placed in a fume cupboard with a good air flow which is checked regularly. A further hazard arises during sectioning since dust particles from resin blocks, and particularly from Spurr's resin, are potentially sensitizing. Blocks should be trimmed with a single-edged razor blade or on an ultramicrotome, and not with a saw, unless there is a very efficient system for dust extraction. Details of the hazards associated with different types of embedding media are given in the following chapters.

5.3

Embedding procedures

The standard method for embedding specimens in capsules or embedding moulds is similar for all types of embedding medium. After fixation (Chapter 3) and dehydration (Chapter 4), the specimens are infiltrated with the embedding medium. Subsequently epoxy resins are cured and acrylic resins are polymerized by heating in an embedding oven or by irradiating with UV light to produce blocks for sectioning. The suppliers of equipment for embedding are listed in the Appendix.

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Biological specimen preparation for TEM

Infiltration of embedding media

The aim of the procedure is to obtain uniform penetration of the embedding medium throughout the specimen. This is achieved by gradually replacing the dehydrating agent, or the intermediate solvent, with the embedding medium. 5.3.1a

S t a n d a r d infiltration m e t h o d s

At the start of the standard procedure the specimens will be in the vials, microtubes or other plastic containers in which they were dehydrated. The dehydrating agent is removed and then the specimen is passed through a series of mixtures of the dehydrating agent and the liquid embedding medium, until it is in the pure embedding medium. The speci­ mens are then transferred to capsules or embedding moulds for curing or polymerization of the resin. The timing and number of steps in the process depend on the type of embedding resin and on the size and nature of the specimen. For example, longer infiltration times are required for large specimens and dense tissues, and when thick cell walls are present, as in plants. Infil­ tration of embedding resins can be done in an automatic processor, such as the Leica Lynx (see Sect. 3.4), but these units are expensive and so are only justified when large numbers of samples are processed routinely. Furthermore, modifications to the standard embedding media and processing schedules proposed in this book may well be required to obtain consistent results. 5.3.1b

Continuousinfiltration

Some delicate tissues and cells are susceptible to damage during infiltration with viscous epoxy resins, and procedures involving the continuous increase in the concentration of the embedding medium are then preferable to the standard step-wise method. No sudden changes in viscosity then occur and the specimen is not subjected to mechanical or osmotic stress. Overall shrinkage of the specimen is also reduced. A simple and well-tried method is to use a specimen rotator (Steinbrecht and Ernst 1967). After dehydration in ethanol and propylene oxide, tissue pieces are placed in 2 to 4 ml of propylene oxide in vials on a rotator in an efficient fume cupboard. The liquid should completely cover the bottoms of the vials when they are in the inclined position. The device is

ChapterS: Embeddingmethods

set to rotate slowly and then a similar quantity (2 to 4 ml) of complete epoxy resin embedding medium is added dropwise. The uncovered vials are kept rotating overnight (minimum 8 h), allowing the propylene oxide to evaporate almost completely. A change of complete resin mixture can then be made before curing, but this is not essential, since small residual amounts of propylene oxide do not affect the curing of epoxy resins (Sect. 4.3.1). Another simple method of continuous infiltration is described by Marchese-Ragona and Johnson (1982) and is suitable for low viscosity epoxy resins. A 4 to 6 mm layer of pure ethanol is placed on top of a 15 to 20 mm layer of Spurr's resin in a glass vial with an air-tight cap. The vial is shaken very gently to cause the boundary between the ethanol and the resin to become diffuse, resulting in an ethanol/resin layer 1 to 2 mm in depth. The dehydrated specimen is transferred from 100% ethanol to the vial and sinks until it meets the ethanol/resin gradient. As the exchange from ethanol to resin proceeds the specimen sinks into pro­ gressively increasing concentrations of resin until it reaches pure Spurr's resin. The specimen is then passed through 2 or 3 changes of fresh resin before curing. More complex systems have been devised by Peters (1980) and by Rostgaard and Tranum-Jensen (1980). 5.3.1c Vacuuminflltration The application of a slight vacuum sometimes aids the infiltration of the embedding medium into large or dense specimens, or into those that contain large air spaces, such as botanical or freeze-dried tissues, al­ though this is not necessary for the majority of specimens. It is only necessary to apply the vacuum at the initial steps of the procedure to accelerate the replacement of the dehydrating agent by the resin, other­ wise there is a danger that volatile components of the embedding medium, such as accelerators, will be removed, resulting in significant changes in the resin formulation and unsatisfactory blocks (Mollenhauer 1993). Consequently vacuum infiltration is not suitable for low viscosity epoxy resins containing the accelerator dimethylaminoethanol, such as Spurr's original mixture, or for acrylic resins. Vacuum embedding chambers are available commercially (see Appendix), but the simplest method of vacuum infiltration is to place the specimens in the infiltrating solutions in uncovered containers in a vacuum desiccator or an old plate desiccator. The pressure is reduced to

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about half an atmosphere (or 380 mm Hg), but not less, by pumping with a rotary pump, and this partial vacuum is maintained during infil­ tration with the embedding medium. The vacuum should be released gently over several minutes at the end of each stage of infiltration. 5.3.2

Embedding moulds

Specimens are embedded in capsules or flat embedding moulds to produce blocks of a convenient size and shape for ultramicrotomy (Reid and Beesley 1991). Gelatin capsules are inexpensive and give cylindrical blocks with dome-shaped ends (Fig. 5.1) in a range of sizes, as listed in Table 5.2. Sizes 00 and 0 are convenient for general use. Gelatin (or polypropylene) capsules are suitable for embedding in acrylic resins, since they are impervious to gases and so completely exclude oxygen. Specially designed polyethylene capsules, such as BEEM capsules, produce blocks with ends in various forms, such as a truncated pyramid or conical tip (Fig. 5.2), which minimize the amount of trimming required before sectioning and limit the movement of the specimen within the capsule. These capsules have hinged caps and are available in sizes 00 and 3, together with BEEM capsule holders which hold a number of capsules upright during filling with resin and curing. Polypropylene capsules of similar design are available for embedding at the higher temperatures that can be reached during the polymerization of acrylic resins.

Table 5.2 The sizes of gelatin capsules Capsule size

Volume

Approximate diameter

ml

mm

000

1.37

9.5

00

0.95

8.5

0

0.68

7.0

1

0.50

6.5

2

0.37

6.0

3

0.30

5.5

4

0.21

5.0

Data from Agar Scientific: catalogue number 6.

Chapter 5:

Embedding methods

Fig. 5.1 Gelatin capsules giving cylindrical blocks with dome-shaped ends. (Courtesy of Agar Scientific.)

Fig. 5.2

Polyethylene BEEM capsules of different shapes. (Courtesy of Agar Scientific.)

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Flat embedding techniques have become increasingly popular and only very small specimens, such as centrifuged pellets of cells and organelles and small tissue pieces, which do not require to be sectioned in a particular orientation, are embedded in capsules. Flat embedding moulds are particularly suitable for long specimens and for thin tissue slices. They allow more accurate orientation of the specimen in the block than capsules and they are reusable a number of times. The standard moulds for embedding in epoxy resins are made of flexible white silicone rubber and have an array of cavities, 3 to 5 mm deep (Fig. 5.3.). The moulds have a range of designs to provide blocks with different shapes for sectioning. Some cavities are numbered to aid the recording of samples and others are larger and enable a number of specimens to be embedded in one block. Spurr's resin sometimes interacts with white silicone rubber and is cured in special green rubber moulds, such as those supplied by Agar Scientific, which are more resistant to attack and so have a longer life. Acrylic and melamine resins also interact with silicone rubber and in addition the moulds have to withstand the high temperatures that can be reached during polymerization. Consequently these resins are poly­ merized in polypropylene moulds. Transparent versions of flat embedding moulds are available. These make the specimens more easily visible and are suitable for polymerization of resins by UV irradiation.

Fig. 5.3

Shaped silicone rubber moulds. (Courtesy of Agar Scientific.)

ChapterS:

Embeddingmethods

Fig. 5.4 A set of specimens embedded in a block of Lowicryl K4M polymerized in an aluminium dish.

All these types of capsules and embedding moulds are obtainable from general suppliers (see Appendix). A number of other containers have been found to be useful as embedding moulds, including polyethylene weighing dishes (Glauert 1975). Aluminium dishes, which are available in two sizes of 53 mm base diameter X 17 mm high, or 43 mm X 12.5 mm, are suitable for embedding a number of specimens at low temperature in Lowicryl K4M (Fig. 5.4). It is also possible to make moulds of the required size in the laboratory from aluminium foil by wrapping it around metal blocks of the required dimensions and then pressing the blocks onto a hard surface. The caps of BEEM polyethylene capsules make convenient flat embedding moulds for small specimens, as do the polyethylene caps of some glass vials. The capsules and embedding moulds must be completely dry and gelatin capsules are best stored in a desiccator or heated in an oven at 60°C, overnight or longer, before use. 5.3.3

Curing and polymerization o f embedding media

5.3.3a Embedding in capsules

Capsules are mounted in racks, such as BEEM capsule holders, in preparation for the transfer of specimens and filling with resin. Transparent versions are available for UV polymerization. Alternatively

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a holder for capsules is easily made in the laboratory by punching an array of holes in the lid of a cardboard box (Fig. 5.5). Following infiltration with the embedding medium, each specimen is removed from the vial with fine tweezers or a pipette and placed in the bottom of a capsule. The capsule is filled about two-thirds full with fresh embedding medium and the specimen is guided back to the exact bottom of the capsule, if necessary. Viscous embedding media should be warmed to about 60°C just before use. A label indicating the experiment and sample number is placed in each capsule in such a position that the number is visible from outside the capsule (Fig. 5.5). Use a pencil or typewriter and good quality glazed paper or hardened filter paper. Do not include more information than is necessary. Most details, such as the date, are best recorded in a laboratory notebook. It is usually advisable not to include experimental details, such as length of incubation or type of treatment, since this may inadvertently influence the microscopist during viewing and recording of electron micrographs. Labels should be as 'blind' as possible. A typical label will indicate the subject of the investigation or the initials of a collaborator (e.g. AW), the number of the experiment in the series (e.g. 19), and the number of the sample in the

Fig. 5.5

A cardboard lid makes a convenient holder for a set of capsules during curing of

an epoxy embedding medium. A label is about to be inserted into one of the capsules.

Chapter 5:

Embedding methods

experiment (e.g. 3); so the label will be AW.19.3, as in Fig. 5.5. The choice of labelling method depends on the working practices within the laboratory and only general guidelines are suggested here. More details, such as the species of animal or type of experiment, may have to be included, using an alphabetic code, when sections are to be examined by a number of different microscopists. It is strongly recommended that the final prints are labelled on the front, with the exposure number, the magnification and the experiment number, so that it is not necessary to keep turning the print over to find this information. A cap is not required for capsules containing Araldite and Epon epoxy resins, but it is advisable to cover Spurr's medium and its modifications, since some of the components of the medium are volatile. Capsules containing acrylic resins are filled as full as possible and the caps are placed on firmly to exclude oxygen entering from the air. These precautions are not necessary if the embedding oven is filled with nitrogen gas. 5.3.3b

Flat embedding

Specimens are transferred to flat embedding moulds in the same way and orientated as required. A potential problem with these moulds is that the tissue may sink to the bottom of the mould during curing or poly­ merization. This can cause difficulties when trimming, since the region of interest may lie at the bottom of the block. One solution to this problem when using an epoxy resin, such as Araldite, is to half fill the mould with resin and cure it. The tissue is then transferred to this hard base and the mould is filled up with fresh resin, with the result that the specimen is embedded in the middle of a flat block. Alternatively, the first layer of resin is not cured completely, but is only allowed to become too viscous for the specimen to sink into it. It is then easier to obtain a block of uniform hardness. Specimens sometimes move as hardening of the embedding medium commences and it is then necessary to adjust the position and orientation of the specimen in the mould when the sur­ rounding medium has just started to become viscous. Vibratome tissue slices and similar specimens tend to curl during fixation and dehydration. They can be flattened by placing them in a small drop of Araldite, or other epoxy resin, on the end of an Araldite block which has been trimmed on an ultramicrotome to produce a flat surface. A carbon-coated coverslip, 10 mm in diameter, is then placed

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carbon-side down on top of the slices to flatten them. The Araldite is cured by heating at 60°C for 24 to 48 hours and the carbon-coated coverslip is removed with a little mechanical pressure. If necessary, the coverslip can be immersed briefly in liquid nitrogen to separate it from the Araldite block. 5.3.3c

Methods of curing and polymerization

The embedding medium is cured or polymerized by placing the capsules and embedding moulds in an oven at the selected temperature (usually 60°C) for the required time. In general, a higher temperature should not be used, since this will modify the hardening process. More cross-links may be formed and the blocks will become brittle and less easy to section. In exceptional circumstances, such as when results are required quickly, the temperature is increased, but then the specimens should be as small as possible, with a maximum thickness of 0.1 mm in at least one dimension. An epoxy resin must then be used, since higher temperatures enhance the uneven polymerization of acrylic resins and the poor preser­ vation of ultrastructure. When a vacuum oven (see Appendix) is available, the pressure can be reduced to about half an atmosphere (or 380 mm Hg) for the first 10 to 15 min to aid the final infiltration of the resin, but there are inherent dangers from hot vapours and so this should only be done when it is absolutely necessary and not routinely. Capsules containing acrylic resins should be placed in holes drilled in an aluminium block so that the heat evolved during polymerization is conducted away as efficiently as possible and all the resin is kept at a uniform temperature. Any unwanted resin remaining at the end of the experiment should be cured or polymerized before disposal, since it is then relatively inert. 5.3.3d

Ultra-violet and microwave irradiation

Embedding media can also be cured or polymerized by UV irradiation, instead of by heat. A long wavelength of 350 to 360 nm and a low inten­ sity are required. Suitable lamps, with 6 W fluorescent tubes, are available from general suppliers (see Appendix). The capsules or embedding moulds are placed at a distance of 250 to 300 mm from the lamp. Uniform irradiation of the specimens is required for consistent results and is more easily obtained in a special chamber, such as the Agar

ChapterS:

Embeddingmethods

polymerization unit (see Appendix). Alternatively a suitable chamber can be constructed in the laboratory, such as the unit illustrated in Fig. 8.10 in Sect. 8.8.2. The only advantage of UV irradiation is that it enables resins to be hardened at lower temperatures. This is of little value in the curing of epoxy resins, since the temperature cannot be reduced much below room temperature and the process is a lengthy one. In contrast, some acrylic resins can be polymerized at very low temperatures by UV irradiation and this procedure and the special apparatus required are described in detail in Sect. 8.8. Resins can also be cured or polymerized by irradiation with electro­ magnetic waves in a microwave oven of the type used for fixation (Sect. 3.11), with the advantage that the procedure is very rapid. It is essential to use a commercial unit designed for laboratory use (see Appendix) to obtain reproducible results. The electromagnetic field is not uniform and is affected by the samples and by any other items, such as a water load, placed in the oven. Consequently the oven has to be calibrated before each embedding to locate suitable positions for the embedding moulds, thus increasing the total time required for curing or polymerization. A detailed description of the necessary procedures is given by Login and Dvorak (1994) and their 'Tool Book' should be consulted before using a microwave oven for embedding. Promising results have been obtained with Epon media. Experiments have also been made with Spurr's epoxy resin and with acrylic resins, but the dangers associated with the hot vapours of these resins are such that these procedures should be dis­ couraged. It is always essential to operate a microwave oven in an efficient fume cupboard and to follow all the safety precautions discussed in Sect. 5.2.

5.4

Embedding methods for cell monolayers

Cell monolayers pose particular problems during embedding, especially when the study concerns the interaction between the cells and the sub­ strate on which they have grown. These very thin layers of cells are easily disturbed (Fig. 5.6) and so it is preferable not to remove them from the substrate before embedding. Consequently the substrate is chosen so

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Fig. 5.6 An electron micrograph of the junction between two human endothelial cells grown on a Falcon tissue culture dish. The cells were fixed and dehydrated in situ and then floated off the dish in propylene oxide, before embedding in Epon. The layer of extracellular material (arrowhead) which had coated the culture dish is no longer flat, indicating that the monolayer had been disturbed during its release from the dish. Scale bar = 0.5 μπι. (Reproduced from Kuhn 1981, with permission.)

that the hardened block can be separated from it or the cells are grown on a substrate which can be sectioned. 5.4.1

Removal of hardened blocks from the substrate

The separation of an embedded monolayer from the substrate is easy when the cells are grown on a plastic surface. Most types of culture dishes and coverslips are suitable, including those made from poly­ styrene (e.g. Falcon), polyethylene (Thermanox) and polyester (Melinex) (see Appendix). Alternatively the cells are cultured in the snap-on cap of a BEEM capsule (Beesley 1978). The monolayer of cells is fixed, dehydrated and infiltrated with the embedding medium (which is usually Araldite and/or Epon) by standard methods. Gelatin or BEEM capsules are then filled with the embedding medium and are inverted over selected areas of the monolayer. The embedding medium is then cured in the usual way and the capsule is removed from the substrate, leaving the monolayer embedded in the surface of the block. It is easy to remove capsules containing Epon (but not Spurr's resin) if this is done after about 12 hours at 60°C before the

ChapterS: Embeddingmethods

resin has completely hardened. After removal the blocks are returned to the embedding oven for final curing. Alternatively, fully hardened blocks are separated from the substrate immediately after removal from the embedding oven, while they are still warm. Acrylic resins shrink during polymerization and separate more easily from surfaces than epoxy resins, so that coverslips can be gently levered away from the block with the corner of a razor blade. In this 'inverted capsule' technique the layer of cells is embedded in the surface of the block and sections can only be cut parallel to the plane of the monolayer. Very careful and accurate microtomy is required to obtain the sections. Sectioning is easier if a flat embedding method is used and the surface of the block is covered with a thin layer of resin. The monolayer on a plastic coverslip is processed through fixation, dehydration and infiltration with resin in the dish and then the coverslip is removed and inverted over a flat embedding mould, containing fresh embedding medium. The embedding medium is then cured, the block removed from the mould and the coverslip separated from the surface of the block. The monolayer of cells at the surface of the resultant thin resin sheet is then covered with another layer of resin. The simple method (Fig. 5.7) developed by Audrey Glauert and Ron Parker (unpublished observations, 1980) for cells cultured on Melinex (Fig. 5.8) is described here to provide general guidelines. The method is as follows: 1. Preparation of Melinex coverslips for cell cultures. Cut out square, 15 X 15 mm, coverslips from a Melinex sheet, 100 to 175 μηι thick. Clean each coverslip by wiping it with tissue soaked in ethanol. Sterilize in ethanol (70%), two or more changes, or store in 70% ethanol. Air dry and keep sterile. Place in a Petri dish and incubate in serum-containing medium, overnight at 37°C, to remove toxic substances. Remove the medium and wash in fresh medium. Add cells and incubate in the usual way. 2. Fixation and dehydration. Fix the monolayer at the incubation temperature by immersion in an aldehyde fixative as described in Sect. 3.9. Remove the dish from the incubator and replace the fixative with buffer, containing calcium chloride, at room temperature. The fixed cultures can be stored at 4°C in buffer, so long as it contains 2 to 3 mM calcium chloride to stabilize lipids.

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(d)

(f)

(g) 1

Fig. 5.7

i

Embedding method for a monolayer of cells cultured on a square plastic

(Melinex) coverslip. (a) The cells are fixed, dehydrated and infiltrated with an epoxy resin, such as Araldite, in a small glass vial, (b) The coverslip is removed from the vial and (c) inverted over the plastic cap of a specimen vial containing epoxy resin, (d) The resin is cured and (e) the Melinex coverslip is peeled away, (f) The resin block is removed by bending the plastic cap, and (g) the block is now ready to be re-embedded in a flat embedding mould, with the cell layer uppermost.

Chapter 5:

Embeddingmethods

Transfer the monolayer of cells on the Melinex coverslip to a glass vial (Fig. 5.7a) containing buffer and then fix it in osmium tetroxide, wash in water, fix with uranyl acetate, dehydrate in ethanol and propylene oxide (which does not affect Melinex) and infiltrate with Araldite and/or Epon by standard methods. 3. Embedding. Remove the coverslip from the vial (Fig. 5.7b) and invert it over a resin-filled plastic container, such as the cap of a specimen vial (Fig. 5.7c), and cure the resin (Fig. 5.7d). The Melinex is then easily peeled away (Fig. 5.7e). Remove the block containing the embedded monolayer by bending the plastic mould (Fig. 5.7f). Place the block (Fig. 5.7g) in a flat embedding mould with the cell layer uppermost and cover it with a thin layer of fresh embedding medium. Cure the resin to produce a block in which the monolayer is protected during sectioning. 4. Sectioning. Examine the resulting disc by light microscopy and mark selected cells with a felt-tip pen. Cut out chosen areas of the monolayer into small blocks, 1 to 2 mm 3 in size, with a small saw and section them in the plane of the monolayer or at right angles to it. Melamine resins are also suitable as substrates for cell cultures (Westphal et al. 1988). Melamine is not available in the form of a coverslip, but a thin foil is prepared by dipping a glass slide or coverslip into a melamine solution. The coverslips are flamed for sterilization and to harden the melamine. Care must be taken at this stage because melamine produces cyanide gas when it burns. 5.4.2

Monolayer embedded and sectioned on the substrate

When the substrate can be sectioned it does not have to be removed from the embedded layer of cells. Damage to the cells is avoided and it is possible to study the relationships between the cells and their substrate in detail. The substrate must be suitable for the good attachment and growth of cells and must section easily without separating from the monolayer. Sectioning is not easy with some of the earlier substrates, such as Nucleopore filters, silicone rubber membranes and Falcon plastic Petri dishes (see Glauert 1975 for references) and these have been super­ seded by thin coverslips made of Araldite or Epon. Alternatively cells are cultured on permeable substrates, such as Millipore filters or dialysis

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Fig. 5.8 An electron micrograph of a macrophage cultured on a layer of fibrin (F) on a Melinex coverslip. The position of the coverslip, before its removal during embedding, is visible as a straight dense line (arrowhead), indicating that there has been no disturbance of the relationship benveen the cell and its substrate during processing. Scale bar = 0.5 μηι.

membranes. These allow better diffusion and infiltration of culture media, fixatives and embedding media. Slides made out of epoxy resins, such as Araldite, Epon or Spurr's resin, make excellent substrates, so long as care is taken to remove all toxic factors by treatment with salt solutions before the cells are added. Special moulds for making epoxy resin slides are available from some

Chapter 5:

Embedding methods

Fig. 5.9 A baby hamster kidney cell cultured on an Epon substrate and embedded in Epon. Scale bar = 2 μηι. (Reproduced from Grinnell et al. 1976, with permission.)

general suppliers (see Appendix), but it is easy to make a slide by curing a thin (2 to 4 mm) layer of the epoxy resin in a container with a flat level bottom, such as one of the larger flat silicone rubber embedding moulds (Sect. 5.3.2). The epoxy resin slides are sterilized by irradiation with UV light and transferred to a polystyrene tissue culture dish. The cells are added and incubated for the required time and are then fixed, dehydrated and infiltrated with the same epoxy resin in situ (Fig. 5.9). The monolayer of cells is then embedded by covering it with a thin, 0.1 to 0.5 mm, layer of embedding medium, or by inverting the slide over a flat mould containing the embedding medium. The resulting 'sandwich' is sectioned in the same way as in the flat embedding method (Sect. 5.4.1). Alternatively, monolayers grown on resin strips, 10X4 X 3 mm, are processed in small vials and the strips are placed longitudinally in BEEM capsules for embedding (Grinnell et al. 1976). The same Epon mixture is used for embedding as was used for the preparation of the slides to ensure that the final 'sandwich' has a uniform hardness and consistency.

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Millipore filters (see Appendix) are suitable substrates, and in general small pore sizes of 0.22 or 0.45 μιτι are chosen, since larger pores allow the cells to migrate into the filter. Propylene oxide dissolves the filter and is avoided during processing. The whole filter is embedded in Araldite or Epon in a flat embedding mould, or the filter is cut into a number of strips which are packed into a BEEM capsule for embedding. Dialysis membranes also provide a permeable substrate which can be sectioned (Hackett and Sullivan 1982; Muller et al. 1991). They are transparent and resistant to the solvents used during processing and can be embedded and sectioned without distortion.

References Acetarin, J.D., Carlemalm, E., Kellenberger, E. and Villiger, W. (1987) Correlation of some mechanical properties of embedding resins with their behaviour in microtomy. Journal of Electron Microscopy Technique 6, 63-79 Beesley, J.E. (1978) A new technique for preparing cell monolayers for electron microscopy. Stain Technology 53, 48-50 Braunfeld, M.B., Koster, A.J., Sedat, J.W. and Agard, D.A. (1994) Cryo automated electron tomography: towards high-resolution reconstructions of plastic-embedded structures. Journal of Microscopy 174, 75-84 Causton, B.E. (1980) The molecular structure of resins and its effect on the epoxy embedding resins. Proceedings of the Royal Microscopical Society 15,185-189 Causton, B.E. (1986) Does the embedding chemistry interact with tissue? In The Science of Biological Specimen Preparation, Muller, M., Becker, R.P., Boyde, A. and Wolosewick, J.J. (eds.), pp. 209-214, SEM Inc., AMF O'Hare, Chicago Causton, B.E. (1988) The hazards associated with embedding resins. Microscopy and Analysis, January 1998,19-21 Coetzee, J. and Van der Merwe, C.F. (1989) Extraction of carbon 14-labeled compounds from plant tissue during processing for electron microscopy. Journal of Electron MicroscopyTechniquell, 155-160 Fehrenbach, H., Richter, J. and Schnabel, Ph.A. (1991) Improved preservation of phospholipid-rich multilamellar bodies in conventionally embedded mammalian lung tissue - an electron spectroscopic study. Journal of Microscopy 162, 91-104 Glauert, A.M. (1975) Fixation, dehydration and embedding of biological specimens. In Practical Methods in Electron Microscopy, Vol. 3, Part I, A.M. Glauert (ed.), NorthHolland, Amsterdam Grinnell, F., Tobleman, M.Q. and Hackenbrock, C.R. (1976) Initial attachment of baby hamster kidney cells to an epoxy substratum. Journal of Cell Biology 70, 707-713 Hackett, C.J. and Sullivan, K. (1982) A convenient culture chamber for observation and embedding of macrophage monolayers for transmission electron microscopy. Journal of Microscopy 126, 207-210 Horobin, R.W. (1983) Staining plastic sections: a review of problems, explanations and possible solutions. Journal of Microscopy 131, 173-186

Chapter 5:

Embedding methods

Kuhn, H. (1981) A simple method for the preparation of cell cultures for ultrastructural investigation. Journal of Histochemistry and Cytochemistry 29, 84-86 Lewis, P.R. and Knight, D.P. (1992) Cytochemical staining methods for electron microscopy. In Practical Methods in Electron Microscopy, Vol. 14, A.M. Glauert (ed.), Elsevier, Amsterdam Login, G.R. and Dvorak, A.M. (1994) The Microwave Tool Book. Beth Israel Hospital, Boston Marchese-Ragona, S.P. and Johnson, S.P.S. (1982) A simple method for the progressive infiltration of resin into a dehydrated biological sample. Proceedings of the Royal Microscopical Society 17, 311-312 Mollenhauer, H.H. (1988) Artifacts caused by dehydration and epoxy embedding in transmission electron microscopy. In Artifacts in Biological Electron Microscopy, R.F.E. Crang and K.L. Klomparens (eds.), pp. 43-64, Plenum Press, New York and London Mollenhauer, H.H. (1993) Artifacts caused by dehydration and epoxy embedding in transmission electron microscopy. Microscopy Research and Technique 26, 496-512 Muller, W.H., Van der Krift, T.P., Knoll, G., Smaal, E.B. and Verkleij, A.J. (1991) A preparation method of specimens of the fungus Peniallium chrysogenum for ultrastructural and immuno-electron microscopical studies. Journal of Microscopy 164, 29—41 Peters, K.-R. (1980) Improved handling of structural fragile cell-biological specimens during electron microscopic preparation by the exchange method. Journal of Microscopy 118, 429—441 Reid, N. and Beesley, J.E. (1991) Sectioning and cryosectioning for electron microscopy. In Practical Methods in Electron Microscopy, Vol. 13, A.M. Glauert (ed.), Elsevier, Amsterdam Rostgaard, J. and Tranum-Jensen 1 J. (1980) A procedure for minimizing cellular shrinkage in electron microscope preparation: a quantitative study of frog gall bladder. Journal of Microscopy 119, 213-232 Steinbrecht, R.A. and Ernst, K.-D. (1967) Continuous penetration of delicate tissue specimens with embedding resin. Science Tools (LKB), p. 24 only Weibull, C. and Christiansson, A. (1986) Extraction of proteins and membrane lipids during low temperature embedding of biological material for electron microscopy. Journal of Microscopy 142, 79-86 Westphal, C., Horler, H., Pentz, S. and Frosch, D. (1988) A new method for cell culture on an electron-transparent melamine foil suitable for successive LM, TEM and SEM studies of whole cells. Journal of Microscopy 150, 225-231

6 Embedding in epoxy resins

The epoxy resins have considerable advantages as embedding media for electron microscopy, in comparison with the acrylic resins, as discussed in Sect. 5.1. They change very little in volume during curing, harden uniformly, have excellent sectioning properties and are very stable in the electron microscope. Consequently they are the resins of choice when the accurate preservation of ultrastructure is paramount.

6. /

The components o f epoxy resin embedding media

These embedding media consist basically of the epoxy resin itself and an anhydride hardener, which reacts with the epoxide and hydroxyl groups of the resin molecules to give three-dimensional, so-called cross-linked, structures (Glauert and Glauert 1958). This process is not strictly a poly­ merization, but an addition reaction, which accounts for the small accompanying shrinkage. The high temperatures required for the reaction can affect components of biological specimens, and so a tertiary amine is added as an accelerator to enable curing to be obtained in less than 24 hours at temperatures as low as 60°C. These compounds pro­ mote epoxy-to-epoxy and epoxy-to-hydroxyl reactions, but do not themselves serve as direct cross-linking agents. Various plasticizers, reactive flexibilizers, or other modifiers, are added to reduce the visco­ sity of the embedding medium and to improve the sectioning properties of the final block. All the components of epoxy resin embedding media are irritating and potentially allergenic, particularly when they are hot, and it is important to always follow the safety precautions outlined in Sect. 5.2. The hazards associated with particular components are listed in Table 6.1.

Glauert and Lewis:

Biological specimen preparation for TEM

Table 6.1 Hazards in handling epoxy resin embedding media Component

Examples

Potential hazards

Resins

Aromatic 3

Araldite

Low toxicity

Aliphatic' 5

Epon

Low toxicity; hot vapours may be carcinogenic

Based on VCHD

Spurr

Highly toxic and carcinogenic

DDSA, etc

Hot vapours toxic

BDMA, etc

Hot vapours highly toxic

BDE

Little information available, but probably mildly toxic

BGE

Highly toxic, possibly carcinogenic

Plastidzers

DBP

Vapours may be mutagenic

Intermediate solvents

Propylene oxide

Highly volatile and flammable; carcinogenic

Hardeners

Aliphatic anhydrides 0 Accelerators

Amines^ Flexibiltzerse

The information about hazards with these components of embedding media is incomplete, and only the known hazards are listed here. All the components are, however, irritant and potentially allergenic. a.Table6.6; b.Table6.8; c.Table6.2; d.Table6.3; e.Table6.4.

6.1.1

Epoxyresins

An epoxide is a compound containing an oxygen atom bonded in a triangular arrangement to two carbon atoms in an oxyrane ring structure (Fig 6.1a). The adjective epoxy indicates a compound which is related to or is derived from an epoxide, while an epoxy resin is a synthetic thermosetting resin containing epoxy groups. The epoxy resins in embedding media for electron microscopy are of three basic chemical types (Causton 1980,1981):

Chapter 6:

Embedding

in epoxy resins

(a) The diglycidyl ether of bisphenol A (DGEBA) (Fig. 6.1a) is the product of the reaction between bisphenol A and epichlorohydrin. It is an aromatic epoxy resin with two types of chemically reactive group: epoxide end groups and hydroxyl groups spaced along the length of the chain. The viscosity of the resin depends on its molecular weight and increases with the value of n. Consequently a resin with n = 0 is to be preferred for electron microscopy. The aromatic bisphenol A moiety gives the cured resin both toughness

Fig. 6.1

T h e structural f o r m u l a e of epoxy resins.

(a) Diglycidyl ether of bisphenol A ( D G E B A ) . T h e o x y r a n e ring is s h o w n in bold at the beginning of the f o r m u l a . (b) Triglycidyl ether of glycerol. (c) Vinylcyclohexene dioxide ( V C H D ) .

I 77

I

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Fig 6.2

Biological specimen preparation for TEM

T h e s t r u c t u r a l f o r m u l a e of a n h y d r i d e h a r d e n e r s f o r e p o x y resins.

(a) D o d e c e n y l s u c c i n i c a n h y d r i d e ( D D S A ) . (b) M e t h y l N a d i c a n h y d r i d e ( M N A ) . (c) N o n e n y l s u c c i n i c a n h y d r i d e ( N S A ) . (d) « - H e x e n y l s u c c i n i c a n h y d r i d e ( H X S A ) . (e) n - O c t e n y l s u c c i n i c a n h y d r i d e ( O S A ) .

Chapter 6:

Embeddinginepoxyresins

and stability under irradiation by the electron beam (Sect. 5.1.8). The use of Araldite (Ciba-Geigy) bisphenol A resins in embedding media is particularly well established. (b) The di- and tri-glycidyl ethers of diols and polyols are aliphatic epoxy resins with lower beam stability than the aromatic resins (Sect. 5.1.8). Epon 812 (Shell) (Fig. 6.1b) and its various 'replacements', which will all be referred to as 'Epon', are of this type. In general they have lower viscosities than the bisphenol A resins. (c) Vinylcyclohexene dioxide (VCHD), a cycloaliphatic diepoxide (Fig. 6.1c), is the basis for the low viscosity epoxy resin embedding media. It has a compact structure and yields highly cross-linked polymers with good high temperature resistance and stability in the electron beam. Resins based on bisphenol A are relatively inert, although they are capable of causing skin sensitization and the inhalation of hot vapours must be avoided. Some resins of the 'Epon' type are classified as mildly carcinogenic, while YCHD is a known carcinogen. 6.1.2

Hardeners for epoxy resins

The hardeners in epoxy resin embedding media are usually acid anhydrides (Table 6.2), that is, substances obtained by removing the elements of water from a compound, especially from an acid. They are prone to hydrolysis and should be stored dry at room temperature. The hot vapours of these anhydrides are toxic and so special care has to be taken during the curing of the embedding medium. Dodecenylsuccinic anhydride (DDSA) was selected for the original Glauert Araldite formulation (Glauert et al. 1956; Glauert and Glauert 1958). It is a long-chain anhydride (Fig. 6.2a) and so confers plasticity on the cured resin. DDSA is a low temperature hardener and is therefore reactive at 60°C. It is the only hardener required in embedding media based on the aromatic Araldite resins and in the mixtures of Araldite and 'Epon' introduced by Mollenhauer (1964). Embedding media based on Epon 812 and similar aliphatic epoxy resins do not give blocks of sufficient hardness with DDSA alone and consequently Luft (1961) proposed the use of a second hardener, methyl endo-methylene tetra-hydro-phthalic anhydride, which is known as methyl Nadic anhydride (MNA). Nadic is the registered trade mark of

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Table 6.2 Anhydride hardeners for epoxy resin embedding media

Dodecenylsuccinic anhydride

DDSA

Viscosity

Density

cP at 25°C

g/ml

290-295

1.00

MW

266

Methyl Nadic anhydride

MNA

225-275

1.24

178

Nonenylsuccinic anhydride

NSA

117-120

1.00-1.05

224

Nonenylsuccinic anhydride"'

NSA

103

1.08

224

J?-Octeny lsuccinic anhydride

OSA

32

1.00

210

n-Hexenylsuccinic anhydride

HXSA

20-26

1.00-1.10

182

cP, centipoise (1 cP=10 s NSA,

3

pascal-seconds); MW, molecular weight.

K-95, specially distilled; see text.

the Allied Chemical Corporation of the USA. MNA is often referred to as NMA in the USA. The unsaturated 'Nadic' ring of MNA (Fig. 6.2b) enhances the beam stability of the cured resin. MNA is a high temperature hardener and is normally used at temperatures above 60°C to form cross-links with epoxy resins. It reacts with permanganates and retains this capacity after curing. Consequently MNA interferes with permanganate staining of ultrathin sections, and it should not be included in embedding media for specimens fixed with permanganates (Sect. 2.8.4). Nonenylsuccinic anhydride (NSA) (Fig. 6.2c), n-hexenylsuccinic anhydride (HXSA) (Fig. 6.2d) and n-octenylsucdnic anhydride (OSA) (Fig. 6.2e) are low viscosity hardeners and are mainly used as components of the low viscosity epoxy resin embedding media. The formulation for NSA had to be changed by the manufacturers in January 1994 to meet USA government regulations. The 'new' NSA is named K-95-specially distilled (see Table 6.2). It has very similar properties to the original NSA and no changes to the formulations for embedding media are required.

Chapter 6: Embeddinginepoxyresins

6.1.3

Accelerators for epoxy resins

The accelerators used with epoxy resins are tertiary amines and pre­ cautions have to be taken during curing, since their vapours are highly toxic when they are hot. 2,4,6-Tris(dimethylaminomethyl)phenol (DMP-30) and benzyldimethylamine (BDMA) (Table 6.3) are the accelerators in Araldite, Έροη' and similar embedding media. DMP-30 (Fig. 6.3a) was included in the original Araldite embedding medium (Glauert and Glauert 1958), but it has a higher viscosity than BDMA and so there are difficulties in mixing it evenly with the other components of the embedding medium and it diffuses into tissues more slowly (Causton 1980). In addition, DMP-30 is hygroscopic and is readily inactivated unless it is stored carefully to keep it dry. Consequently DMP-30 has been the source of many problems of inadequate embedding through the years (Glauert 1987). BDMA (Fig. 6.3b) has none of these disadvantages and is just as effective as DMP-30 as an accelerator. To compensate for its lower activity, however, OH

(a)

-N(CH,

(CH3)2N-

N(CH3)2

CH, (C)

HO-

-CH,

-CH0 CH,

Fig. 6.3

The structural formulae of accelerators for epoxy resins.

(a) 2,4,6-7Vz's(dimethylaminomethyl)phenol (DMP-30). (b) Benzyldimethylamine (BDMA). (c) Dimethylaminoethanol (DMAE).

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Table 6.3 Accelerators for epoxy resin embedding media

Benzvldimcthvlamine

BDMA

Dimethvlaminoethanol

DMAE

2,4,6-7"ns(dimethylaminomethvl) phenol

DMP-30

Viscosity

Density

cP at 25°C

g/ml

0.85 3.32 20.50

0.93 0.90 0.97-1.00

cP, centipoise. Trade names: BDMA, DY 062 (Ciba-Geigy); DMAE, S-I (Pennsalt) DMP-30, HY 960 and DY 064 (Ciba-Geigy).

its concentration should be increased from the 1.5 to 2.0% usually required with DMP-30 to the higher end of the range 2.5 to 3.0%, but not above 3.0%. The concentration of the accelerator is generally critical in epoxy resin embedding media, the optimum depending upon the resin and anhydride used and the curing schedule. Unfortunately DMP-30 was chosen by Luft (1961) for his popular embedding medium based on Epon 812 and it has been widely used in commercial embedding kits. Some suppliers are now recommending BDMA as a substitute for DMP-30 and it is hoped that other suppliers will follow their example. Dimethylaminoethanol (DMAE) (Table 6.3 and Fig. 6.3c) was intro­ duced by Spurr (1969) as an accelerator for low viscosity embedding media and it has been widely used. Recent studies, however, suggest that BDMA is a more suitable accelerator for these embedding media also. 6.1.4

Additives for epoxy resin embedding media

Various modifiers are added to epoxy resin embedding media to reduce the viscosity of the medium and to improve the sectioning properties of the final block.

Chapter 6:

6.1.4a

Embeddinginepoxyresins

Plasticizers

Dibutyl phthalate (DBP) is the only plasticizer widely used in epoxy resin embedding media. It fulfils two functions: to reduce the viscosity of the embedding medium, thus aiding infiltration into the specimen, and to improve sectioning of the final block. Epoxy resin embedding media containing DBP have ideal sectioning properties and are easier to cut into ultrathin sections than any other embedding medium for electron micro­ scopy. DBP is already present as a component of some commercially available Araldite epoxy resins. Additional DBP was included in the original Araldite M (CY 212) formulation (Glauert et al. 1956), but it was soon found that no additional DBP is required with this resin, although it may be beneficial with other Araldites, such as MY 753 and GY 502, to give them the same excellent sectioning properties as CY 212. There is suggestive evidence that some phthalic esters are mutagenic in mice. Consequently persistent exposure to the fumes of DBP should be avoided. 6.1.4b

Reactiveflexibilizers

The reactive flexibilizers (which are also known as reactive diluents) (Table 6.4) have advantages over plasticizers since they react with the other components of the embedding medium and become an integral part of the cross-linked polymer. Ideally they should react with the anhydride hardener at the same rate as the epoxy resin. These flexibilizers are very efficient in reducing the viscosity of Araldites and other DGEBA resins, but do not appear to give the final block the same excellent sectioning properties as DBP. Although their primary purpose is to reduce viscosity, they can also act as curing agents. Some of the flexibilizers used in early formulations, such as Cardolite NC-513 and Thiokol LP-8, are not recommended, since they are less effective than those listed in Table 6.4. DER 732 (Dow) is an aliphatic polyglycol diepoxide flexibilizer (Fig. 6.4a), with a viscosity of 55 to 100 cP at 25°C, and is pale yellow in colour. It is also marketed as Epi-Rez 502. DER 732 has been included in formulations for embedding media based on Maraglas 655 (Erlandson 1964), DER 332 (Lockwood 1964) and Epon 812 (Kushida 1966b), but has been replaced by DER 736 for most purposes. DER 736 (Dow) is a diglycidyl ether of polypropylene glycol (Fig. 6.4a) with a shorter chain length than DER 732 and a consequent lower viscosity of 30 to 60 cP at 25°C. DER 736 is most widely known as a component of Spurr s low viscosity epoxy resin embedding medium and

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BiologicalspecimenpreparationforTEM

its modifications, but has also been used in an Epon 812 formulation (Kushida 1967). The same flexibilizer is available from Fullam as Effapoxy in a low viscosity kit (No. 51400) in which it is the only epoxide component in an embedding medium which also contains the anhydride hardeners DDSA and MNA. 1,4-BHtanediol diglycidyl ether (BDE) (Fig. 6.4b) is best known by the Ciba-Geigy trade name Araldite RD-2. It is a reactive flexibilizer with an even lower viscosity (19 cP at 25°C) than DER 736 (Table 6.4) and is a component of the Ladd (RD-2) ultra-low viscosity medium developed by John P. Gray and Allen Lovey at Ladd Research Industries in 1974. A purer form of BDE, the reactive diluent Araldite DY 026, is manufactured by Ciba-Geigy at Basle and has the very low viscosity of 10 cP at 25°C. Another 'especially pure' form of BDE is marketed by Ladd as Aquembed and is described as a direct replacement for the water-miscible aliphatic epoxy resin Durcupan (Fluka). BDE is also present in EMbed 812, which was developed by Electron Microscopy

Table 6.4 Reactive flexibilizers for epoxy resin embedding media

Trade name

Viscosity

WPE

cP at 25°C

Density g/ml

DER 732

Diglycidyl ether of a polypropylene glycol

55-100

305-335

1.07

DER 736

Diglycidyl ether of a polypropylene glycol

30-60

175-205

1.00

Araldite RD-2

1,4-Butanediol diglycidyl ether (BDE)

19

136

1.05

Araldite DY 026

1,4-Butanediol diglycidyl ether (BDE)

10-15

110-115

1.05

' Araldite DY 026 is a purer form of RD-2. cP, centipoise; WPE, weight per epoxide. Suppliers: Dow Ciba-Geigy

DER 732 and 736 Araldite RD-2 and DY026.

Chapter 6:

E m b e d d i n g i n e p o x y resins

Sciences as a replacement for Epon 812, although its major component (60%) is DGEBA. η-Butyl glycidyl ether (BGE) (Fig. 6.4c) is a reactive flexibilizer with the relatively high viscosity of 116 cP at 25°C. It is already present in the DGEBA resins Araldite 506 (Ciba-Geigy), Epon 815 (Shell), DER 334 (Dow) and, possibly also, Maraglas 655 (Marblette). All reactive flexibilizers with low molecular weights, such as BDE and BGE, are toxic and are more hazardous than the liquid epoxy resins, except for VCHD (see Table 6.1). BGE is particularly unpleasant and is a possible carcinogen. It must be handled with extreme care and excessive exposure to it must be avoided. Consequently, BGE cannot be recom­ mended for general use and it certainly should not be used as an intermediate solvent (as proposed by Kushida 1963). It is therefore preferable to add BDE as a reactive flexibilizer when it is necessary to lower the viscosity of an epoxy resin embedding medium.

CH,

/°\ (a)

CHo

(b)

CH2

CH

(C)

CH3

CH2

-CH

-CH

CH,-

CH,-

/ CH

-CH,-

/°\

0

\ CH,

/0X CH2

O

(CH2)4

0

CH2

CH2

CH

CH

CH2

κ °\

Fig. 6.4

CH2

CH2

O

CH2

The structural formula of reactive flexibilizers for epoxy resins.

(a) Diglycidyl ether of polypropylene glycol: DER 732, n=9; DER 736, n=4. (b) 1,4-Butanediol diglycidyl ether (BDE)(Araldite RD-2). (c) η-Butyl glycidyl ether (BGE).

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6.1.5

BiologicalspecimeripreparationforTEM

The anhydride:epoxide ratio

For normal industrial use epoxy resin mixtures are formulated so that there is an equivalence of anhydride groups (in the hardener) and epoxide groups (in the epoxy resin), and the curing conditions are selected so that the addition reaction goes to completion. The ratio of anhydride:epoxide (A:E) is consequently 1.0:1.0. It is determined by the formula:

A:E

=

Weight of anhydride in grams Weight of epoxy resin in grams ; : ; MW of the anhydride WPE of the epoxy resin

where MW is the molecular weight of the anhydride and WPE is the weight per epoxide of the epoxy resin, or the number of grams of resin which contain 1.0 gram equivalent of epoxide. When the components of the embedding medium are measured by volume, this formula becomes:

A:E =

Volume of anhydride in ml X density in g/ml

Volume of epoxy resin in ml X density in g/ml

MW of anhydride

WPE of epoxy resin

In the original Araldite formulation of Glauert and Glauert (1958) an A:E of 1.0:1.0 was selected and was obtained with approximately equal volumes of Araldite M (CY 212) and DDSA. There was a departure from industrial practice, however, since the curing temperature was set at 60°C or below and so was far too low for all the possible cross-links between the resin and the hardener to be formed. This turned out to be a blessing in disguise, since it is now known that a fully cross-linked polymer would be too brittle to section. Subsequently, Luft (1961) suggested that the amount of cross-linking should be limited by reducing the A:E ratio and proposed a ratio of 0.7:1.0 (or 0.7) for embedding media based on Araldite GY 502 or Epon 812. At one time it was thought that the A:E ratio had to be adjusted with considerable accuracy (see discussion in Glauert 1991), but it is now clear that the actual value of this ratio is not critical, so long as the curing temperature of 60°C is not exceeded. The degree of cross-linking rises only slowly as the A:E ratio is increased. At higher temperatures the

Chapter 6:

Embeddinginepoxyresins

formation of cross-links progresses much faster and a lower ratio is required to obtain satisfactory blocks. An analysis of the A:E ratios actually used and the results of tests in the laboratory suggest that higher values of 0.8 to 0.9 should be selected for the aromatic Araldites and the Araldite/Έροη' formulations than for the aliphatic 'Epons', where the range 0.70 to 0.75 appears to be more suitable. It cannot be emphasized too strongly that these values are only guidelines and there is no need to adhere to them rigorously. 6.1.6

Curing temperatures for epoxy resins

A temperature of 60°C is recommended for the curing of epoxy resin embedding media for electron microscopy. The formation of cross-links then reaches a maximum in 16 to 24 hours. A linear polymer with few cross-links is favoured at lower temperatures of curing and consequently Luft (1961) suggested a three-stage curing process with incubation at 35°C, and then at 45°C, and finally at 60°C, to improve the sectioning properties of the final block. Similar blocks are obtained, however, when epoxy resins are cured in one stage at 60°C, and so the more lengthy three-stage process is unnecessary, although it may have some value when embedding temperature-labile specimens. Higher curing temperatures produce a more rapid cure and a greater degree of cross-linking, with a consequent modification of the final polymer and its sectioning properties (Causton 1980). At temperatures of 80°C and above the addition reaction in Epon 812 embedding media becomes exothermic and at a curing temperature of IOO 0 C temperatures of over 160°C are reached (D.P. Kiefer, unpublished observations, 1986). Thus high curing temperatures should be avoided when the resin is to be sectioned for electron microscopy. On rare occasions, such as in the assessment of some pathological specimens, a very rapid procedure may be essential and then higher temperatures must be used. It is advisable, however, to start the cure at 70°C and only to raise the temperature, say to 95°C, after 45 min. This avoids any exothermic reaction and mini­ mizes the possible solvent action of the embedding medium at high tem­ peratures, producing blocks with better sectioning properties for both light and electron microscopy.

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6.2

Biological specimen preparation for TEM

Epoxy resin embedding media

Epoxy resins have become the basis for the standard embedding media for electron microscopy, since they are easy to handle and are capable of preserving ultrastructural detail down to the macromolecular level, as in the classic study of muscle fibres by Reedy and Reedy (1985) illustrated in Fig. 2.12 in Sect. 2.8.2. 6.2.1

The development of epoxy resin embedding media

The first epoxy resin embedding medium for electron microscopy (Table 6.5) was based on a Shell epoxy resin (designated EPO) and an aliphatic polyamine hardener (Maaloe and Birch-Andersen 1956). The mixture had a limited compatibility with ethanol, was very viscous and set rapidly at room temperature. Consequently it was very difficult to get adequate infiltration of the embedding medium into the specimen. Sub­ sequently, Glauert et al. (1956) tested a series of Ciba epoxy resins (Araldites) and proposed an embedding medium based on Araldite M and the hardener, DDSA, which has none of the disadvantages of the EPO mixture. The final blocks have excellent sectioning properties and the aromatic groups in the resin give the sections great stability under the electron beam (Sect. 5.1.8). Consequently, embedding media based on Araldite M and similar Araldites are still widely used today. Table 6.5 The development of epoxy resin embedding media for electron microscopy

1956

Maaloe and Birch-Andersen describe the first epoxy resin embedding medium

1956

Glauert, Rogers and Glauert introduce Araldite embedding media

1960

Finck describes the first embedding medium based on Epon 812

1961

Luft publishes his classic paper describing embedding media based on Epon 812 and Araldite GY 502

1964

Mollenhauer suggests combining Araldite and Epon 812 in an embedding medium

1969

Spurr introduces a low viscosity embedding medium based on vinylcvclohexene dioxide

1974

Gray and Lovev develop an ultra-low viscosity embedding medium

Chapter 6:

Embedding in epoxy resins

Subsequently Finck (1960) and Luft (1961) investigated the suita­ bility of the Shell aliphatic epoxy resins (Epons, named Epikotes in Europe) for electron microscopy, and the Epon embedding media proposed by Luft (1961) and the mixtures of Araldite and Epon intro­ duced by Mollenhauer (1964) remain popular. In order to obtain an epoxy resin embedding medium with very low viscosity at room temperature it is necessary to use a very different epoxy resin, VCHD (Sect. 6.1.1), and Spurr (1969) developed an em­ bedding medium based on VCHD. The viscosity of this medium was reduced even further in 1974 by John P. Gray and Allen Lovey, who proposed the inclusion of a different hardener and flexibilizer. All the epoxy resins, hardeners, accelerators, plasticizers and flexibilizers mentioned in this chapter are available from general suppliers (see Appendix), as separate items or as part of an embedding kit. 6.2.2 The aromatic Araldite epoxy resins Araldite is the trade name for the epoxy resins manufactured world-wide by Ciba-Geigy, notably at Duxford in the UK, at Basel in Switzerland, and in the USA. A large range of these resins has been developed for industrial use, but only a few of them have been selected as components of embedding media for electron microscopy (Table 6.6).

Table 6.6 Araldite epoxy resins for electron microscopy Aromatic resins based on DGEBA TradeJ

Viscosity

WPE

Density

cP at 25°C

DBP

g/ml

Araldite M (CY 212)

1300-1650

226-242(234)

1.10-1.15 (1.125)

19%

Araldite MY 753

1800-2800

219-235(227)

1.13-1.17(1.15)

17%

Araldite GY 502

2100-3600

222-238(230)

1.13-1.17(1.15)

17%

Araldite GY 6005

7500-9500

182-196(189)

1.17

DGEBA, diglycidyl ether of bisphenol A; WPE, weight per epoxide DBP, content of dibutyl phthalate by weight; cP, centipoise. Average values for the WPE and density are given in brackets.

0%

189

I 90

Glauert and Lewis:

Biological specimen preparation for TEM

Araldite M was the basis of the first epoxy resin embedding medium developed in 1956 by Glauert and her colleagues (Glauert et al. 1956; Glauert and Glauert 1958). It is an aromatic DGEBA resin (Fig. 6.1a in Sect. 6.1.1) with the addition of 24 grams of the plasticizer DBP for every 100 grams of resin, so that the content of DBP is about 19% by weight. This resin was originally manufactured at both Duxford and Basel and after some years the designation was changed to Araldite CY 212. Manu­ facture at Duxford has now ceased, but the same resin is still available from Ciba-Geigy at Basel, again as Araldite M. These changes in name have resulted in some confusion among microscopists and suppliers alike, and this resin will probably continue to appear in catalogues for a time under the designation CY 212. Consequently both names will be used here. There is the added problem that the original Glauert formulation was acquired by Fluka of Basel in Switzerland in the early 1960s and is marketed in kit form under the trade name Durcupan ACM (where ACM stands for Araldite Casting resin M). Unfortunately the data sheets circulated with these kits are very out of date and give misleading information. The majority of suppliers do not seem to be aware that Durcupan ACM is Araldite M under another name. In these circum­ stances it is advisable to purchase Araldite M (CY 212) instead of Durcupan ACM, so as to obtain the resin at a lower price and with more accurate information. Araldite GY 502 is a DGEBA resin which is similar to Araldite M (CY 212) but contains only 17% DBP by weight (Table 6.6). It is manufactured in the USA and was the first 'American' Araldite to be tested (Finck 1960). Subsequently it was introduced by Luft (1961) in his classic paper on 'improvements' in epoxy resin embedding methods. An identical resin, Araldite MY 753, is manufactured at Duxford. Since these resins contain less DBP than Araldite M (CY 212) they are considerably more viscous. Araldite GY 6005 is a pure DGEBA resin containing no DBP and has a very high viscosity (Table 6.6). Heating to 60°C dramatically decreases the viscosity of all these resins. For example, the viscosity of Araldite MY 750, a pure DGEBA resin with no added plasticizer or flexibilizer, decreases from 12,000 to 16,000 cP at 25°C to 220—260 cP at 60°C (Ciba-Geigy data sheet no. R21c). Similar decreases in viscosity have been reported with the aromatic DGEBA resins, DER 330, 331 and 332, manufactured by Dow.

Chapter 6:

6.2.3

Araldite embedding m e d i a

6.2.3a

T h e standard embedding m e d i a

Embedding in epoxy resins

Embedding media based on Araldite have three basic components: the Araldite resin; an anhydride hardener, DDSA (Sect. 6.1.2); and an amine accelerator, BDMA (Sect. 6.1.3). Standard formulations for each of the Araldite resins are listed in Table 6.7. These are based on the properties of the resins, hardeners and accelerators discussed in Sect. 6.1 and differ from many of the formulations published in other textbooks or proposed in data sheets and in technical notes accompanying embedding kits. Fre­ quently it appears that no accurate calculations have been made of the relative amounts of the resin and hardener to be used. For example, the A:E ratios for all the Araldite GY 6005 media in present use are too low, being 0.5 to 0.6. As a result these media continue to harden after curing for 24 hours at 60°C, since cross-links take longer to establish.

Table 6.7 Standard Araldite embedding media Araldite

Araldite

Araldite

M (CY212)

GY 502

GY 6005

or MY 753 Araldite M (CY 212)

20.0 ml (22.5 g) 20.0 ml (23.0 g)

Araldite GY 502 or MY 753

20.0 ml (23.5 g)

Araldite GY 6005 Hardener, DDSA

22.0 ml (22.0 g)

Plasticizer, DBP Accelerator, BDMA

1.2 ml (1.3 g)

23.0 ml (23.0 g)

28.0 ml (28.0 g)

0.5 ml (0.6 g)

5.4 ml (5.6 g)

1.3 ml (1.4 g)

1.6 ml (1.7 g)

(2.8 to 3.0%)

The curing time for these Araldite media is 16 hours at 60 C. The amount of DBP added to Araldite GY 502, MY 753 and GY 6005 is that required to make them equivalent to Araldite M (CY 212). DDSA, dodecenylsuccimc anhydride; DBP, dibutyl phthalate BDMA, benzyldimethylamine.

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The calculated A:E ratios for the standard media proposed here vary from 0.85 to 0.86, using the average values for the WPE and density of the Araldite given in Table 6.6, and thus are well within the range (0.8 to 0.9) suggested for Araldite embedding media in Sect. 6.1.5. It is easier and simpler to measure the components of the embedding medium by volume rather than by weight, thus avoiding the need to weigh small amounts of viscous liquids and enabling the epoxy resin and the hardener to be measured while they are warm. The warming of the resin and hardener to reduce their viscosity is an essential step in obtaining complete mixing of these two components. Measurement by weight, how­ ever, appears to be standard practice in some laboratories and so the equivalent amounts are also given in grams in Table 6.7. It must be emphasized that the amounts of the Araldite epoxy resin and the hardener, DDSA, need only be approximate, while the accelerator, BDMA, should be measured accurately and its concentration should not exceed 3%. 6.2.3b

Adjustment o f t h e content o f dibutyl phthalate

The plasticizer DBP is added to Araldite resins to reduce the viscosity of the embedding medium and to improve the sectioning properties of the final block (Sect. 6.1.4a). Araldite M (CY 212) contains 19% DBP by weight and this appears to be about optimal for most biological specimens. Araldite GY 502 and Araldite MY 753 contain only 17% DBP (Table 6.6) and so it is advisable, although not essential, to add a small amount of DBP to bring the concentration in the mixture of resin and plasticizer up to approximately 19%, as suggested in Table 6.7. Araldite GY 6005 is an unplasticized DGEBA resin and provides the opportunity to vary the amount of plasticizer for special purposes. Some workers in materials science use Araldite GY 6005 with no added DBP (Ulan et al. 1990), but for most biological tissues these media will be too viscous and also the blocks will be difficult to cut. The amount of DBP suggested in Table 6.7 will give the medium approximately the same content of DBP as the other standard media and this is the maximum amount of DBP that should be required. 6.2.3c

Adjustment o f t h e hardness o f the final block

In the unlikely event that a softer block is required with these aromatic Araldite resins, this can be obtained by increasing the amount of DBP or

Chapter 6:

E m b e d d i n g i n e p o x y resins

by adding a small amount of a reactive flexibilizer, such as BDE (Araldite RD-2 of Ciba-Geigy, or its purer form Araldite DY 026) or a diglycidyl ether of polypropylene glycol (DER 736 of Dow) (Table 6.4 in Sect. 6.1.4b). In the rare event that a harder block is required, this is best achieved by replacing some of the DDSA with the hardener MNA. Since MNA has a higher anhydride content than DDSA (Table 6.2 in Sect. 6.1.2), each ml of DDSA should be replaced with 0.5 ml of MNA to maintain the same A:E ratio. It may then be necessary to reduce the amount of the accelerator BDMA added to ensure that its concentration does not exceed 3%. 6.2.4

Alternative aromatic epoxy resins

Some DGEBA resins contain a reactive flexibilizer (Sect. 6.1.4b) as an alternative to the plasticizer DBP. The most notable feature of these resins is their very low viscosity in comparison with resins containing DBP. For example, the viscosity of the pure DGEBA resin Araldite GY 6010, which is 12,000 to 16,000 cP at 25°C, is reduced to 500 to 700 cP when the resin contains 11% BGE, as in Araldite GY 506. In comparison, the presence of 17% DBP only reduces the viscosity of GY 6010 into the range 2100 to 3600 cP, as in Araldite GY 502. Reactive flexibilizers may be attractive in producing low viscosity, but sectioning of the resulting blocks may not be quite as easy as for resins containing DBP. 6.2.4a

DGEBA resins containing BGE

In addition to Araldite GY 506 (Ciba-Geigy), BGE is also a component of Epon 815 (Shell) and DER 334 (Dow). Araldite GY 506 is the basis for a Fullam embedding kit, and is also a component of an Araldite/Έροη' medium that is still recommended by Mollenhauer (1993). The other resins have also been included in embedding kits from various suppliers. There is considerable concern, however, about the toxicity and possible carcinogenicity of BGE (Sect. 6.1.4b) and Ciba-Geigy have withdrawn Araldite MY 752, which is the UK equivalent of Araldite GY 506. Similarly, Dow have replaced DER 334 with DER 324, which contains an alternative aliphatic glycidyl ether flexibilizer. In these circumstances, it is recommended that embedding media containing BGE are avoided. BDE is relatively harmless.

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Biological specimen preparation for TEM

DGEBA resins containing BDE

The only proposed embedding medium based on DGEBA containing the flexibilizer BDE (Sect. 6.1.4b) is EMbed 812, which was developed by Electron Microscopy Sciences (EMS) as a replacement for Epon 812. It differs from the other 'Epons', however, in that it does not consist only of aliphatic epoxy resins. In fact, a major component (more than 60%) of EMbed 812 is an aromatic DGEBA resin, which is combined with an aliphatic resin and BDE. It has a viscosity of 140 to 180 cP at 25°C, a WPE of 150, and a density of 1.10 g/ml, and thus has similarities to the aliphatic 'Epon' resins (see Table 6.8), in spite of its content of DGEBA. It gives excellent results when used with the hardeners DDSA

Table 6.8 'Epon' epoxy resins for electron microscopy Aliphatic epoxy resins based on di- and tri-glvcidvl ethers of glycerol

Supplier

Trade name

Viscosity

WPE

cP at 25°C

Density g/ml

Shell

Epon S12

120-210

140-170

1.15-1.22

Agar

Agar 100

150-170

145-160

1.22

Fullam

Epox 812

180

155

1.14

Ladd

LX-112

185

141-149

1.23

Nisshin

Quetol 812

140

141

1.23

Polysciences

Poly/Bed 812

140-175

142-149

1.22

Taab

Taab S12

150-170

145-160

1.22

Ted Pella

Eponate 12

130-170

140-160

1.21-1.25

EMS 1 "

EMbed 812

140-180

150

1.10

cP, centipoise; VC'PE, weight per epoxide. Note: Details have only been included if they have been confirmed directlv bv the supplier. Not all of these products mav still be available. ' Electron Microscopy Sciences (EAlS) developed and market EMbed 812 as a replacement for Epon S12. However, it is not an aliphatic epoxy resin, since it contains 60% aromatic diglvcidvl ether of bisphenol A (DGEBA) (see Sect. 6.2.4b).

Chapter 6:

Embedding in epoxy resins

and MNA in Luft's formulation (see Table 6.9 in Sect. 6.2.6a). In addition, an EMS embedding kit is available which contains both EMbed 812 and Araldite GY 502, with DDSA as the only hardener. The sectioning properties of the final block are improved, because of the presence of DBP, and much better preservation of ultrastructure is obtained than with EMbed 812 alone. We are grateful to Stacie Kirsch of EMS for providing information about EMbed 812 and its use. 6.2.5

The aliphatic 'Epon' epoxy resins

An epoxy resin similar to Araldite M was difficult to obtain in the USA in the late 1950s and there was an apparent variability from one batch of resin to another. Consequently Finck (1960) and Luft (1961) investigated the suitability of the Shell epoxy resin Epon (or Epikote) 812 for em­ bedding media for electron microscopy, and Luft (1961) proposed his formulation which is still popular today. Epon 812 is an aliphatic epoxy resin based on di- and tri-glycidyl ethers of glycerol (Fig. 6.1b in Sect. 6.1.1) and it originally appeared to have the advantage of world-wide uniformity. However, in 1973 Luft reported that there had been a gradual drift upwards of the epoxide equi­ valent over the years, as discussed in Sect. 6.1.5, and wide ranges of values for viscosity, WPE and density have been quoted in various publi­ cations. The variability of the WPE of Epon 812 has led to unnecessary concern about the adjustment of anhydride:epoxide ratios. An even greater problem arose in about 1984 when it seems that Shell ceased manufacture of Epon 812. This led to the development of Epon 812 'replacements' by many suppliers of embedding kits (Table 6.8). Most of these replacement resins, which will all be referred to as 'Epon', claim to have been manufactured to tighter specifications than Epon 812, but the range of values given for the WPE is sometimes as wide as 130 to 170. For the present purpose it will be assumed for simplicity that all the 'Epons' have an average WPE value of 150 and a density of 1.23 g/ml. These aliphatic, 'Epon'-type, resins all have considerably lower viscosities than the aromatic Araldites, but they do not have the same uniformity and a wide range of 120 to 210 cP is given for the viscosity of Epon 812 at 25°C. This range is smaller for the replacement resins (Table 6.8) and, in general, these resins have a lower viscosity than Epon 812.

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Biological specimen preparation for TEM

'Epon'embedding media

For practical purposes it is the viscosity of the complete embedding medium, consisting of resin, hardeners and accelerator, which is of importance. Unfortunately mixtures of 'Epon' and anhydride hardeners begin to increase in viscosity as soon as they are brought together, as a consequence of the interaction between the hydroxyl groups in the resin and the anhydride groups in the hardener. This interaction occurs in the absence of the accelerator and in the cold. For example, it has been found that stock mixtures of Poly/Bed 812, DDSA and MNA have a viscosity of 300 cP immediately after mixing, a value considerably higher than for the resin alone. The addition of the accelerator BDMA, at a concen­ tration of 1.4%, increases the viscosity to 380 cP, and this increase is greater when the accelerator DMP-30 is used. Further increases in visco­ sity occur with time. These observations suggest that the curing of 'Epon' commences at room temperature (25°C) as soon as the resin and hardener are mixed. Markedly lower increases in viscosity with time are observed with embedding media based on the aromatic Araldites (Luft 1973), pre­ sumably as a consequence of the presence of fewer hydroxyl groups in the epoxy resin, but it is always advisable to use freshly prepared em­ bedding media. 6.2.6a

Luft's A and B mix 'Epon' embedding medium

To obtain blocks of sufficient hardness with the aliphatic Έροη' resins it is necessary to use a second anhydride hardener, MNA, in addition to DDSA. The two hardeners have different molecular weights (Table 6.2 in Sect. 6.1.2) and consequently Luft (1961) proposed that two separate mixtures should be prepared as follows: Mixture A Mixture B

Epon 812 62 ml Epon 812 IOOml

DDSA IOOml MNA 89 ml

Both the A and B mixtures had an A:E ratio of 0.7 with the samples of Epon 812 available to Luft in 1961. Some problems have arisen due to the fact that there is 100 ml of DDSA in mixture A and 100 ml of Epon 812 in mixture B, with the result that inaccurate tables have been published in some textbooks (for example, Hayat 1989). Consequently we have

Chapter 6:

E m b e d d i n g i n e p o x y resins

Table 6.9 Standard A and B mix Έροη' embedding media (after Luft 1961)

Mixture A

MixtureB

'Epon'

20 ml (24 g)

DDSA

31 ml (31 g)

Έροη'

20 ml (24 g)

MNA

17 ml (21 g)

Table 6.10 Variations of hardness with the amounts of DDSA and MNA in Έροη' embedding media (after Luft 1961) ml

ml

ml

ml

ml

Mixture A

10

7

5

3

0

Mixture B

0

3

5

7

10

0.3

0.3

0.3

0.3

BDMA (maximum)

soft —

0.3 —>• hard

Curing time is 16 hours at 60°C.

recalculated the amounts of resin and hardeners based on 20 ml of Έροη' in both mixtures (Table 6.9). Assuming average values of 150 for the WPE and 1.23 g/ml for the density of the Έροη', the A:E ratios of both standard mixtures A and B are between 0.71 and 0.74, and thus are within the range (0.70 to 0.75) suggested for Έροη' embedding media (Sect. 6.1.5). Once the mixtures A and B have been prepared, it is a simple matter to combine them in different proportions to obtain blocks of a range of hardness, as shown in Table 6.10. It is advisable to start with the 5:5 ratio of A:B and then to change to 7:3 (for a softer block) or 3:7 for a harder block). The accelerator, BDMA, is added to all the final mixtures of A plus B at a concentration of 2.5 to 3.0%. 6.2.6b

Single-mix 'Epon' embedding media

It is usually more convenient to prepare an Έροη' embedding medium in one step, instead of using the two mixtures A and B. In consequence various single-mix formulations have been proposed in textbooks and in

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Table 6.11 Standard single-mix Έροη' embedding media Soft (7:3)

Medium (5:5)

Έροη'

20 ml (24 g)

20 ml (24 g)

DDSA

22 ml (22 g)

16 ml (16 g)

5 ml

MNA BDMA (2.8 to 3.0%)

Hard (3:7)

20 ml (24 g) 9 ml

(9 g)

(6g)

8 ml (IOg)

12 ml (15 g)

1.4 ml (1.5 g)

1.3 ml (1.5 g)

1.2 ml (1.4 g)

Curing time is 16 hours at 60°C. DDSA, dodecenvlsuccinic anhydride; MNA, methyl Nadic anhydride BDMA, benzvldimethylamine.

the data sheets accompanying embedding kits. The basis for these sug­ gested media is not at all clear, since few of them follow the principles established by Luft (1961) and some of them diverge quite widely from his formulations. Consequently we have calculated a standard set of single-mix embedding media and these are listed in Table 6.11. They correspond to the soft (7:3), medium (5:5) and hard (3:7) embedding media of Luft (1961). Assuming an average WPE for the Έροη' of 150 and an average density of 1.23 g/ml, the calculated A:E ratios of these formulations are in the range 0.70 to 0.74. The accelerator, BDMA, is added to each for­ mulation at a concentration of 2.5 to 3.0%. Satisfactory blocks should be obtained with all the samples of 'Epon' listed in Table 6.8. 6.2.6c

'Epon' embedding media with lower viscosities

The hardener DDSA has the highest viscosity (290 to 295 cP at 25°C) of the anhydrides used in epoxy resin embedding media (Table 6.2 in Sect. 6.1.2) and various modifications have been suggested to obtain an Έροη' embedding medium with a lower viscosity by using a different hardener. The most promising of those at present available was developed by Kushida who reduced the viscosity by replacing both DDSA and MNA with the hardener NSA (viscosity 117 to 120 cP at 25°C), which is well known as a component of Spurr's low viscosity medium, in mixtures based on Epon 812 (Kushida 1971) or Quetol 812 (Kushida et al. 1983). The accelerator DMP-30 should be replaced with BDMA (2.5 to 3.0%) in

Chapter 6 :

Embedding in epoxy resins

these media. They must be freshly prepared, since the viscosity increases rapidly after mixing the components. 6.2.7

Ar a l d i t e / ' E p o n '

embedding media

Mollenhauer (1964) tested a range of embedding media and proposed a formulation containing both Araldite M and Epon 812. This medium has proved to be popular in comparison with Έροη' alone, since sectioning is much easier, presumably as a result of the DBP in Araldite M (Sect. 6.1.4a). The aromatic Araldite M also gives the sections greater stability in the electron beam. There is no evidence, however, to suggest that these Araldite/Έροη' media are preferable to Araldite alone, and they do not appear to have been adopted by Araldite users to the same extent. The inclusion of two different epoxy resins is an added complication, both in practice and in the calculation of A:E ratios. In consequence, some of the Araldite/Έροη' formulations suggested in data sheets for embedding kits have low A:E ratios and are not recommended. A set of standard Araldite/'Epon' embedding media is proposed here (Table 6.12). Assuming average values of 150 for the WPE and 1.23 g/ml for the density of the Έροη', and the average values for the WPE and density of the Araldites shown in Table 6.6, the calculated A:E ratios of these media are in the range 0.84 to 0.85. Table 6.12 Standard Araldite/Έροη' embedding media Araldite

Araldite

M (CY212)

GY 502 or MY 753

Araldite M (CY 212)

12 ml (14 g)

Araldite GY 502 or MY 753

12 ml (14 g)

Έροη'

20 ml (24 g)

20 ml (24 g)

Hardener, DDSA

50 ml (50 g)

50 ml (50 g) 0.3 ml (0.4 g)

Plasticizer, DBP Accelerator, BDMA (2.75 to 2.95%)

2.4 ml (2.6 g)

Curing time is 16 hours at 60°C. DDSA, dodecenylsuccinic anhydride; DBP, dibutyl phthalate BDMA, benzyldimethylamine.

2.4 ml (2.6 g)

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Biological specimen preparation for TEM

Storage and mixing of the components of the media

Araldite and Έροη' epoxy resins and the anhydride hardeners, DDSA and MNA, can be stored indefinitely (certainly for many years) if they are kept in air-tight, securely sealed containers at room temperature, preferably with a 'head' of inert gas. They should not be stored in a refrigerator or deep-freeze cabinet, since this increases the risk that water vapour will condense inside containers which are opened before they reach room temperature. The anhydride NSA readily absorbs water, even from the air, and this hardener, and embedding media containing it, must be kept dry in a desiccator. The accelerator BDMA, and other amines, are readily inactivated by water. They should be stored at room temperature in a firmly stoppered bottle and preferably in a desiccator. Problems can still arise with these amine accelerators during use. Each time a bottle is opened there is a possibility that moisture will be introduced from the atmosphere, particularly when the humidity is high. Consequently the containers should remain open for as short a time as possible, and it is probably prudent to replace BDMA with a fresh sample every 6 months. DMP-30 should be replaced more frequently since it is particularly sensitive to inactivation by water vapour. Complete mixing of epoxy resins and anhydride hardeners is necessary to obtain uniform blocks and is only possible if they are first warmed to reduce their viscosity. At room temperature, no amount of stirring or shaking gives complete mixing, even with a mechanical mixer or stirrer. Luft (1973) graphically described the results of mechanical mixing at room temperature, including the entrainment of bubbles of air and stratification. Unfortunately it seems that weighing and mixing of resins and hardeners at room temperature is a common practice in some laboratories, and certainly there is no mention of warming in many data sheets. The resins and hardeners may appear to be fullv mixed, but this will not be so. It is sensible to follow the practice of polymer chemists and to warm the resin and hardeners to obtain complete mixing, as originally recommended by Glauert and Glauert (1958). Heating to 60°C dramatically decreases the viscosity of epoxy resins, but has no other effect on them. Consequently thev can be heated by placing the containers in an oven as often as required. The warming of resins and hardeners enables embedding media to be prepared easily and

Chapter 6:

Embedding inepoxyresins

quickly without using any special apparatus. The vapours of warm epoxy resins, anhydride hardeners and amine accelerators are irritant and sensitizing (see Table 6.1 in Sect. 6.1) and so mixing must take place in a fume cupboard or in a very well ventilated area of the laboratory. It is important to be familiar with all the safety precautions described in Sect. 5.2 before starting work. 6.2.8a Preparation o f epoxy resin embedding media

The standard procedure is as follows: 1.

Place the containers of epoxy resin(s) and hardener(s), and a glass graduated cylinder and a glass conical flask, in an oven at 60°C and leave for at least 10 min. A longer time does no harm. 2. Pour the required volume of the warm epoxy resin (or resins) into the warm graduated cylinder and then add the required volume of warm hardener (or hardeners). Immediately pour the mixture of resin and hardener into the warm conical flask. Shake the mixture gently by rotating the flask by hand for a few minutes until mixing is complete. A few bubbles may form but these will soon disperse. The mixture will now appear clear and structureless. Do not use a stirring rod, since this is not necessary and may introduce bubbles. 3. Add any required flexibilizer or plasticizer and then add the required amount of the accelerator, BDMA, measured accurately with a graduated pipette or in previously calibrated drops. Continue shaking the flask for a further minute or two. The temperature will be falling steadily and so little curing is initiated. 4. The embedding medium is now ready for use. The whole procedure takes less than 30 min and so the embedding medium can be freshly prepared during the final stages of dehydration of the specimens or whenever it is required. The graduated cylinder and conical flask should be drained immediately after use by inverting them over disposable containers. They can then be used again and no washing up is required. The surplus embedding medium collected in the containers should be left to harden, after which it can be disposed of easily and safely.

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This simple method of preparing epoxy resin embedding media is only possible when the amounts of the components are measured by volume. For those who wish to continue to use the more arduous method of measurement by weight, it is necessary to warm the components again after they have been measured and before they are mixed together, if complete mixing is to be achieved. It is not easy to weigh these viscous liquids when they are at room temperature and the whole procedure takes considerably longer than when measurements are made by volume. 6 . 2 . 8 b Storage o f e p o x y r e s i n e m b e d d i n g m e d i a

Mixtures of aliphatic Έροη' epoxy resins and anhydride hardeners begin to increase in viscosity as soon as they are prepared, and this increase is even faster after the accelerator has been added (Sect. 6.2.6). Storage of mixtures of epoxy resins and hardeners should therefore be avoided, despite the fact that this seems to be common practice in some laboratories. These increases in viscosity occur even in the cold and contamination with water vapour is a danger for mixtures stored in a refrigerator or deep-freeze cabinet. In particular, complete embedding media containing the accelerator should not be stored. It is strongly recommended that epoxy resin embedding media be freshly prepared just before use by the simple procedure described above.

6.2.9

E m b e d d i n g m e t h o d s for e p o x y r e s i n s

6 . 2 . 9 a T h e s t a n d a r d p r o c e d u r e for i n f i l t r a t i o n w i t h e p o x y r e s i n s

Araldite and 'Epon' are directly soluble in ethanol and acetone, but the diffusion process is slow, so that removal of every last trace of the

Table 6.13 A standard infiltration schedule for epoxy resins

Dehydrating agent:propylene oxide

1:1

Propylene oxide Propylene oxide:epoxy embedding medium

10 min 10 min

1:1

1 h or longer

Epoxy embedding medium

overnight

Epoxy embedding medium

2 h or longer

Chapter 6:

E m b e d d i n g i n epoxy resins

dehydrating agent is difficult, and any remaining ethanol (or acetone) has an adverse effect on the properties of the final block (Mollenhauer 1988). This problem is overcome by using the intermediate solvent, propylene oxide, which reacts with the anhydride hardener in the embedding medium, so that small amounts remaining in the tissue do no harm. Care has to be taken, since propylene oxide is highly volatile and flammable and also carcinogenic (see Table 6.1). Following dehydration, the specimens remain in the same vials or tubes during infiltration with the epoxy resin. It is essential to warm the embedding medium briefly just before use, since this greatly decreases its viscosity and ensures good infiltration. A standard infiltration schedule is given in Table 6.13. The procedure is carried out at room temperature in a fume cupboard, taking the safety precautions described in Sect. 5.2, and is as follows: 1. Remove the dehydrating agent (usually ethanol or acetone) with a pipette and place it in a waste bottle. 2. Immediately replace the dehydrating agent with equal volumes of the dehydrating agent and propylene oxide, adding sufficient liquid to ensure that the specimen is well covered. Leave for 10 min. 3. Remove the mixture with a pipette. Do not remove all the mixture, since propylene oxide is very volatile and the specimen may dry out. 4. Immediately replace the mixture with propylene oxide, leave it for 10 min, and then remove most of the propylene oxide. 5. Immediately replace the propylene oxide with a mixture containing equal volumes of propylene oxide and freshly prepared epoxy resin embedding medium. Shake the vial gently to mix the components, place it on a specimen rotator (Sect. 3.4), and leave it for 1 h, or longer. 6. Replace the mixture with epoxy resin embedding medium, which has been warmed in an oven to 60°C to reduce its viscosity. Leave the vial overnight on the rotator with the cap off (if possible) to allow any remaining propylene oxide to evaporate. 7. Replace the embedding medium with fresh warm medium and leave the vial for 2 h, or longer. 8. Transfer the specimens to capsules or embedding moulds filled with fresh warm medium in preparation for curing the epoxy resin, as described in Sect. 5.3.3.

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Place all used dehydrating agents, intermediate solvents, liquid embedding media and mixtures containing them in special waste disposal bottles for disposal (see Sect. 5.2). Leave mixtures of propylene oxide and the embedding medium uncovered in a fume cupboard after use to allow the propylene oxide to evaporate. Place any remaining excess embedding medium and the medium removed during infiltration in small plastic containers and allow it to harden before disposal. Similarly allow any embedding medium remaining in the vials to harden before the vials are discarded at the end of the procedure. No washing up is then required. This standard infiltration schedule is suitable for the majority of specimens and has been used successfully for many years with embedding media based on Araldite M (CY 212), one of the most viscous media. 6.2.9b Modified infiltration procedures

Incomplete infiltration can occur with large specimens and dense tissues and when thick cell walls are present, as in plants. In the final block the specimen will then be less hard than the surrounding embedding medium. Adequate infiltration is achieved by one or more of the following procedures: i.

Making sure that dehydration is complete by increasing the times of soaking in the dehydrating agent and in propylene oxide. ii. Passing the specimens more gradually from propylene oxide to the embedding medium, by using mixtures with ratios of 2:1,1:1 and 1:2. iii. Applying a vacuum during the final stages of infiltration (step 7), as described in Sect. 5.3.1c.

Table 6.14 Infiltration schedule for epoxy resins with acetone as the intermediate solvent (Mollenhauer and Droleskey 1985)

Acetoneiepoxyembeddingmedium

1:1

Acetone:epoxy embedding medium

1:3

2h 2h

Epoxy embedding medium

overnight

Epoxy embedding medium

2 h or longer

Chapter 6:

Embedding in epoxy resins

All dehydrating agents and intermediate solvents dissolve lipids. The extraction of lipids is particularly marked with propylene oxide (Chapter 4) and cannot be completely prevented by fixation. Propylene oxide also dissolves some plastics. Consequently it is sometimes necessary to avoid propylene oxide and to use acetone as the intermediate solvent. The application of a vacuum during the final stages of infiltration by the em­ bedding medium will ensure that all traces of the acetone are removed. A typical infiltration schedule, omitting the use of propylene oxide, is given in Table 6.14. Again adjustments to the ratios of acetone to epoxy resin, and to the times for each step in the procedure, may be required to obtain better infiltration. A modified infiltration procedure is also required when acetonitrile (AN) is selected as a less toxic alternative to propylene oxide (Sect. 4.3.2). AN can also be used as the only dehydrating agent, as suggested by Edwards et al. (1992) (see Sect. 4.4.2). Their schedule for infiltration of EMbed 812 at room temperature is given in Table 6.15. Infiltration of embedding resins can be done in an automatic pro­ cessor, such as the Leica Lynx (Sect. 3.4), instead of by hand, but these units are very expensive and so are only justified when large numbers of samples are processed routinely. In addition, modifications to the standard embedding media and processing schedules proposed in this book may well be required to obtain consistent results.

Table 6.15 Infiltration schedule for epoxy resins after acetonitrile dehydration (Edwards et al. 1992)

50% Acetonitrile in water

15 min

70% Acetonitrile in water

20 min

90% Acetonitrile in water

2 X 10 min

100% Acetonitrile Acetonitrile:epoxy embedding medium Epoxy embedding medium

3 X 20 min 1:1

Ih and 18 h 3 X2h

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BiologicalspecimenpreparationforTEM

Infiltration after dehydration by freeze-substitution

Specimens dehydrated at -80 to -90°C by freeze-substitution, as described in Sect. 4.5, are embedded in an epoxy resin at room temperature when good preservation of ultrastructure is the main aim of the experiment. An example of the excellent results obtained is illustrated in Fig. 6.5. A variety of schedules, with different substituting fluids and fixatives, have been proposed (see review by Nicolas and Bassot 1993). The procedures are long and so it is preferable to use one of the commercial units supplied by Leica and Balzers (see Appendix). The schedule must be adjusted for every new specimen and antigen to be detected, and so those included here are only given to provide guidelines.

Fig. 6.5 Electron micrograph of outer ciliary epithelial cells in the eye of an albino rat. Cryofixation by high-pressure freezing, dehydration by freeze-substitution in acetone containing 2% osmium tetroxide, and embedding in Epon. Note the excellent preservation of cytoplasmic organelles, such as the centrioles. Scale bar = 0.5 μπι. (Reproduced from Eggli and Graber 1994, with permission.)

Chapter 6:

Embeddinginepoxyresins

i.

Humbel and Muller (1986), for various tissues. Freeze-substitution in methanol + 1% OsO 4 + 0.4% uranyl acetate + 3% glutaraldehyde + 3% water, -90°C, 8 h; -60°C, 8 h; -30°C, 8 h; 0°C, 30 min. Anhydrous acetone wash. Infiltration with Araldite/'Epon'. Cured at 60°C. ii. Graham and Beveridge (1990), for bacteria. Freeze-substitution in acetone + 2% OsO 4 + 2% uranyl acetate, -80°C, 72 h. Temperature rise to room temperature. Pure acetone, 6X15 min. Infiltration with Epon at room temperature. Acetone:Epon, 1:1, overnight. Fresh Epon. Cured at 60°C for 36 h. iii. Nicolas and Bassot (1993), for dinoflagellates. Freeze-substitution in acetone+ 5% OsO 4 , -90°C, 3 days. Temperature rise at 10°C/h to -30°C. -30°C, 2 h. Temperature rise to room temperature, 1 h. Pure acetone wash. Infiltration with Epon. iv. Eggli and Graber (1994), for rat ciliary body. Freeze-substitution in acetone + 2% OsO 4 , -90°C, 8 h; -60°C, 8 h; -30°C, 8 h (with constant stirring). Pure acetone, temperature rise slowly from -30°C to +4°C. Infiltration with Epon 812 at room temperature. Cured at 60°C. 6.2.9ά

The curing of epoxy resins

After infiltration by the embedding medium is complete, the specimens are transferred to dry capsules or flat embedding moulds, which are then filled with freshly prepared embedding medium, as described in Sect. 5.3.3. It is easier to dispense the embedding medium into the capsules or embedding moulds if it is first warmed briefly to 60°C. The capsules and moulds are then placed in an embedding oven and the embedding medium is cured by heating at 60°C. The vapours from the hot em­ bedding medium are toxic, irritant and sensitizing (see Table 6.1) and so the embedding oven must be placed in an efficient fume cupboard or vented directly to the outside of the laboratory. The initial application of a vacuum of about half an atmosphere (or 380 mm Hg), as described in Sect. 5.3.1c, can sometimes help infiltration of the embedding medium into large or dense specimens, or into plant cells, but this should only be done when it is absolutely necessary. Blocks of Araldite and 'Epon' can be sectioned after curing over­ night (about 16 hours) at 60°C, but the sectioning of some embedding media improves if a longer time of 24 to 48 hours is used. None of these

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times are critical and even longer times (up to a week) at 60°C will not change the blocks significantly. This curing procedure produces blocks with good sectioning properties (Reid and Beesley 1991) with all the formulations proposed here. The temperature of curing should not be increased above 60°C, except in special rapid embedding schedules (see below), since this will alter the hardening process and may result in uneven or brittle blocks (see discussion in Sect. 6.1.6). The temperature of 60°C used in the standard curing schedule is too high for some immunocytochemical studies, but very long incubation times of many days are required to harden epoxy resins at lower tem­ peratures. The effect of heat on the specimen can be reduced by starting at the lower temperature of 35°C, as in Luft's (1961) original procedure, and then increasing the temperature in steps of 45°C and 60°C to com­ plete the cure. With this schedule the majority of the epoxy resin will have immobilized (but not hardened) before the temperature rises and so the adverse effects of heat on antigens will be minimized. 6.2.9e Rapid infiltration and curing o f epoxy resins When it is necessary to obtain quick results, such as in some pathological investigations, the whole procedure from fixation to embedding is performed as rapidly as possible. The specimen should have a maximum thickness of 0.1 mm in at least one dimension and constant agitation is essential during infiltration of the embedding medium. In a typical rapid procedure the total fixation time is reduced to 1 hour or less and dehydration to 30 min or less (Sect. 10.5.1). The infiltration of the embedding medium into the specimen and the subsequent curing of the resin are the most time-consuming steps in the procedure, but the total infiltration time can be reduced from 20 hours or more to less than 1 hour using propylene oxide as the intermediate solvent in the schedule given in Table 6.16, which is modified from Bencosme and Tsutsumi (1970). Table 6.16 Rapid infiltration schedule for epoxy resins

Propylene oxide

2X5min

Propylene oxiderepoxv embedding medium

1:1

5-15 min

Propylene oxide:epoxy embedding medium

1:3

15-20 min

Epoxy embedding medium

10 min

Chapter 6:

Embedding in epoxy resins

Schedules for the rapid infiltration of epoxy resins using microwave irradiation have also been proposed (for example, see Giberson et al. 1995), but little time is saved, as compared with the schedule given here. In addition, it is inadvisable to use flammable solvents in a microwave oven. These solvents should not be used at concentrations above 50%, since they may ignite and the safety risk in using them is just not worth the potential saving of time. Araldite and Epon resins can only be cured rapidly by using high temperatures. The preferred procedure is to place the capsules or embedding moulds containing the specimens in the embedding medium at 70°C for 45 min and then at 95°C for a further 45 min. The embedding oven must be placed in an efficient fume cupboard when these high tem­ peratures are used. Specimens should not be placed directly in an oven at a high temperature of 95 to 100°C, since this procedure is more likely to result in uneven and brittle blocks (Sect. 6.1.6). Rapid curing of epoxy resins can also be obtained in a microwave oven, as discussed in Sect. 5.3.3d. The main effect of microwaves is to raise the temperature very rapidly, so that curing of Epon embedding media occurs within as little as 15 min. Embedding media containing the hardener MNA discolour during curing by microwave irradiation, and so it is advisable to use an Araldite resin, which only requires the hardener DDSA, in place of Epon. As with fixation (Sect. 3.11), microwave irradia­ tion is obviously of value when rapid processing is paramount, although the time taken to calibrate the oven for each embedding has to be taken into account. Initial experiments have given some promising results, but the best methods for carrying out this relatively new procedure require further study and the advantages of microwave irradiation for routine ultrastructural studies have yet to be firmly established.

6.3

Low viscosity epoxy resin embedding m e d i a

In order to obtain an epoxy resin embedding medium of very low viscosity it is necessary to select vinylcyclohexene dioxide (VCHD), the least viscous of the epoxy resins used in electron microscopy, or one of the Quetol media based on ethylene glycol diglycidyl ether. In addition,

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flexibilizers and hardeners must be chosen carefully to achieve a complete embedding medium of low viscosity. Spurr's embedding medium

6.3.1

The first epoxy resin embedding medium with a very low viscosity (60 cP at 25°C) was developed by Spurr in 1969 and is still popular today. It is based on VCHD (ERL-4206 of Union Carbide; Araldite RD-4 of CibaGeigv), which has a viscosity of 7.8 cP at 25°C. This epoxy resin is easy to distil and has good reproducibility, but unfortunately it is a known carcinogen and has a very much higher toxicity than the other epoxy resins used in electron microscopy (see Table 6.1 in Sect. 6.1). Consequently all solutions containing VCHD must be handled with great care in an efficient fume cupboard, using special gloves, as discussed in Sect. 5.2. Table 6.17 Spurr's low viscosity epoxy resin embedding medium Standard

Modifications

A

B

(in grams)

Firm

Hard

VCHD

10.0 6.0

Component

DER 736 NSA DMAE(S-I)

C

D

E

Soft

Rapid cure

Slow cure

10.0

10.0

10.0

10.0

4.0

8.0

6.0

6.0

26.0

26.0

26.0

26.0

26.0

0.4

0.4

0.4

1.0

0.2

0.7

0.7

0.7

1.8

0.3

8.0

8.0

3.0

16.0

or BDMA-·

Cunng time (h) at 60 c Ct at 70 C C

;

16-24 8.0

BDMA is preferable to DM AE.

+Curing at 60°C is preferable to 70°C for the standard medium (see text). BDMA, benzyldimethylamine; DM AE, dimethylaminoethanol DER 736, diglycidyl ether of a polypropylene glycol NSA, nonenvlsuecinic anhydride; VCHD, vinvlcvclohexene dioxide.

Chapter 6:

E m b e d d i n g i n e p o x y resins

VCHD is a cycloaliphatic diepoxide (Fig. 6.1c in Sect. 6.1.1) with a molecular weight of 140, an epoxide equivalent (WPE) of 74 to 78 and a density of 1.10 g/ml. It has a compact structure and yields highly crosslinked polymers with good high-temperature resistance. On its own it produces very hard blocks and so a reactive flexibilizer is added to the embedding medium. Spurr (1969) selected as the flexibilizer a diglycidyl ether of polypropylene glycol (DER 736 of Dow) (Sect. 6.1.4b), which has a moderately low viscosity of 30 to 60 cP at 25°C, with NSA as the hardener. NSA has a lower viscosity than the anhydrides DDSA and MNA which are components of the Araldite and Έροη' embedding media (Table 6.2 in Sect. 6.1.2). Spurr (1969) added the accelerator DMAE (S-l of Pennsalt) to the medium at a concentration of about 1%, and the block then hardens in about 8 hours at 70°C or in 16 to 24 hours at 60°C. A faster or slower cure is obtained by increasing or decreasing the concentration of the accelerator (Table 6.17). More recent studies have shown that BDMA is preferable to DMAE as the accelerator. It has a lower viscosity (see Table 6.3 in Sect. 6.1.3) and is also less volatile, so that it is much more suitable for vacuum infiltration (Sect. 5.3.1c). BDMA is therefore recommended for Spurr's medium and for the other low viscosity epoxy resin embedding media. The hardness of the final block is modified by adjusting the relative amounts of VCHD and DER 736. The A:E ratio of the standard medium is 0.71. This increases to 0.76 or decreases to 0.67 as the amount of DER 736 is varied from 4.0 to 8.0 g in the hard (B) and soft (C) modifications (Table 6.17), but Spurr (1969) reported that it was not necessary to adjust the A:E ratio to be exactly the same as the 0.71 of the 'standard' medium. The speed of the cure can be modified by increasing or decreasing the amount of the accelerator added. With the higher concentrations the viscosity of the embedding medium increases more rapidly at room temperature and so care has to be taken to obtain good infiltration. 6.3.1 a

Preparation o f Spurr's embedding m e d i u m

The anhydride hardener NSA, and to a lesser extent VCHD and DER 736, can absorb moisture from the air and the resultant blocks are then brittle. Consequently the components of the embedding medium should be stored either in a vacuum desiccator or over dry, heat-treated molecular sieve (BDH type 5A, 1/16 inch pellets, for example). Care must be taken not to disturb the molecular sieve when dispensing the

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components. Molecular sieve should not be used to dry the complete embedding medium, since it appears to take up VCHD preferentially, resulting in an incompletely cured, soft block. The embedding medium is prepared in an efficient fume cupboard by adding each component (measured by weight) in turn to a disposable polypropylene embedding beaker. The same amounts can be used when measurement is by volume, since the densities of most of the components are close to 1.0 g/ml. The VCHD, DER 736 and NSA have a low viscosity and can be easily mixed together at room temperature with gentle swirling by hand or with a magnetic stirrer. The accelerator is then added and the complete medium is mixed immediately by brief stirring. The viscosity of the freshly prepared standard medium is 60 cP at 25°C, but this increases significantly within 24 hours, particularly when the medium contains DMAE. Freshly prepared media should always be used. Spurr's medium is compatible with all standard dehydrating and intermediate fluids (Sect. 4.3). With ethanol and acetone it is important to remove all last traces of the dehydrating agent during infiltration of the specimen with the embedding medium. 6.3.1 b Infiltration schedules for Spurr's medium

The mixtures of Spurr's medium and the dehydrating agent are prepared just before use with the same special safety precautions as for the pre­ paration of the embedding medium. Dehydrated specimens in vials or other containers are infiltrated with Spurr's medium in a fume cupboard at room temperature. A typical schedule is given in Table 6.18. It can be varied quite widely, depending on the type of specimen. Some workers find it convenient to infiltrate small specimens during the afternoon and then to cure the resin overnight. The solutions are added and removed from the specimen with fine pipettes and are placed in waste bottles for disposal after use (see Sect. 5.2). Delicate tissues and cells, which are easily affected by changes in the density and viscosity of infiltrating fluids, can be infiltrated by a continuous method in which there is a progressive exchange of ethanol and Spurr's medium, as in the simple procedure of Marchese-Ragona and Johnson (1982) described in Sect. 5.3.1b.

Chapter 6:

Embedding in epoxy resins

Table 6.18 Infiltration schedule for Spurr's medium

Dehydrating agent:Spurr's medium

1:1

30 min-2 h

Dehydrating agent:Spurr's medium

1:3

30 min-2 h

Spurr's medium (for small specimens) or Spurr's medium (for large specimens)

4-6 h 4-6 h, followed by overnight

Specimens which have been dehydrated by freeze-substitution (Sect. 4.5) are infiltrated with Spurr's medium at room temperature by the standard method. The specimens are used primarily for ultrastructural studies and so fixatives are added to the substituting fluid, as for specimens embedded in Araldite or Epon (Sect. 6.2.9c). A typical schedule (Edelmann 1989) is as follows: Freeze-substitution in acetone + 2.5% osmium tetroxide + 0.2% uranyl acetate, -80°C, 2 days. Temperature rise at 3°C/h to -50°C. -50°C, 8 days. Temperature rise at 5°C/h to room temperature. Acetone rinse. Infiltration with Spurr's medium at room temperature. Alternatively the specimens are infiltrated with Spurr's medium at low temperatures. The method described by Hippe-Sanwald (1993) is as follows: Freeze-substitution in methanol+1% osmium tetroxide + 0.5% uranyl acetate + 3% glutaraldehyde + 3% water, -90°C, 8 h; -60°C, 8 h; -35°C, 8 h. Infiltration with VCHD as an intermediate solvent: at -35°C, methanol:VCHD (1:1), 4 h; VCHD, 16 h. At -15°C, VCHD, 4 h; VCHD:Spurr's medium (1:1), 5 h. At +4°C, VCHD:Spurr's medium (1:3), 16 h; Spurr's medium, 24 h.

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6.3.1 c

Biological specimen preparation for T E M

Embedding procedure for Spurr's medium

Following infiltration, the specimens are placed in dry capsules or embedding moulds containing the embedding medium (this can be from the same batch as used for infiltration). Capsules containing very large or dense specimens or tissues dehydrated by freeze-drying may benefit from being placed under a vacuum of about half an atmosphere (or 380 mmHg) for a short period to aid penetration of the embedding medium, but then BDMA should be selected as the accelerator. It is important to place the caps on the capsules and to cover the moulds before putting them in the embedding oven to avoid loss of volatile components from the embedding medium and also to prevent absorption of moisture from the air and the consequent production of blocks which are brittle or have a sticky, uncured surface layer. The whole procedure must be carried out with great care in a fume cupboard and with the use of protective clothing, because of the dangers of exposure to warm VCHD. Spurr's standard medium cures in 16 to 24 hours, when heated at 60°C, but longer times can be used if that is more convenient, with little effect on the characteristics of the final block. The use of an oven temperature of 60°C is preferable to heating at 70°C for 8 hours, as originally proposed by Spurr (1969), since the lower curing temperature results in more repro­ ducible blocks. Shorter or longer times are recommended for modified versions of the standard medium (see Table 6.17). Temperatures higher than 70°C accelerate curing, but tissue damage may then occur and there may also be an increase in the shrinkage of the resin during curing. Shrinkage is usually slight at temperatures of 70°C or lower. The blocks of media containing DMAE are white to pale yellow in colour, the colour increasing as the concentration of DMAE is raised, while with BDMA the colour is darker, ranging from pale yellow to light amber. The blocks are stable during storage and the removal of gelatin capsules by soaking in tap water does not make the resin tacky or notice­ ably change its physical properties (Spurr 1969). Aldrich and Mollenhauer (1986) observed that there was a move­ ment of Spurr's resin relative to the tissue at the surfaces of blocks which had been trimmed and then stored for 6 months. This suggests that sections should be examined as soon as possible after they have been cut (which is good general practice), especially when making studies at high magnification.

Chapter 6:

6.3.2

Embedding in epoxy resins

Ultra-low viscosity epoxy resin embedding media

The viscosity of Spurr's medium was reduced still further in an UltraLow Viscosity (ULV) embedding medium developed by John P. Gray and Allen Lovey at Ladd Research Industries in 1974. This formulation, which was described by Mascorro et al. (1976) and is named Ladd (RD-2), contains the hardener HXSA (Section 6.1.2) and the flexibilizer BDE (Araldite RD-2)(Sect. 6.1.4b), and has a viscosity of 20 cP at 25°C. The hardness of the final block is adjusted by varying the amount of Araldite RD-2. This medium can be improved by replacing RD-2 with DER 736 as the flexibilizer, in a formulation named Ladd (DER). A further modification became necessary recently since the hardener HXSA is no longer available. It has been replaced with OSA (Sect. 6.1.2). This increases the viscosity of the complete medium, but it is still less than 30 cP. The recommended medium (which differs from those available at present in embedding kits) contains VCHD (10.0 g), DER 736 (1.5 g) and OSA (21.0 g), with BDMA (0.7 g) as the accelerator. The curing time is about 24 hours at 60°C, and the medium is handled with great care in the same way as Spurr's medium. 6.3.3

Quetol 651 epoxy resin embedding medium

Hiroshi Kushida has developed a series of embedding media based on the Quetol resins manufactured by the Nisshin Company of Tokyo. The earliest of these media contains the epoxy resin Quetol 651 (Kushida 1974), which is an ethylene glycol diglycidyl ether with the theoretical structure shown in Fig. 6.6. Quetol 651 has a low viscosity of 15 cP at 25°C, an epoxide equivalent of 115 and a density of 1.15 g/ml. It is miscible with ethanol and acetone, and also with water, so that it could presumably be used as a dehydrating agent in a similar way to Durcupan, but this does not appear to have been tried.

/°\ CH9 CH-CH2 Fig. 6.6

0

CH2

CH2

0

CH2

The structural formula of the epoxy resin Quetol 651.

CH

C

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Chapter 6 :

Embedding i n epoxy resins

The embedding medium consists of Quetol 651 (33 ml), the hardener NSA (67 ml) and the accelerator DMP-30 (1.5 to 2.0%). Replacement of the DMP-30 with BDMA (2.5 to 3.0%) is advisable. The medium is prepared and infiltrated into specimens in the same way as Spurr's medium and has the great advantage that Quetol 651 does not have the carcinogenic properties of VCHD. Kushida (1985) has also proposed a number of modifications to the original Quetol 651 medium in which a second hardener, MNA, is added to increase the hardness of the final block. The version of this medium proposed by Johnson (1981) and recommended by Taab Laboratories has an A:E ratio of 1.03 and is not advised. It is claimed that these modified Quetol 651 media are particularly suitable for comparative observations on semithin sections by light and electron microscopy and for the study of thick sections at 300 to 400 kV, but these claims have not been supported by comparative studies with other embedding media. 6.3.4

Properties of low viscosity epoxy resin embedding media

The contrast in sections of Spurr's medium stained with uranyl acetate and lead citrate is well known to be lower than in sections of Araldite or Έροη', particularly for plant tissues, and comparative studies have shown that the contrast is even lower in sections of the ULV epoxy resins (Oliveira et al. 1983; Mollenhauer and Droleskey 1985). There is a pro­ gressive decrease in contrast in stained sections from Quetol 651, in a for­ mulation with NSA as the hardener, to Spurr to Ladd (DER 736) to Ladd (RD-2) (Fig. 6.7), suggesting that there are differences in the uniformity of infiltration by the various media (see Sect. 5.1.7). Regions of the specimen which are fully infiltrated by all the components of the embedding medium, such as in Spurr's medium, will be only lightly stained. The good staining reported by Mascorro et al. (1976) with Ladd (RD-2) probably resulted from incomplete infiltration of the embedding medium, since they used a very rapid schedule. Thus to improve contrast when using ULV epoxy resins, Ladd (DER 736) should be chosen in

Fig. 6.7 A comparison of the contrast in sections of rat liver embedded in (a) Ladd resin containing Araldite RD-2, (b) Ladd resin containing DER 736, (c) Spurr's resin, and (d) Quetol 651. Scale bar = 0.5 μιη. (Reproduced from Mollenhauer and Droleskey 1985, with permission.)

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preference to Ladd (RD-2). Contrast can also be improved by the addition of 1% 'Epon' (Pelco Eponate 12, for example) to the embedding medium. Bozzola and Russell (1992) suggest a formulation consisting of equal parts of Spurr's medium and an Έροη' medium for plant tissues, which is intermediate in viscosity and in staining properties between the two types of epoxy resin. These low viscosity epoxy resin embedding media can cause structural changes in tissues, which are not seen with Araldite and 'Epon'. In general, swelling occurs during infiltration (Sect. 5.1.3) and shrinkage during curing, but these effects are usually slight, except with Ladd (RD-2) where broken membranes have been observed in bacteria (Ringo et al. 1979) and in mammalian tissues (Aldrich and Mollenhauer 1986). Swelling of tissues and tissue components may also occur when the sections are floated on the water surface in the knife trough (Mollenhauer 1988). The water may interact with the tissue in regions of the block in which the resin is incompletely cured. This discussion illustrates the fact that compromises have to be made in the choice of an embedding medium. At present, the best formulation for a low viscosity epoxy resin embedding medium, giving minimal dimensional changes and good contrast after section staining, appears to be the medium which used to be marketed as the Pelco Ultra-Low Viscosity medium, containing VCHD, OSA, DER 736 and BDMA (see Sect. 6.3.2).

6.4

VVater-miscible epoxy resins

Embedding resins that are miscible with water can act as dehydrating agents, thus avoiding the use of ethanol, acetone and propylene oxide (Chapter 4), and so preserving some components of the specimen which might otherwise be extracted. It is important to realize, however, that these resins are organic solvents and it is a fallacy to assume that they will dissolve less lipid. They cause extraction, although the components extracted may be different than with conventional dehydrating agents. Epoxy compounds in aqueous solution react readily with proteins and nucleic acids and therefore tend to act as fixatives (Gibbons 1959). The water-miscible acrylic resins are now more popular than the watermiscible epoxy resins, since they can be used for dehydration and

Chapter 6:

Embedding in epoxy resins

embedding at low temperatures, but a description of the epoxy resins is included here for completeness. 6.4.1

Aquon

The first water-miscible epoxy resin to be tested for electron microscopy was Aquon, the water-miscible fraction of Epon 812 (Gibbons 1959). Aquon did not become commercially available and had to be extracted from Epon 812 in the laboratory by a rather lengthy procedure. Con­ sequently it was soon superseded by a Ciba aliphatic epoxy resin (originally named X133/2097) which was introduced by Staubli (1960) and is marketed by Fluka as Durcupan (see Appendix). Quetol 651 epoxy resin (Sect. 6.3.3) is also miscible with water, but does not appear to have been used as a dehydrating agent. 6.4.2

Durcupan

Durcupan is an aliphatic polyepoxide and is completely miscible with water. The original embedding medium contained Durcupan resin, the anhydride hardener DDSA (Sect. 6.1.2), the plasticizer DBP (Sect. 6.1.4a) and the accelerator DMP-30 (Sect. 6.1.3). The blocks were difficult to section, possibly as a result of the high concentration (5.9 to 7.0%) of DMP-30 used. This embedding medium was modified by Kushida (1964, 1966a) to give blocks with better infiltration and sectioning properties. It is unfortunate that Staubli's original formulation is still recommended by suppliers of embedding kits. It appears that the most suitable of the published Durcupan formulations contains Durcupan (100 ml), MNA (120 ml), Thiokol LP-8 (20 to 35 ml) and DMP-30 (1.5 to 2.0%) (Kushida 1966a). The replacement of the flexibilizer Thiokol LP-8 with DER 736 would probably improve sectioning still further, and it is also advisable to replace 1.5 to 2.0% DMP-30 with 2.5 to 3.0% BDMA as the accelerator. Durcupan is handled in the same way as other low viscosity epoxy resins (Sect. 6.3.1) and the same safety precautions must be taken. 6.4.2a

D e h y d r a t i o n a n d e m b e d d i n g in D u r c u p a n

Specimens are dehydrated in Durcupan resin and are then infiltrated with the complete embedding medium (Durcupan resin plus hardener plus flexibilizer plus accelerator). The dehydration schedule proposed by

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Table 6.19 Dehydration schedule for Durcupan epoxy resin (Staubli 1963)

50% Durcupan resin in water

15-30 min

70% Durcupan resin in water

15-30 min

90% Durcupan resin in water

15-30 min

Durcupan resin

30-60 min

Dureupan resin

30-60 min

Staubli (1963), which is considerably shorter than his original procedure (Staubli 1960), is given in Table 6.19. Specimens are dehydrated with Durcupan at room temperature with constant agitation on a rotary shaker (see Sect. 3.4). 30% Durcupan in water should be avoided as the tissue tends to swell in the same way as it does in 30% aqueous ethanol or acetone. After dehydration the specimens are infiltrated at room temperature with the Durcupan embedding medium, which is not water-miscible, using the schedule given in Table 6.20. They are then transferred to capsules or embedding moulds with fresh Durcupan mixture. The embedding medium is cured by heating at 50°C for about 48 hours. 6.4.2b

Conventional embedding after Durcupan dehydration

An alternative solution to the difficulty of sectioning the original Durcupan medium is to embed the specimens in a conventional waterimmiscible medium after dehydration in Durcupan resin. Embedding is still achieved without passing the specimen through reactive solvents, such as propylene oxide. Specimens dehydrated in Durcupan can be embedded in a polyester or methacrylate resin (Kushida 1965), but it is preferable to use an epoxy resin (Staubli 1963). Durcupan is a low Table 6.20 Infiltration schedule for Durcupan epoxy resin (Kushida 1966a)

Durcupan:Durcupan embedding medium

3:1

Ih

Durcupan:Durcupan embedding medium

1:1

Ih

Durcupan:Durcupan embedding medium

1:3

Ih

Dureupan embedding medium

1-2 h

Dureupan embedding medium

1-2 h

Chapter 6:

Embedding in epoxy resins

viscosity epoxy resin and forms cross-links with other epoxy resins to become part of the final polymer. The specimens are passed from Durcupan resin to the embedding medium through a series of mixtures of increasing concentration of the embedding medium (for example 3:1, 1:1, 1:3 for 1 hour each) and then to 100% embedding medium. Following infiltration into the specimen, the embedding medium is cured in the usual way. 6.4.3

Aquembed

Aquembed was introduced by Ladd (see Appendix) as a substitute for Durcupan (Fluka). It is an especially pure form of the water-miscible aliphatic polyepoxide BDE (Sect. 6.2.4b), with a viscosity of less than 10 cP at 25°C, a WPE of 102 to 105, and a density of 1.07 g/ml. Ladd currently advise using similar formulations, dehydration and embedding procedures for Aquembed as originally proposed by Staubli (1960) for Durcupan. It would be preferable, however, to use the modified embedding medium and procedures recommended by Kushida (Sect. 6.4.2).

References Aldrich, H.C. and Mollenhauer, H.H. (1986) Secrets of successful embedding, sectioning, and imaging. In Ultrastructure Techniques for Microorganisms, Aldrich, H.C. and Todd, W.J. (eds.), pp. 101-132, Plenum Press, New York Bencosme, S.A. and Tsutsumi, V. (1970) A fast method for processing biologic material for electron microscopy. Laboratory Investigation 23,447^50 Bozzola, J.J. and Russell, L.D. (1992) Electron Microscopy. Principles and Techniques for Biologists. Jones and Bartlett, Boston Causton, B.E. (1980) The molecular structure of resins and its effect on the epoxy embedding resins. Proceedings of the Royal Microscopical Society 15, 185-189 Causton, B.E. (1981) Resins: toxicity, hazards and safe handling. Proceedings of the Royal Microscopical Society 16, 265-271 Edelmann, L. (1989) The contracting muscle: a challenge for freeze-substitution and low temperature embedding. Scanning Microscopy Supplement 3, 241—252 Edwards, H.H., Yeh, Y.-Y., Tarnowski, B.I. and Schonbaum, G.B. (1992) Acetonitrile as a substitute for ethanol/propylene oxide in tissue processing for transmission electron microscopy. Microscopy Research and Technique 21, 39-50 Eggli, P.S. and Graber, W. (1994) Improved ultrastructural preservation of rat ciliary body after high pressure freezing and freeze substitution: a perspective view based upon comparison with tissue processed according to a conventional protocol or by osmium tetroxide/microwave fixation. Microscopy Research and Technique 29, 11—21

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Erlandson, R.A. (1964) A new Maraglas, D.E.R. 732, embedment for electron microscopy. Journal of Cell Biology 22, 70+-709 Finck, H. (1960) Epoxy resins in electron microscopy. Journal of Biophysical and Biochemical Cytology 7, 27-30 Gibbons, I.R. (1959) An embedding resin miscible with water for electron microscopy. Nature (London) 184, 375-376 Giberson, R.T., Smith, R.L. and Demaree, R.S. (1995) Three hour microwave tissue processing for transmission electron microscopy: from unfixed tissue to sections. Scanning 17, Suppl. V, 26-27 Glauert, A.M. (1987) Accelerators for epoxy resins. Proceedings of the Royal Microscopical Society 22, 264 only Glauert, A.M. (1991) Epoxy resins: an update on their selection and use. Microscopy and Analysis, September 1991,15-20 Glauert, A.M. and Glauert R.H. (1958) Araldite as an embedding medium for electron microscopy. Journal of Biophysical and Biochemical Cytology 4, 191-194 Glauert, A.M., Rogers, G.E. and Glauert, R.H. (1956) A new embedding medium for electron microscopy. Nature (London) 178, 803 only Graham, L.L and Beveridge, T.J. (1990) Evaluation of freeze-substitution and conventional embedding protocols for routine electron microscopic processing of eubacteria. Journal of Bacteriology 172, 2141-2149 Hayat, M.A. (1989) Principles and Techniques of Electron Microscopy; Biological Applications, 3rd edn. Macmillan, Basingstoke Hippe-Sanwald, S. (1993) Impact of freeze substitution on biological electron microscopy. Microscopy Research and Technique 24,400-422 Humbel, B. and Muller, M. (1986) Freeze substitution and low temperature embedding. In Science of Biological Specimen Preparation, Muller, M., Becker, R.P., Boyde, A. and Wolosewick, J.J. (eds.), pp. 175-183, SEM Inc., AMF O'Hare, Chicago, Illinois Johnson, J.E. (1981) Transmission and scanning electron microscopy. In Current Trends in Morphological Techniques, Johnson, J.E. (ed.), CRC Press, Boca Raton Kushida, H. (1963) An improved epoxy resin 'Epok 533', and polvethylene glycol 200 as a dehydrating agent. Journal of Electron Microscopy 12, 167-174 Kushida, H. (1964) Improved methods for embedding with Durcupan. Journal of Electron Microscopy 13, 139-144 Kushida, H. (1965) Durcupan as a dehydrating agent for embedding with polyester, styrene and methacrylate resins. Journal of Electron Microscopy 14,52-53 Kushida, H. (1966a) Further improved method for embedding with Durcupan. Journal of Electron Microscopy 15, 94-95 Kushida, H. (1966b) New embedding with D.E.R. 732 and Epon 812. Journal of Electron Microscopy 15, 96-98 Kushida, H. (1967) A new embedding medium employing D.E.R. 736 and Epon 812. Journal of Electron Microscopy 16, 278-280 Kushida, H. (1971) A new method for embedding with Epon 812. Journal of Electron Microscopy 20, 206-207 Kushida, H. (1974) A new method of embedding with a low viscosity epoxy resin "Quetol 651".Journal of Electron Microscopy 23, 197 only Kushida, H. (1985) Embedding method for electron microscopy in biology. TokaiJournal of Experimental and Clinical Medicine 10, 557-571 Kushida, H., Kushida, T., Yonehara, K. and Nakazawa, E. (1983) A new embedding medium available for stereoscopic observation of semi-thin sections under 200 kV transmission electron microscope. Journal of Electron Microscopy 32, 61-65

C h a p t e r 6:

E m b e d d i n g i n e p o x y resins

Lockwood, W.R. (1964) A reliable and easily sectioned epoxy resin embedding medium. Anatomical Record 150, 129-140 Luft, J.H. (1961) Improvements in epoxy resin embedding methods. Journal of Biophysical and Biochemical Cytology 9, 409-414 Luft, J.H. (1973) Embedding media - old and new. In Advanced Techniques in Biological Electron Microscopy, Koehler, J.K. (ed.), pp. 1-34, Springer-Verlag, Berlin and Heidelberg Maaloe, O. and Birch-Andersen, A. (1956) On the organization of the 'nuclear material' of Salmonella typhimurium. Symposia of the Society for General Microbiology 6, 261-278 Marchese-Ragona, S.P. and Johnson, S.P.S. (1982) A simple method for the progressive infiltration of resin into a dehydrated biological sample. Proceedings of the Royal Microscopical Society 17, 311-312 Mascorro, J.A., Ladd, M.W. and Yates, R.D. (1976) Rapid infiltration of biological tissues utilizing n-hexenyl succinic anhydride (HXSA)/vinyl cyclohexene dioxide (VCD), an ultra-low viscosity embedding medium. Proceedings of the 34th Annual Meeting of the Electron Microscopy Society of America, 346-347 Mollenhauer, H.H. (1964) Plastic embedding mixtures for use in electron microscopy. Stain Technology 39,111-114 Mollenhauer, H.H. (1988) Artifacts caused by dehydration and epoxy embedding in transmission electron microscopy. In Artifacts in Biological Electron Microscopy, Crang, R.F.E. and Klomparens, K.L. (eds.), pp. 43-64, Plenum Press, New York and London Mollenhauer, H.H. (1993) Artifacts caused by dehydration and epoxy embedding in transmission electron microscopy. Microscopy Research and Technique 26,496-512 Mollenhauer, H.H. and Droleskey, R.E. (1985) Some characteristics of epoxy embedding resins and how they affect contrast, cell organelle size, and block shrinkage. Journal of Electron Microscopy Technique 2, 557-562 Nicolas, M.-T. and Bassot, J.-M. (1993) Freeze substitution after fast-freeze fixation in preparation for immunocytochemistry. Microscopy and Research Technique 24, 474-487 Oliveira, L., Burns, A., Bisalputra, T. and Yang, K.-C. (1983) The use of an ultra-low viscosity medium (VCD/HXSA) in the rapid embedding of plant cells for electron microscopy. Journal of Microscopy 132,195-202 Reedy, M.K. and Reedy, M.C. (1985) Rigor crossbridge structure in tilted single filament layers and flared-X formations from insect flight muscle. Journal of Molecular Biology 185,145-176 Reid, N. and Beesley, J.E. (1991) Sectioning and cryosectioning for electron microscopy. In Practical Methods in Electron Microscopy, Vol. 13, Glauert, A.M. (ed.), Elsevier, Amsterdam Ringo, D.L., Cota-Robles, E.H. and Humphrey, B.J. (1979) Low viscosity embedding resins for transmission electron microscopy. Proceedings of the 37th Annual Meeting of the Electron Microscopy Society of America, 348-349 Spurr, A.R. (1969) A low-viscosity epoxy resin embedding medium for electron microscopy. Journal of Ultrastructure Research 26, 31-43 Staubli, W. (1960) Nouvelle matiere d'inclusion hydrosoluble pour la cytologie electronique. Comptes Rendus des Seances de PAcademie des Sciences 250,1137-1139 Staubli, W. (1963) A new embedding technique for electron microscopy, combining a water-soluble epoxy resin (Durcupan) with water-insoluble Araldite. Journal of Cell Biology 16, 197-201

223

7 Embedding in acrylic resins

7.1

Acrylic resins for electron microscopy

Acrylic resins are transparent, colourless polymers formed from sub­ stituted derivatives of acrylic acid (Fig. 7.1). Acrylic monomers have a very low viscosity and are polymerized by radical chain reactions in which the double bond in the acryl group is the centre of the reaction. Monomers having one acryl group, such as methyl methacrylate, form linear polymers, while polyfunctional monomers, such as ethylene glycol dimethacrylate, form cross-linked, three-dimensional, insoluble structures (Fig. 7.1). Both types of polymer are thermoplastic and are brittle below their glass transition temperature (T g ), which is defined as the temperature at which glassy amorphous polymers become flexible or rubber-like because of the onset of the free rotation of covalent bonds (Gerrits and Horobin 1996). Consequently plasticizers, such as poly­ ethylene glycol, have to be added to some formulations to decrease the T g and thus to improve the sectioning properties of the blocks at room temperature or below (Gerrits et al. 1990). 7.1.1

The properties of acrylic resins

Acrylic monomers retain their low viscosity as the temperature is lowered, and so specimens can be infiltrated and the resin can be poly­ merized at low temperatures. This is their main advantage as embedding media for electron microscopy. The monomers of some acrylic resins, such as 2-hydroxyethyl methacrylate (glycol methacrylate, GMA), 2-hydroxypropyl methacrylate (HPMA), the London resins (LRs) and the Lowicryls, are polar and are miscible with water, and consequently they can be used for partial or complete dehydration (Sect. 4.4.4). Some of these, such as Lowicryl K4M, will polymerize in the presence of considerable amounts of water and the final polymer is hydrophilic.

I

10

Glauert and Lewis:

Fig. 7.1

The

Biological specimen preparation for TEM

structural

formulae

of

acrylic

resins

and

the

mechanism

of

their

polymerization. (a) Acrylic acid. (b) A m e t h a c r y l a t e in w h i c h R is an alkyl g r o u p . (c) A dimethacrylate cross-linker in which t w o methacrylate groups are linked by a dialcohol. (d) T h e m e c h a n i s m b y w h i c h an individual step in t h e p o l y m e r i z a t i o n of an acrylic resin adds a n e w molecule t o t h e m o n o m e r . T h e n e w b o n d f o r m e d is s h o w n elongated. I s ' represents t h e initiator molecule w h i c h p r o d u c e s an u n p a i r e d electron ( O ) at the next c a r b o n a t o m .

In contrast, the polymers of GMA and H P M A are hydrophobic, since most or all of the hydrophilic groups of the monomers appear to be used up during polymerization. Acrylic resin monomers extract lipids and other components from tissues. For example, substantially more lipid is extracted from sciatic

Chapter

7:

Embedding in acrylic resins

nervous tissue fixed with glutaraldehyde and osmium tetroxide during embedding in LR White than in conventional epoxy resins (Kent 1992), and extraction of 14 C-Iabelled compounds from plant tissues is 40-times higher than with Spurr s epoxy resin (Coetzee and Van der Merwe 1989). The lipids in tissues fixed only with aldehydes are particularly vulnerable to extraction during infiltration with water-miscible acrylic resins, which are powerful lipid solvents for neutral lipids and some phospholipids, even at —20°C (see Glauert 1975). During dehydration and infiltration with Lowicryl resins, a temperature of -50°C or lower is required to reduce the extraction of lipids significantly from glutaraldehyde-fixed cells of Acboleplasma laidlawii (Weibull and Christiansson 1986) and to obtain improved preservation of myelin (Kent 1992) 7./.2

T h e polymerization o f acrylic resins

Acrylic resins are polymerized by heat or by UV irradiation, and the process is controlled by the addition of initiators, such as dibenzoyl peroxide, which produce free radicals when they decompose, and accelerators, such as the aromatic tertiary amines, which promote the action of free radical-producing agents by, for example, accelerating the decomposition of dibenzoyl peroxide. The polymerization of acrylic resins is accompanied by considerable shrinkage (10 to 20%, by volume) and impurities in the embedding medium or localized deposits of osmium, or even components of the specimen itself, can initiate polymerization, which may then begin in one region before another. This process is accentuated by the fact that the polymerization reaction is exothermic. The heat generated cannot escape through the surrounding resin and so 'hot spots' develop in which the reaction is accelerated. The resulting uneven polymerization causes stresses within the hardening block, thus damaging the specimen, and the final block has regions of different hardness. Various procedures have been advocated to reduce 'polymerization damage', such as the addition of 0.01% uranyl nitrate to the embedding medium, so that there is a more uniform initiation of polymerization, or the addition of crosslinking agents, such as divinyl benzene or dimethacrylates, which reduce the shrinkage during polymerization.

2Π

228

Glauert and Lewis:

7. 1.3

Biological specimen preparation for TEM

Stability in the electron b e a m

A big disadvantage of acrylic embedding media for ultrastructural studies is that the sections are unstable in the electron beam. As much as 70% of the resin may be lost under normal operating conditions and this loss is followed by a flow of the remaining material and a consequent distortion of structures within the specimen. This damage occurs at beam intensities lower than those normally used for examining specimens and is quite different from the larger scale changes, such as movement of the section and the appearance of holes, which are often quoted as indications of 'instability'· This early loss of mass is reduced when divinyl benzene is present as a cross-linking agent, but this increased resistance is not due to the greater number of cross-links, but to the presence of the aromatic ring in divinyl benzene. Increased beam resistance is also obtained by the incorporation of an aromatic acrylic resin into the embedding medium, as in LR White and LR Gold, but the loss of mass during irradiation is still greater than with the epoxy resins (Sect. 5.1.8). 7. 1 . 4

T h e d e v e l o p m e n t o f acrylic resin e m b e d d i n g m e d i a

The non-polar n-alkyl methacrylate esters, η-butyl methacrylate and methyl methacrylate, were the basis for the first embedding media developed specifically for electron microscopy by Newman et al. (1949) (Table 7.1). They enabled much thinner sections to be cut than is possible with paraffin wax, but the preservation of ultrastructure is poor as a result of extraction of tissue components, uneven polymerization and irra­ diation damage in the electron microscope. These embedding media were superseded by the epoxy resins for ultrastructural studies, following the introduction of Araldite in 1956. The polar acrylic embedding media, such as those based on 2hydroxyethyl methacrylate (glycol methacrylate, GMA) (Rosenberg et al. 1960) and 2-hvdroxypropyl methacrylate (HPMA) (Leduc and Holt 1965), have advantages for enzyme cytochemistry and immunocytochemistry, because of their low viscosity, miscibilitv with water, and ability to polymerize at low temperatures. Formulations based on GMA are still advocated for combined light and electron microscopy. Examples are the GMA-Quetol 523 (a methoxy polyethylene glycol 200 methacrylate) mixture of Kushida (1977) and Bioacryl (or Unicrvl; British BioCell), which is a mixture of

Chapter 7:

Embedding in acrylic resins

Table 7.1 Acrylic resins for electron microscopy.

Components of the earliest acrylic embedding media

Introduced by:

Non-polar resins: Methyl methacrylate

Newman et al. (1949)

w-Butyl methacrylate Polar resins for cytochemistry: 2-Hydroxyethyl methacrylate (GMA)

Rosenberg et al. (1960)

2-Hydroxypropyi methacrylate (HPMA)

Leduc and Holt (1965)

Components of polar Lowicryl embedding media Lowicryl K4M

Carlemalm et al. (1982)

2-Hydroxypropyl methacrylate (HPMA) 2-Hydroxyethyl acrylate (toxic) n-Hexyl methacrylate Triethylene glycol methacrylate (cross-linker) Lowicryl Kl IM

Acetarin et al. (1986)

2-Hydroxypropyl methacrylate (HPMA) 2-Hydroxyethyl acrylate (toxic) Ti-Butyl methacrylate 2-Ethoxyethyl methacrylate 2-Methoxyethyl methacrylate 1,3-Butanediol dimethacrylate (cross-linker)

Components of non-polar Lowicryl embedding media Lowicryl HM 20

Carlemalm et al. (1982)

Ethyl methacrylate «-Hexyl methacrylate Triethylene glycol dimethacrylate (cross-linker) Lowicryl HM 23

Acetarin etal. (1986)

Ethyl methacrylate «-Butyl methacrylate 1,3-Butanediol dimethacrylate (cross-linker)

Component of LR White and LR Gold A polyhydroxy-substituted bisphenol A dimethacrylate (cross-linker)

Causton (1984)

229

230

Glauert and Lewis:

Biological specimen preparation for TEM

GMA (40%), HPMA (30%), «-butyl methacrylate (26%) and styrene (4%) (Scala et al. 1992). It is claimed that Unicryl is 'new' and 'unique', but there is nothing particularly novel about its formulation and it is very expensive compared with a standard GMA embedding medium. In addition, styrene is a carcinogen and so great care must be taken when handling Unicryl. This is particularly important at the high temperatures used during polymerization of the resin, when there is the danger of the production of toxic fumes. None of these media can be recommended for electron microscopy, because of their poor preservation of ultrastructure, but GMA has proved to be excellent for a wide range of cytochemical investigations and a standard procedure for embedding in GMA for light microscopy is described in Sect. 7.2.3. The minimum temperature for infiltration and for polymerization of media based on GMA and HPMA is in the region of —30°C, which is not low enough to produce a significant reduction in the modification and extraction of cell components during embedding. Consequently, Carlemalm et al. (1982) and Acetarin et al. (1986) developed the Lowicryl embedding media, which are complex mixtures of acrylic resins (Table 7.1). Lowicryl K4M is very similar to the standard GMA and HPMA formulations, while Kl 1M, HM20 and HM23 are all capable of use at — 50°C or below. They are popular for studies in immunocytochemistry, although again the preservation of ultrastructure is still no more than adequate. The London resins, LR White and LR Gold, were developed with the specific aim of providing an embedding medium with low toxicity, as well as low viscosity (Causton 1981). The composition of these resins has not been published, but they contain a polyhydroxy-substituted bisphenol A dimethacrylate, which reduces the loss of mass from the sections under irradiation by the electron beam. LR White is a powerful lipid solvent (Sect. 7.1.1) and membranes are poorly preserved. It will, however, poly­ merize at low temperatures and is convenient to use, since it is supplied in a pre-mixed form. Consequently it is the acrylic resin of choice for cytochemical studies by combined light and electron microscopy. The components of acrylic resin embedding media and the embedding kits mentioned in this chapter are available from general suppliers (see Appendix).

Chapter 7:

7.1.5

E m b e d d i n g i n a c r y l i c resins

Embedding methods for acrylic resins

7. 1.5a Safe handling of acrylic resins Acrylic resins are highly volatile, even at low temperatures, and can cause irritation and allergic skin reactions (see Table 7.2). Particular care must be taken with those embedding media, such as Lowicryls K4M and Kl 1M, that contain 2-hydroxyethyl acrylate, which is strongly Table 7.2 Hazards in handling acrylic resin embedding media Component

Examples

Potential hazards

Resins Methacrylate Acrylate

Methyl and «-butyl

Mildly toxic

GMA and HPMA

Toxic

2-Hydroxyethvl acrylate

Highly toxic

Cross-linkers Aliphatic Aromatic

Triethylene glycol dimethacrylate

Low toxicity

1,3-Butanediol dimethacrylate

Toxic

Divinyl benzene

Toxic

Styrene

Carcinogenic

N, N-Dimethy laniline

Highly toxic

7V,N,3,5-Tetramethylaniline

Mildly toxic

N t N-Dimethyl paratoluidine

Highly toxic

PEG 400

Low toxicity

2-Butoxyethanol

Highly toxic

2-Isopropoxyethanol

Low toxicity

Dibenzovl peroxide

Potentially explosive

Benzoin methylether

Toxic

Accelerators Aromatic tertiary

Plasticizers

Initiators

The information about hazards with these components of embedding media is incomplete, and only known hazards are listed here. All the components are, however, irritant and potentially allergenic. GMA, 2-hydroxyethyl mcthacrylate; HPMA 1 2-hydroxypropyl methacrylate PEG, polyethylene glycol.

231

232

GIauertandLewis:

BiologicalspecimenpreparationforTEM

sensitizing (Tobler and Freiburghaus 1990). Contact with the skin and eyes must be avoided and the vapours of the resins must not be inhaled, particularly when the resins are hot. In addition, some of the aromatic tertiary amines, such as 7V,7V-dimethylaniline, which are used as accele­ rators, and some plasticizers, such as 2-butoxyethanol, are highly toxic. Consequently all the safety precautions outlined in Sect. 1.4.5 and Sect. 5.2 must be taken, particularly during infiltration with mixtures of resins and dehydrating solvents when toxic and other effects are greatest. Emphasis must be given to the wearing of protective gloves and to the use of an efficient fume cupboard. 7.1.5 b

Storage and mixing of acrylic resins

Acrylic resins should be stored in firmly closed containers in a wellventilated area of the laboratory. As with all potentially hazardous chemicals, minimal stocks should be kept and fresh components should be used whenever possible. The initiator, dibenzoyl peroxide, which aids the polymerization of acrylic resins, must be handled with particular care. It breaks down spontaneously when it gets too hot and can explode. Consequently it is usually supplied as a powder or in a sus­ pension in water. It must be stored in small bottles in a cool place and well away from other flammable materials. Acrylic embedding media should be prepared and dispensed within a plastic tray on a special area of bench to control spillages, as described in Sect. 5.2, and preferably within a fume cupboard containing the necessary balance and other equipment. Excess resin should be placed in waste bottles with tightly fitting lids for disposal (see Sect. 1.4.5c). 7.1.5c

Infiltration and polymerization of acrylic resins

The procedure for the infiltration of acrylic resins into dehydrated speci­ mens in vials or microtubes is the same as the standard procedure for epoxy resins described in Sect. 5.3.1, although shorter times can usually be used at each stage. Any acetone used during dehydration must be completely removed from the resin and oxygen must be excluded, since they both act as radical scavengers and inhibit the polymerization of many acrylic resins. Gelatin capsules are preferred, since gelatin com­ pletely excludes air. Embedding moulds must be covered, unless the resin is polymerized in an atmosphere of nitrogen gas.

Chapter 7:

Embedding in acrylic resins

The main problem during polymerization of acrylic resins by heat or UV irradiation is the rapid rise in temperature which occurs as a result of the exothermic reaction. It is essential to remove the heat evolved as rapidly as possible to prevent an uncontrolled reaction resulting in large local temperature rises and uneven polymerization. The simplest method of conducting the heat away is to place the specimen containers in a cooled block, which acts as a 'heat sink', but this is only effective when very small volumes of resin are polymerized. More efficient methods of temperature control, such as an ethanol bath, are described in the next chapter in Sect. 8.8. 7.2

Glycol methacrylate for light microscopy

Although GMA has disadvantages for ultrastructural studies by electron microscopy (Sect. 7.1.4), GMA media have proved to be excellent for a wide range of cytochemical investigations by light microscopy, and con­ sequently a standard procedure for embedding in GMA is included here to give guidelines for its proper use. In addition GMA is fully miscible with water and enables specimens to be dehydrated and embedded at low temperatures for studies in immunocytochemistry. This account is based on the extensive and critical studies of Peter Gerrits and Richard Horobin (Gerrits and Horobin 1996). 7.2.1

Glycol methacrylate

In its monomeric form, 2-hydroxyethyl methacrylate (GMA) is the ethylene glycol monoester of methacrylic acid (Fig. 7.2). The remaining hydroxyl group of ethylene glycol is largely responsible for the hydrophilic properties of the GMA monomer. It is a hygroscopic, transparent and colourless liquid with a molecular weight of 130, a density of 1.07 g/ml and a viscosity of 5 cP at 30°C. GMA is a mild skin irritant and may cause allergic skin reactions. Gloves should be worn when handling the monomer. Commercial samples of the GMA monomer contain hydroquinone, hydroquinone monomethylether, or methoxyethyl hydroquinone (in concentrations of 200 to 2000 ppm) to block free radicals and thereby prevent spontaneous polymerization. These inhibitors can be removed, but satisfactory polymerization is achieved without removing them if

233

234

Glauert and Lewis:

Biological specimen preparation for TEM

(a)

HO

CH 2

CH 2

OH

CH 3

(b)

I

CH2^C

C OH

CH 3

(C)

I

CH2^c

C 0

CH 2

CH 2

OH

Fig. 7.2 The structural formulae of (a) ethylene glycol, (b) methacrylic acid, and (c) glycol methacrylate.

suitable amounts of the initiator and accelerator are used. The GMA polymer is slightly hydrophobic. GMA has a high glass transition temperature (T g ) of 55°C and con­ sequently a plasticizer has to be added to decrease this temperature for sectioning at room or lower temperatures (Sect. 7.1). Ideally the blocks should be sectioned at a temperature close to their T g . A plasticizer which is widely used in experimental formulations (Ruddell 1967), as well as in commercial GMA embedding kits (such as JB-4, Polysciences), is 2-butoxyethanol (BE) (Table 7.3). Unfortunately BE is a toxic irritant with a high vapour pressure and an unpleasant smell, so that it must be handled with great care. In addition, these small molecules readily migrate from the inside of a resin block to the surface. Consequently blocks can vary in consistency if they are kept for long periods of time and a noxious and unpleasant odour is released during sectioning. These formulations are best avoided. Gerrits et al. (1990) investigated a range of plasticizers for GMA and for routine work now recommend the use of polyethylene glycol (PEG) 400 as an alternative to BE. It has a low toxicity and is a component of Technovit 7100 (Kulzer) and HistoResin (Leica). A different plasticizer with low toxicity, 2-isopropoxyethanol (IP), is used in Technovit 8100 (Kulzer) (Table 7.3). 7.2.2

Polymerization of glycol methacrylate

Gerrits and his colleagues (Gerrits et al. 1991) have made extensive studies of various initiator-accelerator systems for the polymerization of GMA,

G M A + cross-linker, E G D

GMA

Technovit 8100

JB-4

BE

IP

PEG 400

Plasticizer

•

solution

A

B P O (JB-4 Catalyst)

B P O (Hardener I)

B P O (I Iardener I)

Initiator

•

N,N,-Dimethylaniline

UB-4 Solution B)

A/,/V,3,5-Tetramethylaniline (Hardener II)

Barbituric acid derivative (Hardener II)

Accelerator

Modified I m m Gen-its and Eppinger (1995).

BE, 2 butoxyetlunol; BPO, dibcn/oyl peroxide (oxygen donor); EG I), ethylene glycol dimethacrylate; G M A , 2-hydroxyethyl methacrylate; IP, 2 i.sopropoxycthanol; PEC!, polyethylene glycol; XCl, source of chloride ions.

G M A f co-catalyst, XCI

Monomer

Infiltration

Stock solutions

Technovit 7100