Light Microscopy in Biology: A Practical Approach

Light Microscopy in Biology

The Practical Approach Series SERIES EDITOR B. D. HAMES Department of Biochemistry and Molecular Biology University of Leeds, Leeds LS2 9JT, UK

See also the Practical Approach web site at http://www.oup.co.uk/PAS * indicates new and forthcoming titles Affinity Chromatography Affinity Separations Anaerobic Microbiology Animal Cell Culture (2nd edition) Animal Virus Pathogenesis Antibodies I and II Antibody Engineering if Antisense Technology Applied Microbial Physiology Basic Cell Culture Behavioural Neuroscience Bioenergetics Biological Data Analysis Biomechanics—Materials Biomechanics—Structures and Systems Biosensors Carbohydrate Analysis (2nd edition) Cell-Cell Interactions The Cell Cycle Cell Growth and Apoptosis * Cell Separation

Cellular Calcium Cellular Interactions in Development Cellular Neurobiology if Chromatin * Chromosome Structural Analysis Clinical Immunology Complement * Crystallization of Nucleic Acids and Proteins (2nd edition) Cytokines (2nd edition) The Cytoskeleton Diagnostic Molecular Pathology I and II DNA and Protein Sequence Analysis DNA Cloning 1: Core Techniques (2nd edition) DNA Cloning 2: Expression Systems (2nd edition) DNA Cloning 3: Complex Genomes (2nd edition) DNA Cloning 4: Mammalian Systems (2nd edition)

if Drosophila (2nd edition) Electron Microscopy in Biology Electron Microscopy in Molecular Biology Electrophysiology Enzyme Assays Epithelial Cell Culture Essential Developmental Biology Essential Molecular Biology I and I * Eukaryotic DNA Replication Experimental Neuroanatomy Extracellular Matrix Flow Cytometry (2nd edition) Free Radicals Gas Chromatography Gel Electrophoresis of Nucleic Acids (2nd edition) if Gel Electrophoresis of Proteins (3rd edition) Gene Probes 1 and 2 Gene Targeting Gene Transcription * Genome Mapping Glycobiology * Growth Factors and Receptors Haemopoiesis * High Resolution Chromotography Histocompatibility Testing HIV Volumes 1 and 2 if HPLC of Macromolecules (2nd edition) Human Cytogenetics I and II (2nd edition) Human Genetic Disease Analysis

if Immobilized Biomolecules in Analysis Immunochemistry 1 Immunochemistry 2 Immunocytochemistry if In Situ Hybridization (2nd edition) lodinated Density Gradient Media Ion Channels * Light Microscopy (2nd edition) Lipid Modification of Proteins Lipoprotein Analysis Liposomes Mammalian Cell Biotechnology Medical Parasitology Medical Virology MHC Volumes 1 and 2 if Molecular Genetic Analysis of Populations (2nd edition) Molecular Genetics of Yeast Molecular Imaging in Neuroscience Molecular Neurobiology Molecular Plant Pathology and II Molecular Virology Monitoring Neuronal Activity Mutagenicity Testing * Mutation Detection Neural Cell Culture Neural Transplantation Neurochemistry (2nd edition) Neuronal Cell Lines NMR of Biological Macromolecules

Non-isotopic Methods in Molecular Biology Nucleic Acid Hybridisation Oligonucleotides and Analogues Oligonucleotide Synthesis PCR 1 PCR 2 *PCR 3:PCR In Situ Hybridization Peptide Antigens Photosynthesis: Energy Transduction Plant Cell Biology Plant Cell Culture (2nd edition) Plant Molecular Biology Plasmids (2nd edition) Platelets Postimplantation Mammalian Embryos Preparative Centrifugation Protein Blotting if Protein Expression Vol 1 * Protein Expression Vol 2 Protein Engineering Protein Function (2nd edition) Protein Phosphorylation Protein Purification Applications Protein Purification Methods Protein Sequencing Protein Structure (2nd edition) Protein Structure Prediction Protein Targeting Proteolytic Enzymes

Pulsed Field Gel Electrophoresis RNA Processing I and II if RNA-Protein Interactions Signalling by Inositides Subcellular Fractionation Signal Transduction * Transcription Factors (2nd edition) Tumour Immunobiology Last 10 published Chromatin 1.4.98 Drosophila 2/e 19.3.98 Molecular Genetic Analysis of Populations 2/e 19.3.98 Mutation Detection 15.2.98 Antisense Technology 18.12.97 PCR 3:ISH.12.98 Genome Mapping 21.8.98 MHC Volume 2 21.8.97 MHC Volume 121.8.97 Signalling by Inositides In prod HPLC of Mac 2/e RNA-Protein Interactions Growth Factors and Receptors Cell Sepn Light Mic ISH2/e GEP3/e Immobilized Biomolecules Eukaryotic DNA Replication Chromosone Structural Analysis Protein Expression Vol 1

Light Microscopy in Biology Second Edition A Practical Approach Edited by

ALAN J. LACEY Department of Biology and Biochemistry, Brunei, The University of West London, Uxbridge, Middlesex UB8 3PH, UK

OXFORD UNIVERSITY PRESS

OXTORD UNIVERSITY PRESS

Great Clarendon Street, Oxford OX2 6DP Oxford University Press is a department of the University of Oxford and furthers the University's aim of excellence in research, scholarship, and education by publishing worldwide in Oxford New York Athens Auckland Bangkok Bogota Buenos Aires Calcutta Cape Town Chennai Dar es Salaam Delhi Florence Hong Kong Istanbul Karachi Kuala Lumpur Madrid Melbourne Mexico City Mumbai Nairobi Paris Sao Paulo Singapore Taipei Tokyo Toronto Warsaw and associated companies in Berlin Ibadan Oxford is a registered trade mark of Oxford University Press Published in the United States by Oxford University Press Inc., New York © Oxford University Press 1999 All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, without the prior permission in writing of Oxford University Press. Within the UK, exceptions are allowed in respect of any fair dealing for the purpose of research or private study, or criticism or review, as permitted under the Copyright, Designs and Patents Act, 1988, or in the case of reprographic reproduction in accordance with the terms of licenses issued by the Copyright Licensing Agency. Enquiries concerning reproduction outside those terms and in other countries should be sent to the Rights Department, Oxford University Press, at the address above. This book is sold subject to the condition that it shall not, by way of trade or otherwise, be lent, re-sold, hired out, or otherwise circulated without the publisher's prior consent in any form of binding or cover other than that in which it is published and without a similar condition including this condition being imposed on the subsequent purchaser Users of books in the Practical Approach Series are advised that prudent laboratory safety procedures should be followed at all times. Oxford University Press makes no representation, express or implied, in respect of the accuracy of the material set forth in books in this series and cannot accept any legal responsibility or liability for any errors or omissions that may be made. A catalogue record for this book is available from the British Library Library of Congress Cataloging in Publication Data (Data available) ISBN 0 19 963670 2 (Hbk) 0 19 963669 9 (Pbk) Typeset by Footnote Graphics, Warminster, Wilts Printed in Great Britain by Information Press, Ltd, Eynsham, Oxon.

Preface Preparing a new edition of an originally successful first edition must always result in a choice having to be made as to what to leave out to make room for any new material. Optical microscopy has developed very fast in the last ten years and yet the same principles of achieving a good image by the interaction of light with the matter under investigation are still there. Developing techniques in confocal and near-field microscopy have been given chapters as have calcium ion and pH imaging which reflect something of the enormous development of fluorochromes and fluorescent microscopy. Video cameras and video recorders used to record images for visual examination or to lead on to image analysis by subsequent processing in digital or other format have all made large contributions to the development of optical microscopy. Basic optical microscopy in which aspects of resolution of fine detail and the requirement of contrast to make that small detail visible has been retained. The recording of the image by wet chemistry methods is still a routine requirement in many projects but the use of video cameras and printers is beginning to challenge the dark-room photochemical production of still images. The basic immunohistochemistry and that of differential staining particularly in the preparation of tissues and chromosomes has been retained while new emphasis has been given to such topics as fluoroprobes for calcium and pH imaging and microinjection of materials into living cells. The use of optical microscopy techniques such as near-field and nanovid to push the resolution beyond the traditionally accepted diffraction limit attempts to bridge the gap between optical and electron microscopy. Surface details of living cells are very important components in the study of drug reaction and these together with cells to surface contacts are assisted by the techniques of reflection-contrast microscopy and evanescent illumination. The editor wishes to thank all the many colleagues, and particularly Dr Peter Hobson, for their patience in technical discussions. Mr D. J. Thomson is gratefully acknowledged for his help in many practical ways. Thanks are due to the production team of OUP who have been so helpful in preparing this book over a long period. Oxford Jan 1999

A. J. L.

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Contents List of Contributors Abbreviations 1. Basic optical microscopy A. J. Lacey 1. Introduction 2. The microscope and its use 3. Summary of the process of image formation in the microscope Lamp collector lens and condensers

4. 5. 6. 7. 8.

Kohler illumination Resolution in the microscope Magnification Interim summary Contrast methods Bright-field Dark-field Phase-contrast Differential interference contrast (DIC) Fluorescence microscopy Provisional summary of contrast techniques Summary of contrast techniques

9. Recording the image Microscope/camera attachments Summary

Acknowledgements References

2. Introduction to confocal microscopy P. J. Shaw 1. Introduction 2. The problem of out-of-focus light The confocal principle: explanation by ray optics

xix xxi 1 1 1 3 3

7 10 14 14 16 16 17 23 25 26 28 29

29 30 37

43 43

45 45 46 47

Contents Linear, shift-invariant imaging and the point spread function The shape of the point spread function Aberrations and the limits to linear, shift-invariant imaging

48 49 51

J. Practical implementation of confocal scanning systems Point scanning Slit scanning Spinning disc Two-photon imaging

54 54 55 56 57

1. Comparison of conventional, wide-field fluorescence imaging with confocal fluorescence imaging Noise and resolution Out-of-focus light When should confocal microscopy be used?

58 58 60 61

5. Practical examples of specimen preparation for confocal imaging

63

References

3. Video microscopy

70

73

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen 1. Video microscopy and the equipment required Introduction General strategies of electronic image improvement The different video-microscopic techniques Electronic equipment for video microscopy Considerations on the microscope

73 73 77 78 89 101

2. High resolution: video-enhanced contrast microscopy Different types of video-enhanced contrast microscopy Sample preparation Procedure for image generation Interpretation Typical applications and limitations

106 106 108 110 116 117

3. Low light: video-intensified microscopy Introduction Procedure for image generation Typical applications

123 123 123 127

4. Image analysis: video-based techniques for measurements in living cells Spatial measurements and motion analysis Intensity measurements

130 131 132

Contents 5. Documentation and presentation of video microscopy data Video recording Obtaining printouts for presentation and publication Preparing and presenting video sequences

133 133 141 143

Acknowledgements

146

References

146

Further reading

148

4. Microscopy of chromosomes

151

A. T. Sumner and A. R. Leitch 1. Introduction

151

2. Methods of preparing chromosomes Routine preparation of mammalian chromosomes Preparation of cells by cytocentrifugation for immunocytochemical studies of chromosomes Preparation of chromosomes from plant cells Assessment of the quality of chromosome preparations

151 152

3. Uniform (solid) staining of chromosomes

160

4. Chromosome banding The classification of chromosome bands C-banding G-banding Ag-NOR staining for nucleolus organizing regions CREST labelling of kinetochores

161 161 162 164 164 166

5. In situ hybridization Probe preparation In situ hybridization reaction

168 170 173

6. Observation and recording of images of chromosomes Observation of banding with absorbing dyes Observation of fluorescent chromosomes Photography of chromosomes Other methods of image capture

177 178 178 180 181

References

155 156 159

183

5. Immunohistochemistry

185

Michael G. Ormerod and Susanne F. Imrie 1. Introduction

185

2. Antibodies

186

xi

Contents Immunoglobulin structure Polyclonal antisera Monoclonal antibodies Purification of antibodies Specificity of antibody reactions Storage of antibodies

186 187 187 188 188 189

3. Effect of tissue processing on antigens Choosing conditions for processing Revealing hidden antigens

189 189 192

4. Choice of label Fluorescent labels Enzymatic labels Colloidal gold Selecting a label

194 194 194 197 197

5. Methods of application The direct method The indirect method Enzyme-anti-enzyme methods Systems using biotin-avidin Other methods

197 198 198 199 200 201

6. Experimental methods A general method Choosing the correct dilution of antibody Fluorescent labels Peroxidase Alkaline phosphatase Glucose oxidase Galactosidase Immunogold Some general procedures

202 202 203 204 204 207 209 209 209 210

7. Controls and problem solving Controls Problem solving

213 213 213

8. Detecting two antigens on the same section

214

9. Cytological preparations 10. Quantification

215 218

11. Equipment

218

References

219

6. Calcium and pH imaging in living cells Richard M. Parton and Nick D. Read 1. Introduction

221

xii

Contents 2. Study of calcium and pH in living cells

221

3. Imaging intracellular free calcium and pH

222

4. Fluorescent dyes for free calcium and pH Properties of calcium and pH dyes Intracellular dye behaviour Single wavelength dyes, ratiometric dyes, and ratiometric dye-pairs

223 223 225 230

5. Introducing calcium and pH dyes into living cells General considerations Ester loading Low pH loading Scrape loading Electroporation lonophoretic microinjection Pressure microinjection Quantifying the extent of dye loading

231 231 232 235 235 235 236 236 237

6. Equipment for fluorescence microscopy Fluorescence microscopes Objectives Dye excitation sources for ion imaging Filters for ion imaging

237 237 238 239 239

7. Fluorescence imaging systems General requirements Conventional fluorescence imaging Confocal imaging Multiphoton imaging Imaging with multiple detectors

240 240 241 243 244 245

8. Optimizing the performance of imaging systems

245

9. Handling experimental material on the microscope stage

248

10. Digital image processing

249

11. Ratioing

249

12. Quantitative ion imaging Image quality and quantitative imaging Signal-to-noise ratio Numerical data extraction Quantitative ratio imaging Calibration of dye response Statistical analysis of image data

252 252 254 256 257 260 264

13. Visual presentation of image data Visual image enhancement Preparation of digitized figures and plates for publication

267 267 267

14. Combining ion imaging with other experimental techniques

269

15. Critical controls for intracellular ion imaging

270

xiii

Contents Acknowledgements References

271 271

7. Reflection-contrast microscopy J. S. Ploem, F. A. Prins, and I. Cornelese-ten Velde 1. Introduction Methodology Applications Review articles

275 275 275 275 276

2. Optical systems for RCM Early developments in reflected-light microscopy New developments in reflection-contrast microscopy Modern reflection-contrast microscopes 3. Image formation General

276 276 277 280 286 286

Image formation in RCM of living unstained cells Image formation in RCM of stained specimens

289 289

4. Applications General Special applications 5. Specimen preparation General Immunohistochemistry Fixation and embedding Sectioning Immuno- and histochemical staining for RCM Mounting and examining sections on microscope slides 6. Summary Acknowledgements References

290 290 295 298 298 298 301 303 305 307 308 309 309

8. Histomorphometry

311

A. J. Reynolds \. Introduction 2. Microscopy Specimen preparation Obtaining an image

311 313 313 318

Calibration

319

3. Linear measurements Intercept measurement Point counting

321 321 324 xiv

Contents 4. Automated measurement Measurement with an interactive computer system (digitizer tablet) Semi-automatic image analysers

327 327 329

5. Histomorphometry

337

Acknowledgements

338

Instrumentation and sources of supply

338

References

339

9. Near-field optical microscopy

341

Niek F. van Hulst \. Introduction Optical microscopy Probe microscopy Breaking the diffraction limit Scope of this chapter

341 341 342 342 344

2. Instrumentation Probes and distance regulation Antenna-type or 'appertureless' near-field scanning optical microscopy Aperture-type near-field scanning optical microscopy Photon scanning tunnelling microscopy (PSTM) Distance regulation: shear force microscopy Conclusion

344 344

3. Applications Single fluorophores and proteins Monolayers and aggregates Virus Cellular surface and cytoskeleton Chromosomes and fluorescence in situ hybridization

355 355 358 362 362 363

4. Conclusions

367

5. Future outlook Acknowledgements

369 369

References

346 346 349 349 350

370

10. Introduction of materials into living cells 1. Particle bombardment as a means of DNA transfer into plant cells Christian Schdpke and Claude M. Fauquet Introduction Practical considerations Identification of cells transformed with reporter genes xv

373 373 373 374 380

Contents 2. Microinjection as a preparative technique for microscopical analysis 386 H. F. Paterson Advantages of microinjection by glass capillary needle Equipment required for microinjection of adherent mammalian cells Preparation of materials for microinjection Microinjection technique Microscopical analysis of microinjected cells

387 391 392 394

Acknowledgements

396

References

396

11. Surface fluorescence microscopy with evanescent illumination

386

399

D. Axelrod 1. Fluorescence at surfaces TIRF for biochemical samples TIRF for biological samples

399 399 401

2. Theory of TIRF Single interfaces: intensity and polarization Intermediate dielectric layers Intermediate metal film

401 401 405 406

3. Optical configurations Inverted microscope TIR with prism on top Inverted microscope TIR with prism below Upright microscope TIR with prism below Inverted microscope TIR without a prism Rapid chopping between TIR and epi-illumination

407 407 410 411 414 417

4. General experimental suggestions

417

5. Applications of TIRF microscopy

420

6. Comparison with other optical sectioning microscopies

422

Acknowledgements

423

References

423

12. Nanovid microscopy

425

Greta M. Lee 1. Introduction Use of nanovid microscopy to analyse molecular mobility in membranes Use of nanovid microscopy to study the extracellular matrix

xvi

425 428 428

Contents 2. General information on equipment and methods Microscope Camera Image processor and frame grabber Image storage

432 432 433 433 434

3. Image analysis to analyse molecular mobility (single particle tracking)

434

References

435

Appendix I

437

Appendix II

443

Index

447

XVll

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Contributors D. AXELROD

University of Michigan, Biophysics Research Division, 930 North University, Ann Arbor, Michigan 48109-1055, USA. I. CORNELESE-TEN VELDE

Department of Pathology, Leiden University, PO Box 9600, 2300 RC Leiden, The Netherlands. CLAUDE M. FAUQUET

ILTAB/ORSTOM, The Scripps Research Institute, Division of Plant Biology - BCC 206,10550 North Torrey Pines Road, La Jolla, CA 92037, USA. SUSANNE F. IMRIE

34, Springfield Road, Wallington SM6 OBB, UK. A. J. LACEY

Department of Biology and Biochemistry, Brunei, The University of West London, Uxbridge, Middlesex UB8 3PH, UK. GRETA M. LEE

Thurston Arthritis Research Center, CB 7280, 5107 Thurston Building, University of North Carolina, Chapel Hill, NC 27599, USA. A. R. LEITCH

School of Biological Sciences, Queen Mary and Westfield College, University of London, London El 4NS, UK. WILLIMAILE

Institut fur Zelltechnologie e.V., Technologie-Park Warnemiinde, FriedrichBarnewitz-Str. 4, D-18119 Rostock-Warnemiinde, Germany. MICHAEL G. ORMEROD

34, Wray Park Road, Reigate RH2 ODE, UK. RICHARD M. PARTON

Molecular Signalling Group, Institute of Cell and Molecular Biology, University of Edinburgh, Rutherford Building, Edinburgh EH9 3JH, UK. H. F. PATERSON

CRC Centre for Cell and Molecular Biology, Chester Beatty Laboratories, Institute of Cancer Research, 237 Fulham Road, London SW3 6JB, UK.

Contributors J. S. PLOEM Pres. Kennedylan 256, 2343 GX Oegstgeest, The Netherlands. F. A. PRINS

Department of Pathology, Leiden University, PO Box 9600,2300 RC Leiden, The Netherlands. NICK D. READ

Fungal Cell Signalling Group, Institute of Cell and Molecular Biology, University of Edinburgh, Rutherford Building, Edinburgh EH9 3JH, UK. A. J. REYNOLDS

Experimental Techniques Centre, Brunei, The University of West London, Uxbridge, Middlesex UB8 3PH, UK. CHRISTIAN SCHOPKE

ILTAB/ORSTOM, The Scripps Research Institute, Division of Plant Biology - CAL 7,10550 North Torrey Pines Road, La Jolla, CA 92037, USA. P. J. SHAW

Department of Cell Biology, John Innes Centre, Colney, Norwich NR4 7UH, UK. WALTER STEFFEN

Mikroskopiezentrum, Institut fur Zellphysiologie und zellulare Biosysteme, Fachbereich Biologie, Universitat Rostock, Universitatsplatz 2, D-18051 Rostock, Germany. A. T. SUMNER

7 Smileyknowes Court, North Berwick, East Lothian EH39 4RG, UK. NIEK F. VAN HULST

Applied Optics Group, Faculty of Applied Physics, MESA Research Institute, University of Twente, PO Box 217, 7500 AE Enschede, The Netherlands. DIETER G. WEISS

Lehrstuhl fiir Tierphysiologie, Institut fur Zellphysiologie und zellulare Biosysteme, Fachbereich Biologie, Universitat Rostock, Universitatsplatz 2, D-18051 Rostock, Germany. ROBERT A. WICK

PricewaterhouseCoopers LLP, Ten Almaden Blvd., Suite 1600, San Jose, CA 95113, USA.

xx

Abbreviations Ab antibody AFM atomic force microscopy AM acetoxymethyl AOM accousto-optical modulator b.f.p. back focal plane BSA bovine serum albumin CaMV cauliflower mosaic virus CCD charge coupled device CHO Chinese hamster ovary CLSM confocal laser scanning microscope DAB diaminobenzidine DAPI 4' ,6-diamidino-2-phenylindole DIC differential interference contrast DNP dinitrophenol DPX distrene, dibutylphthalate, xylol DTT dithiothreitol EM electron microscopy FAC fluorescent analogue cytochemistry FISH fluorescence in situ hybridization FITC fluorescein isothiocyanate FVN field of view number FWHM full width half-maximum GFP green fluorescent protein gfp gene encoding green fluorescent protein GUS (3-glucuronidase HRP horseradish peroxidase IAD illuminating aperture diaphragm IFD illuminated field diaphragm Ig immunoglobulin IGSS immunogold silver staining ILTAB International Laboratory for Tropical Agricultural Biology LM light microscopy LUC luciferase luc gene encoding luciferase LUT look-up table NA numerical aperture NIH National Institutes of Health NOR nucleolar organizing regions NP-40 Nonidet P-40 NSOM near-field scanning optical microscopy

Abbreviations PA PBS PCR PHA PMSF PO POL p.s.f. PSTM RCM RGB ROI RSE SCV SDS SIT S/N TEM TIFF TIRF TPCLSM uidA VEC VIM X-gluc

'pro analyse' quality reagents phosphate-buffered saline polymerase chain reaction phytohaemagglutinin phenylmethylsulfonyl fluoride peroxidase polarization point spread function photon scanning tunnelling microscopy reflection-contrast microscopy red, green, blue region of interest relative standard error settled cell volume sodium dodecyl sulfate silicon intensifier target signal-to-noise transmission electron microscopy tagged image file format total internal reflection fluorescence two-photon laser scanning microscope gene encoding p-glucuronidase video-enhanced contrast video-intensified microscopy 5-bromo-4-chloro-3-indolyl-p-D-glucuronide cyclohexylammonium salt

xxn

1

Basic optical microscopy A. J. LACEY

1. Introduction The perception of, or the sight of, an object whether in a microscope or in an everyday situation is the result of a complex process. It begins with light interacting with the object. The modified light is collected by the lens of the eye and brought to a focus on the retina of the eye. The nerve responses in the retina are interpreted by the eye/brain system programmed to a greater or lesser extent by previous experience. The area of the specimen seen by the eye is the perception of the extension in x and y directions, while the experience of the of the z direction is the perception of depth.

2. The microscope and its use Protocol 1 gives a practical guide to establishing the potential of the hardware of a microscope. Protocol 1. Examination of the microscope potential Equipment Microscope and accessories

A. Lamp(s) 1. Trace the power supply from the mains to the light source. Discover type of lamp available, i.e. tungsten halogen, high pressure mercury, or other. Locate centring screws for lamp relative to its collector lens. There may be two separate light sources one for transmitted and another for epi-illumination or reflected light work. Locate the slider which blanks off the incident light train. There may be a focus control for the lamp collector lens. Establish its function (see Kohler illumination protocol. Protocol 2).

A. J. Lacey Protocol 1.

Continued

2. Find voltage control for tungsten light and the colour temperature setting. 3. There may be neutral density filters in the light path. B. Condensers 1. Note carrier of condenser and find centration screws. 2. Note type, i.e. universal, bright-field with top lens control by screw off or flip top. 3. Note the appearance of the lamp side of the condenser, i.e. the phase annuli, dark-field stop, or Wollaston prism, illuminating aperture diaphragm, and any centration facility for each of these. Engravings may include lens corrections such as NA (numerical aperture), apochromatic. Direction of shear is shown in DIC condensers. C. Objectives (usually a range of objectives in a rotatable turret) 1. Numerical aperture (NA), e.g. 0.65 or 1.3 (possible iris for control of NA). 2. Magnification 40:1 (or just 40), 100, X100 (infinity corrected) x 40 (angular magnification) x 100. 3. Tube length: 160 mm or infinity (°°). 4. Coverslip (thickness) 0.17 (correction collar for high dry NA). 5. Immersion (otherwise dry): usually oil but sometimes water or glycerine. 6. Lens type: Plan, Phaco, Apo, Fl, etc. 7. Manufacturer and code number. D. Eyepieces Often inserted as a pair in a head with interpupillary distance control. The right-hand one may have a diopter control for balancing the focus of the two eyes. The head may have a slider to bring in a beam splitting prism to take all or some of the light to a camera port. 1. Engravings: (a) Magnification: x 10, x 15. (b) Field of view number (FVN): 18, 14. This gives an indication of the field of view but needs converting to mm by reference to objective mag. and tube factor—see note (a). (c) Type (field): wide-field (WF). (d) Corrections: compensating. (e) High eyepoint: spectacles symbol.

1: Basic optical microscopy 2. Note whether there is a magnification changer present in the tube of the microscope and whether such a changer has a telescope (phase) position and a focus control for it.

Notes: (a) The field of view in mm =

The tube factor may be X 1.25 for a binocular head but it can be found by calculation substituting a measured field of view in the above equation. (b) It is important to match the lenses in use at any one time. Thus the very highest corrected objectives should be used with highly corrected condensers and appropriate eyepieces.

3. Summary of the process of image formation in the microscope The summary is illustrated graphically in Figure 1. (a) (b) (c) (d) (e) (f) (g) (h) (i) (j)

The specimen is illuminated. The specimen and the light interact. Part of the light is scattered. Part of the light is unchanged. The objective collects the light. What is collected by the objective is seen (as an optical transform of the specimen) in the back focal plane (b.f.p.) and passes on through it. The light issuing from the b.f.p. passes to the primary image plane where it forms an interference pattern (a further optical transform). This interference pattern is the primary image produced by the objective and is a magnified replica of the specimen (see Figure la). The light passing on is further modified by angular alteration in the eyepiece (Figure 1b). It passes out through the exit pupil of the microscope and into the entrance pupil of the eye with an angle which is perceived as further magnification (Figure 1b).

3.1 Lamp collector lens and condensers The light issuing from the filament is collected by a lamp collector lens which focuses it on to the illuminating aperture diaphragm. The condenser then col-

Figure 1. (a) Summary of image formation in the microscope. Main steps in the process are indicated on the right. P, Q, R represent features in the specimen which are imaged at P1, Q1and R1. (b) Geometrical optics of the light microscope. fe = focal length of the eyepiece; fo = focal length of the objective; To = tube length (notionally = 160 mm); h = specimen dimension; H = intermediate (primary) image dimension; H' = final (virtual) image dimension; Dv = least distance of distinct vision (250 mm).

lects the light and puts it as a cone whose apex falls into the specimen. This practical process is given in Protocol 2 (Kohler illumination). Condensers are precision lenses with colour and spherical aberration corrections and are often capable of several contrast techniques—see under contrast methods (Section 8).

1: Basic optical microscopy

3.1.1 Light matter interactions The light then interacts with the specimen in some or all of the following ways: absorption transmission reflection refraction diffraction polarization absorption and subsequent re-emission—fluorescence or phosphorescence The cone of light which entered the specimen is scattered to a greater or lesser extent by the specimen (Figure la) and some may be lost out of the sytem. For the simple case the central ray of light passes on up the optical axis

A. J. Lacey of the microscope. Some of its energy may be scattered at an angle by diffraction or refraction from this straight line. The angle, of diffraction for example, will be related to the fineness of the patterns in the specimen. Protocol 3 illustrates this process. 3.1.2 Objectives The objective lens in a microscope collects the undeviated light and the scattered light to an extent determined by the angular aperture of the objective or more precisely by its numerical aperture (NA). That light which is collected by virtue of the numerical aperture passes on through the back focal plane of the objective (b.f.p.) and is brought to a plane (the primary image plane) where the various components of the light (the direct light and the scattered light) interfere with each other and form an interference pattern called the primary image. This image is a magnified replica of the specimen. The NA of the objective is an important component in determining the resolving power of the microscope as shown in Equation 1.

where lambda is the wavelength of light used (e.g. 500 nm or 0.5 jxm for green light) and NAobj the numerical aperture of the objective. The resolving power of the objective can be calculated thus for a 0.65 NA objective as about 500 nm or 0.5 |xm and for a NA of 1.3 as half these values (twice the resolving power) (see Protocol 3 for the explanation in practical terms). One of the engravings on the objectives indicates the primary magnification that is the ratio of the primary image size to that of the specimen. The primary image resulting from the objective is then effectively further magnified by the eyepiece lenses, and seen by the eye. 3.1.3 Eyepieces The light passes on from the primary image plane and is altered in angular terms as it passes through the eyepieces. This alteration is perceived by the eye/brain as the final magnification of the microscope. The eye(s) is placed with its entrance pupil at the exit pupil or Ramsden disc of the microscope. The Ramsden disc can be found by following Protocol 2. The eyes should be set comfortably over the eyepieces—both open even if there is only one eyepiece. They will eventually learn to be in a relaxed state, i.e. focused on infinity and viewing the image as though it were at infinity. The distance apart of the binocular eyepieces (the interpupillary distance) should be set by finding the position of a single image seen through both eyes. The separation given on the scale should be memorized. The left eye system is focused first by the microscope focus control, shielding the right eye by a piece of paper rather than closing it, and then the right eyepiece is brought to a comfortable focus, shielding the left eye, by altering the diopter setting on one of the eyepieces. Again it can be noted for future reference.

1: Basic optical microscopy

4. Kohler illumination In order to obtain an even distribution of light on the specimen and an even background intensity a sequence of lamp collector and of condenser lenses is used. The principles of the method are: (a) A lens in front of the light source places an image of that light source at a position which is not in the plane which contains the specimen. (b) A second lens (the condenser) puts an image of the surface of the first lens onto the specimen to be examined and does so with as wide an aperture as possible to illuminate the specimen. Protocol 2. To obtain Kohler illumination in a transmitted light microscope and to establish conjugate planes Equipment Microscope with light source, illuminated field diaphragm (IFD), illuminating aperture diaphragm (IAD) (an optional extra would be a phase telescope)

Contrasty specimen such as a periodic acid Schiff/Light Green or Masson trichrome stained specimen

Method 1. Examine the microscope (for an unfamiliar machine) carefully to establish the physical control procedures. Trace the pathway from the mains wall switch through to the tungsten or quartz halogen light source with its intensity control filters, IFD, condenser with its IAD, and focus control, objectives (type and range) to the eyepieces (again type and range). There may be supplementary magnification changers in the tube of the microscope and other modifications may include light splitting prism in the trinocular head. 2. The microscope may also be capable of reflected light microscopy with filter blocks appropriate for fluorescence work. These should be slid out of the light train for transmitted work. 3. Switch on the light and check its centration, following the maker's instructions, with respect to the lamp collector lens. 4. Using the contrasty specimen focus an image of it through a low power objective such as x 10. 5. Looking through the eyepiece partially close the IFD until it is seen in the field of view, i.e. in the plane of the specimen. 6. Centre the image of the IFD by the use of centration screws on the condenser. Focus the condenser to give the field iris a sharp image. In moving the condenser focus you may notice that the image of the IFD has a coloured edge. This should be of one colour only and change

A. J. Lacey Protocol 2. Continued

7.

8.

9.

10.

11.

evenly otherwise suspect poor alignment. Check alignment instructions as appropriate. Open the IFD until it is just outside the field of view of the microscope. This procedure will also provide more critical centration. It is at this stage that in really low power objectives the image of the field iris may not fill the field of view. In this case either a supplementary lens may be brought into the light path below the condenser or the flip top lens removed from the condenser. There may then be need to refocus the condenser (step 6). Take out the eyepiece and insert the phase telescope (or use the builtin Bertrand lens), to focus the telescope on the back focal plane (b.f.p.) of the objective. This can be recognized by seeing the image of the IAD in view. Test that this is the case by opening and closing that diaphragm. There may be capacity for centring the aperture image relative to the b.f.p. If this is so then do so but being careful not to meddle with the centring screws of the condenser itself which you have already been set. Adjust the IAD so that it fills about 70% of the b.f.p. of the objective. You will notice that some glare disappears from the side of the body tube of the microscope as you make this adjustment. You may also notice that there is an image of the light source in the centre of this bright illuminating aperture. This is as it should be. If there is a facility for centring the lamp filament then do so with respect to the image of the condenser iris and the b.f.p. These three are said to be conjugate (see later). The presence of a diffuser near the light source may not allow the image of the filament to be seen so the maximum brightness point should be centred. Replace the eyepiece and assume that this is the best situation for the particular combination of condenser and objective settings. Excessive brightness in the image should be reduced by the insertion of neutral density filters into the light path and not by reducing the IAD. For visual microscopy it is permissible to reduce the intensity of the light source but this will increase the warmth of the colours and is not appropriate for colour photomicrography (see later). For a change in objective, i.e. to a x 40, check steps 5-9. You will probably find that all that is required is to increase the IAD to match the increased size of the b.f.p. in the higher numerical aperture (NA) objective. When returning to a lower NA objective remember to again check steps 5-9. To complete the understanding of the conjugate planes in the microscope place a piece of ground glass or lens tissue at right angles to the light path just above the eyepiece. Move the tissue or glass along the light path towards or away from the surface of the eyepiece. You

8

1: Basic optical microscopy will discover a place where the apparent disc of light is at a minimum dimension. This disc of light is the exit pupil of the microscope, otherwise known as the Ramsden disc. In correct conditions it also acts as the entrance pupil of the eye. It is about 3 mm in diameter and is conjugate with the b.f.p. Note for spectacle wearers it is some greater distance (15 mm) from the eyepiece and an appropriate eyepiece will be marked with a engraving of spectacles. 12. Compile a list of the conjugate planes in which that of the b.f.p. is included. Note that this series is as follows: lamp filament, IAD, b.f.p.of objective and Ramsden disc. 13. Find the other series which includes the IFD, the specimen, the primary image, and the retina of the eye (or the emulsion of the film or chip of the video camera). The primary image lies in the eyepiece just outside the inner focal point of the eyepiece eye lens and a diaphragm (the field diaphragm of the eyepiece) there gives the image a clear edge. A field lens also occurs in the eyepiece which controls the area of the primary image entering the eye. 14. Kohler illumination successfully keeps the irregularities of intensity in the light source out of the series which contains the specimen.

These principles are the basis for Protocol 2. The planes which contain the lamp filament or its image comprise one series and those containing the specimen and its image comprise another and are set out in diagrammatic form in Figure 2 and in Table 1.

Figure 2. A ray diagram of Kohler illumination showing the positions of the two sets of conjugate planes. Solid lines represent rays of light moving from left to right arising from the centre of the lamp filament to the near margin of the lens. Dotted line is a ray passing through the centre of the lamp collector lens. Vertical solid lines with arrow heads are lenses, while vertical solid lines without arrow heads are diaphragms (irises). Vertical dotted lines are planes in which specimen, back focal plane (b.f.p) of objective and image are situated. Zigzag represent the lamp filament and its subsequent images.

9

A. J. Lacey Table 1. Conjugate planes in Kohler illumination Series A (aperture set)

Series B (field set)

Lamp filament (source) Illuminating aperture diaphragm (IAD) Back focal plane of objective (b.f.p.) Ramsden disc

Illuminated field diaphragm (IFD) Specimen Primary image Retina of eye

All modern microscopy should be commenced by setting-up Kohler illumination. The capacity of your microscope to set up Kohler illumination is thereby tested. One of the situations often met with is that the lamp collector lens may have a diffuser surface. This is sometimes the surface next to the lamp and it is so treated to break up the image of the irregularities in the light source. Other variations may be in having a ground glass just near the field iris for the same reason. The condenser aperture iris is sometimes at a considerable distance from the front focal plane of the condenser making for difficulty in losing its image from the series conjugate with the specimen.

5. Resolution in the microscope It was stated in Section 3.1.2 above that the NA of the objective is paramount in determining the details in the image. Protocol 3 sets out an example of how to demonstrate this. By combining the magnifying power of the objectives and eyepieces to produce an equal magnification it is readily possible to show that the NA of the objective is the key factor (see Figures 3a-c). Protocol 3a. To demonstrate empty magnification Equipment Microscope with objectives x 40, NA 0.65, and x 10, NA 0. 25, and eyepieces x 25 and X6

The diatom Navicula lyra mounted in DPX or other resin8

Method 1. Observe the diatom with the x 40, NA 0.65 objective together with a x 6 eyepiece to give a total magnification of X 240. Observe the detail present in the image as the objective is capable of resolving spacing of about 500 nm. 2. Repeat with an objective x 10, NA 0.25, and using the x 25 eyepiece. The combined objective/eyepiece will give a magnification of x 250. Note the general outline of the diatom is the same but in the image of its surface there is no detail. 10

1: Basic optical microscopy 3. Infer that there is excess magnification in the second case. The NA 0.25 objective is only capable of resolving spacings of larger than 1200 nm and the pattern in the diatom is smaller. Therefore there is surplus or empty magnification present. 4. The regular pattern of dots in the frustule of the diatom can be seen if the NA of the objective is greater than 0.3. The pattern is not resolved in the case of the lower NA objective. aA test plate can be obtained from Micro Instruments Ltd. in which five diatoms are mounted including N. lyra. Recommended combinations of NA 0.3 and total magnification (x 100) required to see the detail are engraved on the divisions of the test plate. Unless there is more detail revealed by additional magnification then the addition is described as empty magnification (see Figure 3). Some additional magnification however makes for more comfortable viewing.

Protocol 3b. Pleurosigma resolved Equipment Oil immersion objective with numerical aperture 1.3, and with a iris in the b.f.p. capable of restricting the b.f.p. to NA 0.8

Prepared slide of Pleurosigma angulatum

Method 1. Set up Kohler illumination. 2. Find and centre the diatom using the objective with its iris open to full 1.3. 3. Close down the condenser iris to as near a pin-hole as possible. Note image of the specimen contains a hexagonal pattern in the diatom surface (see Figure 3c). 4. Remove eyepiece and observe b.f.p. of objective. Note presence of bright central image of the aperture iris (conjugate) and also the six coloured images of this iris around the margin of the b.f.p. These are the six first order diffraction images surrounding the zero order light (Figure 3a). 5. Close the objective iris noting as you do so that you are occluding the first order diffraction (Figure 3b), i.e. preventing it from leaving the b.f.p. 6. Replace the eyepiece and observe the loss of the hexagonal pattern in the surface of the diatom (Figure 3d). Reopen the iris and retrieve the pattern. 7. Repeat steps 1-6, noting appearance of images and of the altered b.f.p.

11

A. J. Lacey Protocol 3b.

Continued

8. Conclude that the pattern is only present in the image when the first order diffracted light contributes to the formation of the image. When this occurs the microscope is said to have resolved the pattern. (Note Protocol 6 gives a practical illustration of the importance of the zero order light.)

Although Equation 1 gives the basic resolution equation of the microscope it is better to think of the effective NA as being the mean of the NAs of the objective and the condenser. It was mentioned on page 4 that the condenser throws an oblique cone of light on to the specimen. In normal Kohler the

Figure 3, The relationship of the image to what light passes through the back focal plane (b.f.p.) of the objective. The objective is a Zeiss Planapo NA 1.3-0.8, x 100, oil immersion lens. The specimen is the diatom Plourosigma angulatum [scale bar = 20 |j.m). (a) The b.f.p of the objective with fully opened iris allowing the central zero order light and the (six) first order light to pass through to forming the image with detail resolved in (c). In (b) the iris in the objective is (almost) occluding the first order diffracted light and producing an image in (d) which fails to resolve the detail in the diatom.

12

1: Basic optical microscopy illuminating aperture is about 70% of the NA of the objective. Equation 2 is a better version of the resolution formula:

provided NAobj is larger than NAcond. Protocol 4 sets out a technique for making make a test specimen of point sources of light, the testing of the performance of objectives, and the practical illustration of the Rayleigh criterion. For a definition of the Rayleigh criterion see Chapter 3, Figure 4, page 82. Protocol 4. To prepare and use a star test object Equipment and reagents Clean coverslips (No. 1 or 0.16 mm thick) and slide

Vacuum coater DPX or other mounting medium

Method 1. Holding a coverslip by the edges, place it horizontally in a sputter coater. 2. Coat one surface with an appropriate metal alloy until opaque. 3. Blow on the metallized surface of the coverslip to remove any dust particles. 4. Cement the coated side downwards on to a microscope slide. 5. Examine through microscope for the tiniest of holes. 6. Having found the tiniest of holes increase the eyepiece magnification as far as you are able, until the rings of light are visible around the central disc of light. 7. Observe and note carefully the appearance of these Airy disc patterns which are the images of the point sources of light. Use a range of objectives and make careful notes of the appearance above and below focus. 8. Examine the range of Airy disc patterns over the whole field of view looking for distortions at the edges of the field of view. 9. Note again their appearance when viewed in green, blue, or red light. 10. Returning again to white light endeavour to find two holes which are close or touching each other. There will be some holes which look dumb-bell-shaped. Are these two overlapping images or are they one of a single oddly shaped hole? Visual acuity and patient observation with accurate note taking make this specimen a very testing one not only for the student of microscopy but also for

13

A. J. Lacey a biologist who is anxious to know the quality of the performance of his/her objectives and microscope system. In the present context it is only to demonstrate that an image of a point source of light is a diffraction pattern. The radius (d') of the first dark ring is proportional to the wavelength of the light in use (smaller for blue and larger for red) and inversely proportional to the numerical aperture of the lens used to produce it. The formula is given as d' = 0.61 lambda/NA (Equation 1). That figure illustrates the intensity scan of the images of two point sources of light close together. They are sufficiently close for the diffraction rings (first and second order) of one to coincide with the main peak (zero order) of the other. The sum of the zero order peak and the other orders of diffracted light is given by the dotted line. If the difference between the peaks and the central trough of the dotted line is perceptable by the eye (usually taken as 16%) then the two points are resolved as two points (Rayleigh criterion of resolution). Video detection is more sensitive than that of the eye in that it is capable of differentiating less than 16% differences. The Sparrow criterion of resolution is where the dip in intensity between the Airy disc overlapping patterns is made visible through the aid of video-enhanced detection. Space does not allow a fuller treatment of the subject here. Point sources for fluorescence microscopy can be made by utilizing the 0.1 (xm or smaller fluorescent P. S. Speck beads available from Molecular Probes Inc. See Chapter 3 for further information on the point spread function. The Airy disc pattern obtained may show aberrations due to poor setting-up rather than intrinsic faults in the lens systems so beginners must be cautious in rejecting lenses.

6. Magnification Magnification is a two stage process in the microscope. The first stage is where the objective produces a real (inverted) magnified image (the primary image). The second stage when the rays emerging from the b.f.p. of the objective pass through the eyepiece lenses to be thereby altered in angle and to present the primary image to the eye as still further magnified. Total magnification should be at least enough to bring the detail resolved by the microscope to the resolving power of the eyes, i.e. about 2 minutes of arc. Magnification should also be within the limits set by the working pupil of the eye, i.e. within the limits of 170 X NAobj to 1000 X NAobj. Figure 4 illustrates the examination of tissue sections with various combinations of NA in the objective and magnifications.

7. Interim summary Summary of image formation in the microscope so far: (a) The image in a microscope is a replica of the specimen constructed by interference of light waves emanating from the specimen. 14

Figure 4. Magnification and resolution. Three photomicrographs taken through objectives of different apertures and magnifications, all printed at a magnification of 400:1. (a) Objective 4/0.16. Final magnification 2500 x MA, The image is unsharp and obviously over-magnified, (b) Objective 10/0.25. Final magnification 1600 x NA. The image still lacks information in fine details, (c) Objective 25/0.65. Final magnification 615 x NA, In this image the fine detail is sharply rendered. Scale bar = 50 ^m. From ref. 1 with permission.

15

A. J. Lacey (b) The microscope is, or is not, capable of resolving the detail in the specimen, when illuminated by a particular wavelength of light, by virtue of the numerical aperture of the objective. (c) The resolution is made available to the eye by the combined magnifying power of the objective and the eyepieces. (d) Resolution however may not be visible unless contrast is present.

8. Contrast methods The perception of the detail in the image is only possible if there is sufficient contrast presented to the eyes between the background and the specimen and between the various features in the specimen. The normal viewing in a transmitted light microscope is by presenting the specimen on a bright background. If however the background is too bright nothing will be seen. The contrast is proportional to the difference in intensity of the background and the specimen and inversely proportional to the intensity of the backgound. A series of practical protocols is now given to illustrate the achievement of contrast in the image.

8.1 Bright-field Bright-field (Protocol 5) contrast is due to the kind of optical system used, i.e. the specimen is seen against a bright background and its contrast is largely the result of absorption by the stains in the specimen with its required chemical fixation and mounting procedures of that specimen or by the scattering power of the specimen. The numerical aperture of the objective also determines the depth of the material which is in-focus at any one plane of focus at any one time. That is information about the z direction in the specimen likely to be confused by out-of-focus light coming from planes above and below the plane of focus (see later chapters for further treatment of this situation). Sectioning of a large specimen is required to produce as near a single layer in the z direction as possible. However the use of reduced illuminating aperture diaphragm (IAD) in achieving contrast in weakly stained material is reviewed as are the uses of colour filters in stained specimens.

Protocol 5.

Bright-field microscopy; contrast assessment

Equipment and reagents Microscope with Kohler illumination Cheek cells mounted in saliva or onion scale inner epidermis mounted in water

Set of colour filters Masson trichome and/or periodic Schiff stained animal sections

16

acid

1: Basic optical microscopy Method 1. Set up Kohler illumination as in Protocol 2. 2. High contrast situation: examine the stained specimen in white light. Recall that the contrast seen here is by differential absorption/ transmission of the many wavelengths comprising white light. The red portions transmit red light and the green areas are those transmitting green light of about 500 nm wavelength. 3. Observe the effects of using monochromatic light obtained by placing filters in the light path. Use filters of complementary colour to bring about increased contrast for example a green filter will darken a red stained area of the specimen (see Figure 5). Filters of the same colour can reduce the impact of stains and make excessively over-stained areas more transparent. 4. Low contrast situation: examine the cheek cells or the onion cells in Kohler illumination in white light as for step 1. Note virtually no contrast, i.e. nothing visible. There may be some visibility in the onion preparation by virtue of the absorbence of light in the vertical walls. 5. Try closing down the IAD and note the improvement in contrast. The nuclei and the cytoplasm of the cheek cells and also that of the onion cells will become visible. Note however that this is strictly speaking not an appropriate technique as a good deal of the information in the image is 'rotten', i.e. has diffraction rings round small features.

Conclude from this protocol that staining thin sections is a powerful method of obtaining contrast between the specimen and the backgound and between features in the specimen. Often however this staining procedure leads to loss of lifelike dimensions or produces chemical artefacts. For living cells refer to Protocol 3 for phase-contrast, and Protocol 4 for differential interference contrast (DIC).

8.2 Dark-field The theoretical background of the contrast techniques is avoided but the case of high power dark ground (Protocol 6) is used to illustrate, in the image as well as the change of contrast, the importance of the zero order light in image formation. Its occlusion in the b.f.p. can be shown in this case to produce reversed contrast in Figure 6. For low power objectives it is sometimes possible to use the illuminating annulus in the phase condenser appropriate for a higher objective NA condition. The annulus then acts as a kind of simple patch stop. Note: Rheinberg illumination. This is done exactly as for dark ground simple patch stop but substituting complementary colours for the central stop and the surrounding illumination. A pleasing image of the specimens in the colour of the surrounding annulus on a background the colour of the central stop. 17

Figure 5. The effect of colour filters on contrast. The specimen (frog stomach section) is stained with periodic acid Schiff and Light Green, photographed on Kodak Technical Pan film, and developed for medium contrast. The appearance of the micrographs is closely similar to that seen directly with the eye. (at No filter. The tips of the cells, densely stained magenta, appear slightly darker than the rest of the specimen which is in shades of green, (b) Red filter. The magenta areas now appear colourless and the green stained nuclei are dark and more conspicuous, (c) Green filter. The green stained nuclei cannot now be seen, while the magenta stained regions are shown in strong contrast. Scale bar = 50 ^m. From ref. 1 with permission.

1: Basic optional microscope

Figure 6. Signifance of the zero order light in image formation: dark-field image of P. angulatum. Objective and specimen as in Figure 3. Condenser this time illuminating with a wide diameter annulus instead of a central disc, (a) The b.f.p. of the objective with its iris fully open to NA 1.3 accepts and allows through the zero order light together with the (six) first order diffracted light (rings intersecting in the centre to pass through, (c) The image is a bright-field image. (b)The iris in the objective occludes the zero order preventing it from contributing to image formation, (d) The image is seen to be one of reversed contrast.

Protocol 6a. To achieve contrast by the use of dark ground illumination: low power and high power systems Equipment Transmitted light microscope

Simple patch stop

Method 1. Set up Kohler illumination and observe a slide on which there are a number of discretely separated features such as a diatoms or in a plankton preparation. 19

A. J. Lacey Protocol 6a. Continued 2. Recalling that you adjusted the aperture iris to give about 66% illumination of the b.f.p. of the objective, examine the physical condenser iris and note the size of the opening. 3. Prepare a disc of transparent material of the size of the filter housing of the condenser. On to the centre of this disc place a disc of opaque material of the same size or slightly larger than the physical opening of the aperture diaphragm. 4. Place centrally in the light path as near the IAD as possible. 5. Re-examining the specimen you will probably discover that nearly everything is black! Adjust the focus of your condenser slightly upwards and open the IAD. This should enable the diffracting features of the specimen to become brightly visible on a background of minimum intensity.

Protocol 6b. Method for high power dark ground Equipment Universal condenser or specialist dark ground condenser Specimen such as Pleurosigma angulatum as illustrated in Figure 6

Immersion oil Objective with a variable diaphragm in its b.f.p. (usually the NA is given as variable in this type of objective)

Method 1. Set the specimen as for bright-field conditions and note its location by reference to stage settings. 2. Remove the specimen and then for universal condensers place a drop of immersion oil on the top lens of the condenser and also another drop on to the back of the slide in a position approximating to the position of the specimen. 3. Carefully replace the slide on the stage allowing the two drops of oil to merge from their centres outwards thereby eliminating any air bubbles. 4. Recover the location of the specimen by stage controls. 5. For the oil immersion objective now place a drop of oil over the specimen. Bring the objective into focus if necessary observing from the side of the microscope when oil contact is made. 6. Having focused on the specimen which will be in bright-field conditions if the objective iris is open, carefully close down the iris in the objective and the image will become one of reversed contrast. 20

1: Basic optical microscopy Note: careful adjustment of the settings of the iris in the objective will change the contrast from dark-field and back to bright-field again. It is possible by observing the b.f.p. with a telescope with these same changes of the objective iris to see that there is a relationship of the image contrast with the presence or absence of the zero order (illuminating) light. If it does not contribute to image formation then the image is of reversed contrast on a dark dark-field (see Figure 7). 7. The oil should be carefully wiped off with lens tissue from all the surfaces at the end of the viewing session.

For dry systems there are special condensers available which can be inserted once bright-field Kohler illumination has been obtained. No oil is used on the condenser in these cases.

Figure 7. Diffraction in the phase-contrast microscope, (a) Specimen of grating (phase) with line spacing about 10 |j.m. (b) b.f.p. of the phase objective NA 0.25 showing zero order light (0) superimposed on the darker phase plate and the other orders of diffracted light (1 and 2) passing partly through the phase plate but mostly inside and outside of it.

21

A. J. Lacey Protocol 7. To set up phase-contrast (for examining living unstained material) Equipment Phase-contrast objectives and condenser Phase telescope High and low contrast specimens such as in Protocol 2

Green filter Test specimen having a simple periodicity such as a diffraction grating

Method 1. Set up Kbhler illumination as for Protocol 2. 2. Remove eyepiece again and insert telescope focused on the b.f.p. Note that the phase-contrast objective has a phase plate carrying a dark ring (the phase ring) fitted in the b.f.p. The dark appearance is due to there being an absorbing layer in the ring to reduce the relative intensity of the annulus light when passing through it. 3. After replacing eyepiece, observe a specimen of cheek epithelial cells. Note the very poor contrast with normal Kohler illumination. You may have to reduce the illuminating diaphragm even to find them. When you have done so reopen the illuminating aperture fully. 4. Swing in the appropriate sized annulus in the condenser (this may be in the slider supplied with low priced phase condensers, or in a rotatable disc in universal condensers). The bright annulus will be of the right size if the condenser is properly focused but it may not coincide laterally with the dark ring in the b.f.p. 5. Adjust the centration of the bright annulus to fit exactly into the phase ring in the b.f.p. 6. Replace eyepiece and place green filter in the light path. 7. Note the unstained materials (or very weakly stained specimens) have greatly improved contrast. Particularly clear are the nuclei in the cheek and onion cells. They will appear dark green (in positive phase systems) because of their increased optical path over the cytoplasm. Granules in the cytoplasm may also be dark. There is a halo around each feature. This halo is an artefact of the phase-contrast system. 8. Changing to higher objectives repeat steps 4-6. Most microscopes are capable of holding their centration for a considerable number of changes of objectives and their condenser matching annuli but it is worth checking their alignment from time to time.

An experiment to illustrate the working of the phase-contrast microscope is as follows. Replace the living specimens with the diffraction grating and 22

1: Basic optical microscopy observe the b.f.p. of the objective with the telescope. Note the appearance of the central ring of light (the image of the annulus) overlying the phase ring as in the setting up procedure but there are also other rings of light as in Figure 7. These are the first order diffraction images of the annulus. Notice that they largely pass through the b.f.p. outside or inside the phase ring. Only a small portion of each passes through the phase ring itself. It is important in the functioning of the phase-contrast microscope that the light scattered by the specimen passes along a separate light path from the zero order (the central ring of light). The specimen has altered the scattered light by about a quarter of a wavelength (of green light) relative to the unscattered and the phase ring puts in another quarter wave difference thereby producing a half-wavelength in total which will give rise to interference and so an amplitude range in the image. Note: the optical path differences (o.p.d.) are proportional to the product of the thickness (t) of the specimen in the z direction and the difference in refractive index of the specimen (no) to that of the medium (nm) in which it situated (no - nm), i.e. o.p.d. = t (no - nm). The phase-contrast image is a map of the o.p.d. of a specimen defined by the boundaries where changes of o.p.d. produce accentuating haloes.

8.3 Phase-contrast Phase-contrast is described in Protocol 7 and the appearance of the b.f.p. of the objective when being used to view a grating is given in Figure 7. It is seen that the zero order light (the bright annulus) is being reduced in intensity relative to the scattered light (in this case the first order diffracted light) which is passing outside or inside the phase plate. The zero order light is also by virtue of its passing through the phase plate being relatively changed in phase by a quarter of wavelength of green light. 8.3.1 Polarized light Crossed polars is a very impressive technique for producing contrast in plant sections or bone material where form birefringence (due to orientated molecules or microfibrils) is present. Protocol 8 is a very basic illustration of how to determine the permitted vibration direction of an unknown piece of Polaroid. Protocol 8. Determination of permitted vibration direction Equipment Pieces of Polaroid Preferably a polarized light microscope with a rotating stage and slots (but sufficient for this protocol would be two pieces of Polaroid)

23

(About) 20 x 10 cm of glass 3 mm or more thick High intensity light source Section of coniferous wood unstained or very weakly stained

A. J. Lacey Protocol 8.

Continued

A. To determine the permitted vibration direction of a piece of unknown Polaroid from first principles Principle: when an unpolarized beam of light is reflected off a transparent surface the reflected beam is plane polarized and its vibration direction is parallel to that of the surface of the reflecting material and at right angles to the ray direction (see Figure 8). 1. Place the glass on a dark bench surface some distance in front of the light source in such a way as to allow the reflection of the light beam to enter your eye (Figure 8). 2. View the light reflection through the piece of Polaroid. 3. Rotate the Polaroid slowly noting the change of intensity perceived. 4. Obtain the orientation of the Polaroid when the intensity of the beam is at a maximum. 5. Conclude from step 4 that the permitted vibration direction of the Polaroid is now parallel to the glass surface. 6. Check by observing the orientation of the Polaroid when the perceived beam is at a minimum intensity. 7. Conclude from step 6 that the permitted vibration direction is now at right angles to the glass surface. 8. Mark your Polaroid in one corner with an appropriate symbol. B. To obtain contrast in an unstained or very weakly stained radial longitudinal section of conifer wood, between crossed polars 1. Set up Kohler illumination in the usual way. 2. Put in the polarizer usually below the condenser (in a polarized light microscope it is usually on a rotatable tray marked with degrees and a reference notch in the carrier). Rotate the polarizer to put 0°. This may be effectively putting the vibration direction as determined above in an east-west orientation. 3. With no specimen on the stage place the second piece of Polaroid (now called the analyser) in the light path above the objective, either in a purpose built slider or in a simple case as a cap over the eyepiece. 4. Rotate the analyser noting the change of intensity. At minimum intensity the two pieces of Polaroid are at crossed polars position. The background should be as near black as possible. Note the position. Then uncross them and insert the specimen to be examined. 5. Using the polarizer only (or having the two polarizers parallel) the specimen can be rotated to see if there is any change of colour or intensity in its features (if there is change then suspect pleochroism). In conifer wood there is little pleochroism.

24

1: Basic optical microscopy 6. Re-introduce the analyser at the crossed polars position and observe the changes of intensity pattern in the background and in the specimen (see Figure 8). If possible rotate the specimen slowly and note the further changes. The black crosses in the bordered pits are known as isogyres and their presence suggest that the pits are spherulitic in structure. 7. Conclude the image is a map of the degree of birefringence (more accurately the optical path, i.e. the product of the thickness in the z direction multiplied by the birefringence) and its orientation. 8. Further study of the image when the sensitive tint plate is inserted in the slot of the purpose built microscope will provide additional colour contrast. The protocol goes on to show a simple crossed polars and rotation technique for achieving contrast in a section of coniferous wood (Figure 8). Further use of polarized light is discussed in Chapters 3 and 11.

8.4 Differential interference contrast (DIC) DIC techniques use crossed polars and Protocol 9 sets out a technique for examining an unstained biological specimen using a transmitted light microscope fitted with DIC optics. DIC is further utilized in Chapter 3, Section 2.5.6. Protocol 9.

Differential interference contrast (DIC)

Equipment Specimens such as cheek cells taken from the side of the cheek by gentle pressure from a clean finger or wooden spatula and mounted in saliva then covered with coverglass are good material, or an onion skin preparation

Microscope fitted with plan objective lenses and carrying a set of Wollaston prisms appropriate for the objectives and the condenser Polarizer and analyser

Method 1. Read the instruction manual of the microscope and check the presence of the parts required. 2. Set up Kohler illumination with the condenser set to normal brightfield conditions. You may notice the symbol on the condenser—as you sort the parts out in the accessory box—indicating the direction of shear. 3. Insert the polarizer below the condenser. 4. Locate the cells perhaps by reference to the air bubbles in the saliva and again temporarily closing the illuminating aperture diaphragm to improve contrast.

25

A. J. Lacey Protocol 9.

Continued

5. Rotate in the appropriate setting (Wollaston prism) in the condenser to match the objective in use. 6. Remove the eyepiece to observe the b.f.p. of the objective. 7. Cross the polars by inserting the analyser and by tuning the upper Wollaston prism to obtain the minimum intensity band in the b.f.p. Close down the aperture iris to a little larger than the dark band. 8. Replace the eyepiece and note the appearance of the image. The cells will show a 3D quality and the direction of lighting will be apparent. This is the direction of shear referrred to in step 2. 9. Adjust the second Wollaston prism while you watch the image and note that you can change the apparent direction of lighting and even experience a change in the perception of hills and valleys. 10. Infer, and it is important to do so, that the image you see is a result of the settings on the microscope. 11. The settings on the microscope in this way displays a contrast image of the optical path variations present within the specimen and between it and the mounting medium. Each change of optical path is represented by an edge effect giving the image the 3D impression. 12. Colour can be obtained (in some microscopes) by referring back to step 7 and tuning the prism to obtain a coloured band or either side of the zero order grey.

Note: the onion cells may show noticeable crossed polar effects in the vertical walls which will show as birefringent brightness changing with orientation. See Chapter 3, page 120 for further information on DIC.

8.5 Fluorescence microscopy This is a powerful contrast technique often using fluorochromes at such low concentrations that living biological material is unaffected by their presence. Much more information and several protocols are given in Chapters 2,4, and 6. Figure 8. Polarized light microscopy, (a) Method for determining the permitted vibration direction of a piece of polaroid (see Protocol 8A). (b) Appearance of an unstained radial longitudinal section of coniferous wood (placed at 45°) viewed between crossed polars. Spherulitic structures are the bordered pits in the tracheid horizontal walls. Brightness in general terms is an indication of the highly birefringent area of the specimen. Photomicrograph on Film HP4, normal contrast development. Nikon objective x 40, NA 0.65. Bar = 100 urn. (c) Same section as in (b) but viewed with x 44 Spencer apochromatic objective, x 6 compensating eyepiece, recorded with Sony camcorder TR805 using Hi 8 SP, and video printed on a Sony Mavigraph video printer. Tracheid width = 0.40 (im.

26

1: Basic optional microscopy

27

A. J. Lacey

Figure 9. Living onion epidermal cells in different contrast techniques, (a) DIC shear direction NE-SW. (b) DIC shear direction NW-SE parallel to the long (c) Phase-contrast of same field as (a), (d) Jamin-Lebedeff interference double Baker-Smith microscope with X 40 objective, (e) Dark-field transmitted light bar = 100 n.m). (f) Dark-field incident light.

image walls, focus (scale

8.6 Provisional summary of contrast techniques The contrast techniques given above are largely those produced in the microscope techniques themselves. The use of stains and fluorescent probes are portraying maps of the chemical affinities in the specimen. Light is differen28

1: Basic optical microscopy tially absorbed according to wavelength and transmitted or the energy is reemitted at a longer wavelength. The phase-contrast and the DIC techniques are giving contrast to changes in path (optical path) of the light passing through the specimen in the z direction. Polarized light is giving a map of the different absorption properties of the material (pleochroism) or in the case of crossed polars technique a map of the optical path—birefringent portions, their thickness in the z direction, and their orientation. The removal of the background intensity in dark-field microscopy and the objective lens ability to collect the light scattered by sharp changes of refractive index in the specimen can bring about reversed contrast in the image of that specimen. Quantitative techniques are available for measuring optical paths by phasecontrast and by Jamin-Lebedeff interference techniques. Crossed polars microscopy is very powerful in measuring refractive indices and direction of birefringence but biologists have not advanced as far as mineralogists in their utilization of this technique. See Chapter 7 for reflection-contrast techniques. Confocal microscopy (see Chapter 2) is basically a technique for preventing unwanted light from contributing to image formation. Video cameras and associated computer software programs for image reconstruction (Chapter 3) also improve upon the basic microscopy contrast techniques by subtracting, for example, the effects of unwanted light or out-of-focus images from the image which has been formed in the microscope. Video-intensified microscopy (VIM) are helpful in very low light level contexts. Chapter 3, Figure 1 indicates how the conventional techniques so far described can be extended by confocal and video microscopy in the visualization of biological materials.

8.7 Summary of contrast techniques The contrast in the image can be manipulated by preliminary treatment of the specimen such as by sectioning and or staining activities which may seriously alter the specimen. Techniques for achieving contrast in living material are possible by the careful control of the light passing through the back focal plane of the objective (phase-contrast) or by comparing two beams made to illuminate the specimen and then being recombined in interference techniques. The image is the result of interaction of light with the specimen. Resolving power of the microscope is achieved by the ability of the objective to collect the diffracted light. A very useful exercise is to examine the same unstained specimen with a series of contrast techniques (see Protocol 12).

9. Recording the image The recording of the appearance of the image can be achieved by hand-drawing which improves with practice! There are some drawing optical aids which allow the image to be seen at the same time as a pencil line drawn on a sheet of paper. These have value if the dimensions of a feature are required as a drawing of the image of a stage micrometer can be prepared with the same 29

A. J. Lacey

optics. However photomicrographic techniques are widely available and instruction manuals for the equipment are in the main user-friendly.

9.1 Microscope/camera attachments Both traditional photomicrographic and video camera recording techniques require careful attention to the interface between the camera and the microscope to achieve parfocality of the microscope image plane with the plane of the emulsion of the film or the chip of the video camera. When the microscope is used for visual examination of the specimen the rays of light leaving the exit pupil of the microscope are almost parallel and the eye is receiving them in a relaxed state (i.e. focused almost at infinity). The image in visual microscopy is said to be a virtual image. Figure 10 illustrates the options that

Figure 10. Images in the eye and for photographic and video camera recording, (a) Image formation with the eye. The microscope is adjusted so that the primary image falls on the front focal plane of the eye lens in the eyepiece. This means that parallel rays of light will enter the eye and be focused by it on to the retina. (b) The primary image is lowered by increasing the specimen-objective distance. This causes rays leaving the eyepiece to converge and form a real image in a plane suitable for accommodating the film. But performance of the objective is thereby impaired, (c) Specimen and primary image are in the correct positions, but the eyepiece has been raised to increase primary image-eye lens distance and thereby produce a real image on the film. (d) Special projective lens or 'photographic eyepiece' designed to form a real image on the film using the normal settings of the microscope. (e) Normal eyepiece and in conjunction with a converging lens in a special photomicrographic attachment; optically similar to using the eye (a). From ref. 1 with permission.

30

1: Basic optical microscopy are available for altering the virtual image to make it a real image falling on the film or chip. The photomicrographic accessories commonly available with microscopes (or built-in photomicrographic systems) have auxiliary lenses to do this. You should check the equipment you have available for such accessories as projection 'eyepiece', projection lens, or other lenses built-in. A check can sometimes be made on the photo equipment by opening the empty camera and placing a ground glass in the plane of the emulsion when an image will be seen. Most microscope manufacturers provide the required adaptors for attaching a range of video cameras to their instruments. The video camera is usually fitted with a threaded 'C' lens mount or the bayonet ENG lens mount. Some manufacturers publish a document specifically on the microscope camera attachments (see for example Olympus 'Video/Photo adapters'). Space precludes a lengthy treatment of this subject. An important piece of the final print is that it contains a scale bar and this is given emphasis. Other chapters in the book will contain photomicrographic records and the details of the optics used to take them (see for example Chapter 4, Section 6.3). Kohler illumination is again important to ensure that the specimen is lit in a controlled manner, and that the lighting of the background is without distracting features. Kohler was himself a photomicrographer and developed the method of illumination precisely to achieve this control of the light. A single protocol is given here for photomicrography and particularly to illustrate the use of colour filters for black and white pictures (Figure 5) or for the adjustment of colour temperature for colour transparencies. Protocol 10. To make a photomicrograph using (a built-in) 35 mm camera system Equipment • Microscope to which is fitted a camera systern with appropriate microscope/camera interfacing • Photomicrograph recording sheet • Range of neutral density and colour filters

• 35 mm film of known characteristics (i.e. B&W or colour, transparency, daylight/ tungsten light, speed) Colour contrasty specimen

Method 1. Select the film according to the requirements envisaged (see Chapter 4). 2. Check the camera for dust and load the film. 3. Advance the film wind on mechanism checking where possible that the camera rewind knob is rotating to indicate that the film is being transported effectively. 4. Set the film speed (previously verified by experiment) on the control panel on the exposure measuring device. 31

A. J. Lacey Protocol 10. Continued 5. Prepare the record sheet for notes on each exposure. 6. Clean the specimen slide thoroughly. 7. Set up Kohler illumination, selecting objective and approximate field of view of your specimen. Check image is free of blemishes caused by dust in the system. 8. Where necessary ensure that the beam splitting prism in the eyepiece head of the microscope is directing the light to the camera. Observe the format lines in the new image and adjust the field of view of your specimen to the 35 mm format. 9. Defocus the specimen's image and with the eyepieces' focusing device critically focus the cross lines (usually seen as double). 10. Return the specimen image to critical clarity by the microscope focusing system. This procedure has focused the image on the graticule and on the emulsion of the film. 11. If using colour film adjust the lamp voltage to the predetermined best setting (colour temperature). If necessary, insert any correction filters (i.e. Wratten 80A to compensate for tungsten/halogen lighting when using daylight film). 12. Decide whether the exposure meter is required to read the whole field of view and thereby obtain an average value or, as in the case of dark ground or fluorescence images, there should be a spot metering. Read the exposure meter and check that it falls in the range of the of the equipment. 13. If indicated that the exposure is too short insert neutral density filters. If indicating too long increase the illumination intensity or use faster film possible up to ASA 3200. 14. Again check precision of focus. 15. Press the shutter release avoiding touching any part of the bench or machine while the shutter is open. 16. Check wind on is working when shutter is closed. 17. Note the details of the optics and specimen on the record sheet. 18. If the specimen is a very important one it may well be useful for a series of modifications and subsequent exposures made at this point (see for appraisal in step 22). 19. (Optional but valuable.) Take the next exposure with the specimen replaced by a stage micrometer, keeping all optics as before. 20. For black and white films a series of pictures taken of the stained contrasty specimen can be photographed using complementary or same colour filters (see for example Figure 5). There may be considerable

32

1: Basic optical microscopy changes of exposure necessary (made by the camera control system if automatic) when combinations of filters are changed. 21. At the end of the film, rewind into the cassette. It is probably false economy to wait for long periods for the film to be entirely used up. It is better to process promptly even if a few exposures are wasted. Records may be lost, film may not be appropriate for the next occasion, someone else may have taken the film out of the camera! Record books are often used in laboratories but even these may be incomplete, for example they may be without the initials of the last user. 22. Critically appraise your pictures looking for excessive warmth (too much red) or too cool (too much blue) in your colour transparencies. Uneven lighting will show if Kohler illumination has been poorly setup, dust in the system will show up. Inaccurate focusing and also inappropriate combinations of magnification and numerical aperture of the objective used resulting in lack of resolution of detail or inadequate depth of focus, all may lead to frustration. Inappropriate section thickness for the depth of field of the objective may also contribute to dissatisfaction.

Protocol 11. To obtain records of an image using a video camera and video printer, video cassette recorder (Figure 11) Equipment • Microscope with camera port preferably with a trinocular head • Video camera • Interface accessory for camera to microscope (so that the image of the specimen falls on the chip of the camera)

• Video printer with appropriate paper (B&W or colour) • TV monitor • Stage micrometer . video cassette recorder (VCR)

A. Using a video camera and video printer 1. Fit camera to microscope through appropriate interface. Connect camera video output to video printer input. Connect video printer output to monitor input. Where the TV monitor has a video input impedance termination switch this should be set to the 75 ohms position. Check paper in cartridge of video printer. 2. Set up Kohler illumination and observe specimen through the eyepieces. 3. Insert beam splitting prism to direct light to camera. 33

A. J. Lacey

Protocol 11.

Continued

4. Observe monitor. Light intensity will almost certainly have to be reduced and do this by insertion of neutral density filters. Heat filters might also be required to avoid burn out of image on monitor. 5. Adjust field of view portrayed by the monitor and critically adjust focus. 6. The frame/field switch would normally be set to 'frame' so that all the lines of the TV picture are recorded. The 'field' setting produces a picture in which only alternate lines are printed, the scanning time been reduced from 40 to 20 milliseconds. This helps to avoid distortion or blurring of moving objects in the image. Press print button. 7. Note settings on video printer for such as normal, dark, light (B&W printers). The printer's instruction book will help in deciding if there is need for altering the ex-factory settings after viewing the printed image. 8. Repeat the procedure for the image of a stage micrometer using the same optical system. From the print thus obtained place a scale bar by hand on to the print obtained in step 7. 9. Some printers have sub-menus for insertion on to the print of details of the specimen, optics, etc. This information can be added at step 6. 10. Make photocopies of the print showing the scale bar. These, in some cases, keep more satisfactorily than the prints themselves. B. Using a video cassette recorder 1. Connect the camera output to the input of the VCR. Connect the ouput of the VCR to the input of the video printer retaining the monitor connections as before. 2. Audio signals can be fed into the VCR either at the time or in a subsequent viewing of the recorded tape. Domestic VCRs may or may not have a microphone input. If there is no microphone input on the VCR then a microphone amplifier is required as an external accessory found in video processors for editing. 3. Again images of the stage micrometer should be recorded. 4. Sequences of moving objects can be recorded. Take care not to have large changes of field of view made too rapidly. Again avoid too long recordings of situations as viewing subsequently becomes exceedingly tedious. Audio record changes of objective, field, specimen changes, as well as date and name of operator. 5. Prints can be made from the stored images on the tape by switching off the camera and using the PLAY mode on the VCR. Use the appropriate stage micrometer images to insert scale bars on the print as described in part A, step 8. 34

Figure 11. Diagrammatic flow of the information from the light source to the final print (and tape record) in a video printer system. The print is of a specimen of human chromosomes G-banded with trypsin Giemsa stain, courtesy of Mr A. Cuthbert and Dr D. Trott, using microscope Olympus BH2, objective A40PL, NA 0.65, Hitachi CCD KP113 camera, and Mitsubishi Video Copy Processor, paper K75HM. The inset of the wave form is reproduced from ref. 2 with permission.

A. J. Lacey Protocol 12a. To make a colour saturation test specimena Equipment • Microscope preferably with trinocular head and appropriate camera microscope interface • Colour video camera • Colour video printer

• Colour monitor . Film base from a processed Polaroid 'Polachrome' 35 mm film Adhesive tape • Mounting medium such as DPX

Method 1. Take the scrap piece of unprocessed film left at the beginning of a used Polachrome film after processing. 2. Roll the adhesive tape down firmly on to the black side of the film. 3. Peel off the adhesive tape, the emulsion will come away on the tape, leaving the film base with the printed lines on the upper surface. 4. Without turning the film over, mount a small piece of this film on a slide under a coverslip. It is essential that the printed lines are on the upper side of the film, immediately below the coverslip. (The lines are on the less glossy side of the film.) Note: for reflected light the specimen can be be vacuum aluminium coated on the non-emulsion, more glossy side, before mounting on the slide, without a coverslip. aThe method for the test specimen is kindly supplied by D. J. Thomson.

Protocol 12b. To test colour saturation of video camera/video printer, video monitor Equipmenta • As for Protocol 11 but with colour camera, colour video printer, and colour monitor

Method 1. Observe the image of the bands with light set at the correct running voltage. 2. Note fully saturated red, blue, and green bands through the eyepieces. Any yellow colour at the junction boundaries of the red and green bands is an artefact produced by an error in the system such as uncorrected chromatic aberration in the microscope (or camera optics). 3. Send the image to the colour printer and monitor. 4. Use neutral density filters to correct light intensity. White balance is

36

7: Basic optical microscopy

automatic in most cameras and will compensate for variations in colour temperature allowing the intensity of the light to be controlled also by the voltage setting on the microscope. Some cameras have a 'high speed shutter' which can be switched in for moving material. 5. Enter sub-menu details of optics, etc. 6. Print. Repeat with specimen replaced by stage micrometer if scale bar is required to be put on to final print. 7. Visually compare the print with the eyepiece image and/or with a con ventional 35 mm photomiurographic image record (obtained perhaps by Protocol 10). Rank your satisfaction level. The exposure times are automatically determined by the camera factory sellings but by careful consultation of the instruction manual can be overridden.

9.2 Summary (l should he remembered that the specimen-light interaction which is por[rayed in the microscope image is changed by a photochemical process in the photographic system into silver grains or dye particles impressed in paper.

Figure 12. Video print of a 'Polachrome'35 mm film base (see Protocol 11). Olympus BX 50 microscope, UPLANFI, •/-,'. 17, \ 40/0.75 objective, JVC colour video camera TK-128OE, Sony Mavigraph UP 1200 EPM video printer. Width of green band 7/jun.

37

A. J. Lacey The changes in the chemistry are then chemically fixed. This pattern of change is then viewed by the observer's eyes in light reflecting prints or light transmitted in transparencies. In the case of the video process the image is changed into a pattern of electronic charges which are read directly, in an analogue system, or translated into a digitized format in the digital camera or digitizer. The signals are then changed back into light and darkness patterns by a further process of electrons interacting with phosphors in the TV tube. The response in these phosphors is what the eyes see and interpret. Or alternatively the electrons cause a change in reactive paper or cause sublimation of dyes onto paper—the video print. The resulting patterns on the print are again interpreted as patterns of light and dark (or colour) seen by the eye. The relative ease of obtaining a print from the video printer technique whether direct from the camera or from a previously recorded image is very attractive. The economics of the video system compared to that of dark-room aided photography must be a factor in deciding which type of situation a laboratory might choose. The archiving of prints is probably much the same in both cases. The use of cine film for recording moving images is now virtually obsolete. The transfer, however, of achival cine records to video tape, best done commercially, is very satisfactory.

Protocol 13. Preparation of onion skin peels for comparison of contrast techniques (5 with permission) Equipment • Inner bulb scales from fresh onions • 0.5 mm plastic sheeting (easily obtained from empty storage bottles for non-toxic chemicals) • Double-sided adhesive tape (Agar Scientific)

• 1 cm cork borer • Scalpel and artist's water colour paint brush . A range of microscope contrast techniques available preferably on the same machine

A. Preparation of onion skin peels

1. Cut the plastic sheeting into 15 mm X 15 mm pieces. Remove a central inner core with the cork borer, leaving a plastic holder with a hollow centre. 2. Firmly secure one side of the holder to double-sided adhesive tape, trim the tape flush with the edges of the holder, and remove the tape covering the central hollow by cutting round the edge of the hole with a sharp scalpel. 3. Obtain the inner bulb scale of the onion avoiding wetting the surface of the inner epidermis with sap. 4. After removing the remaining protective layer of the adhesive tape, press the plastic holder firmly against the inner epidermis of the scale. 38

1: Basic optical microscopy Cut the epidermis with the scalpel around the outer margin of the holder. 5. Lift the holder off which will remove the epidermis adhering to the plastic holder. Handling the holders by the edge with forceps will protect the epidermis from damage. 6. Store the specimens in water in a Petri dish until required. Any gelatinous cell wall material adhering to the side of the epidermis in contact with the water may be wiped off with the soft paint brush. 7. Mount a holder in water on to a slide. Cover with a glass. Go to contrast techniques. B. For short-term preparations 1. Isolate by a series of shallow cuts a piece of the inner epidermis of an inner scale of an onion bulb. Grip one edge with forceps and strip from the scale. 2. Mount in water on a slide, taking care to keep the outer surface of the skin upwards in order to maintain its orientation to the light and to the eye. Cover with a coverglass. This preparation will last for a considerable time particularly if kept moist and cool. C. Contrast techniques 1. Examine the specimens under each type of contrast technique including transmitted and incident light for bright-field, dark-field, phasecontrast, polarized light (single polarizer and crossed polars), DIC, and epifluorescence (auto and with fluorochromes). Figure 9 shows the appearance of the epidermis under five of these contrast techniques. 2. Prepare a chart such as in Table 2 giving sufficient space for comments on the image of the various techniques utilized. 3. Enter into the chart the levels of brightness or relative intensity of the various features and their backgound. Remember to enter equal brightness or darkness if there is no contrast between features. 4. You may wish to amend the entries given in the Table 2 but be prepared to think through why you disagree with what is entered. Notes: for quantitative data on such features as the mass of the nucleus or the refractive index of the cytoplasm then immersion refractometry using phase-contrast or Jamin-Lebedeff interference microscopy could be tried. Relative dimensions and volume can be obtained by reference to techniques described in Chapter 8. Plasmolysis of the cytoplasm can be achieved by immersing the epidermis in 2% sea salt in water which has a higher osmotic pressure than the vacuolar sap.

39

Table 2. Onion skin examined with various microscope techniques Technique

Background

Onion skin components (see Figure 13) Nucleus

Cytoplasm

(%.13/4 and 4a)

(Fig. 13/5 and 61

Light-matter interaction Walls

Vacuolea,b Horizontal Top

Bottom

Vertical Short

Long

Transmitted Bright-field

Bright

Only visible with aperture closed down (rotten image throughout)

Dark-field

Dark

Sky grey

Phase(+ve) green filter

Bright green

DIC

Bright

Polarized light Bright (plane)

Granules sky grey or bright

Bright in all orientations

Scattering by reflection and refraction Boundaries of refractive index

Dark with halo Dark granules with haloes round edge Movement

Bright Bright (on either side of dark line)

Map of optical path in z direction = t(no - nm)

Visible (3D effect)

Bright Bright Map of optical path as above (but (but see under polarized light) with boundaries accentuated by 3D effect)

Granules clearly visible Movement

As bright-field

Polarized light Dark (plane) with crossed polars Crossed polars Red one colour + sensitive tint

Diffraction effects and absorption map

Very slight white

No change on rotation of specimen

No pleochroism apparent

Bright Bright (at certain orientations only)

Map of birefringence, o.p.d. and orientation =t(n11- n1)

Blue (or yellow)

Indicate direction of birefringence ±(n11-n1)

Yellow (or blue)

AVEC-DIC

Blue (or yellow)

Transvacuolar strands and ERd

Yellow (or blue)

See DIC above but with enhanced contrast Indicates direction of birefringence ± (n11-n1)

Fluorescence see under incident fluorescence Confocalc Incident Bright-field

Bright

with surface replica and sputter coating Dark-field Dark

Some brightness but excessive Visible(?) Map of reflectiveness glare Negative or positive of surface characteristics depending on single or double replication Clear contour of brightness Grades into horizontal walls Map of surface light scattering features Dark channels in wall clear

Fluorescence Primary Dark Fluorescence Acridine Orange Dark Fluoroprobes Dark

No primary fluorescence apparent

Apple green

(If over-stained yellow green) Map of chemical affinity for Acridine Orange

Various colours depending on probes and irradiating wavelengths (Vacuole visible with LYCHb)

Confocalc aThe vacuole can also be inferred by achieving plasmolysis when mounting medium such as 2% sea salt in water has a higher osmotic pressure than the cell sap. b Oparka et al. (4) has described the use of LYCH on onion preps. c Confocal microscopy is clearly potentially very powerful for optically sectioning living onion cells. d Video microscopy using VEC and VIM (3).

A. J. Lacey

42

1: Basic optical microscopy Figure 13. Diagrammatic three-dimensional appearance of onion epidermal cell in its relation to the light in a microscope and the observer's eye. (a) How the cells appear seen from the top (x and y directions) (see Figure 9). 1. Vertical walls in the (z) axial plane (a) short and (b) long. 2. Horizontal wall in the plane of the stage of the microscope. Upper wall, (a) upper surface (b) inner surface. 3. Lower horizontal wall, (a) inside cell surface (b) outer. 4. Nucleus with (4a) nucleolus. 5. Cytoplasm with granules (with insert courtesy of Dr N. Allen and ref. 3 with permission sought). 6. Vacuole.

Figure 13 gives a three-dimensional impression of what the epidermal cells can be interpreted as being like after examination with a wide range of techniques. The scientific literature is adding information continuously to this impression of the physicochemistry of onion epidermal cells. Video microscopy from the laboratory of Professor Allen (3) and fluorochrome studies by Drs Oparka and others (4) have greatly expanded the information on the vacuole and its contents.

Acknowledgements Inevitably in this introductory chapter or summary of basic microscopy there are a number of points which were derived from colleagues and teachers over the years and their contribution is hereby gratefully recognized. Especial thanks for his reading the whole script and amending the text and the protocols on the video recording go to Mr D. J. Thomson. Thanks too to Dr Peter Evennett for permitting his figures to illustrate much of the chapter. Thanks are due for the protocol on preparation of the onion epidermis given by Dr Oparka and for the illustration of the details of the endoplasm to Dr Nina Allen. Colleagues Drs Cuthbert, Trott, and Jobling have kindly allowed the author access to their video printers.

References 1. Evennett, P. J. (1989). In Light microscopy in biology: a practical approach (ed. A. J. Lacey), p. 61. IRL Press, Oxford. 2. Trundle, E. (1987). Newnes television and video engineer's pocket book. Butterworth Heinemann, Oxford. 3. Allen, N. S. and Brown, D. T. (1988). Cell Motil. Cytoskel, 10,153. 4. Oparka, K. J., Murant, E. A., Wright, K. M., Prior, D. and Harris, N. (1991). J. Cell Sci., 99, 557. 5. Oparka, K. J. and Read, N. D. (1994). In Plant cell biology: a practical approach (ed. N. Harris and K. J. Oparka), p. 27. IRL /OUP Press, Oxford.

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2 Introduction to confocal microscopy P. J. SHAW

1. Introduction Optical microscopy is inherently limited in resolution by the wavelength of the light used in the technique; in practice this resolution limit is generally about 0.25 jjim. However, there are several factors which make optical methods very powerful. One major advantage comes from the combination of microscopy with fluorescent probes. Almost any biological component can be specifically labelled with a fluorescent tag, sometimes called a fluorochrome, and then imaged by virtue of its fluorescent light emission. The available fluorescent probes include dyes for specific components, such as nucleic acids, reporters for particular ions such as calcium, modified DNA and RNA sequences, which are used for fluorescent in situ hybridization, and antibodies raised against almost any biological component imaginable. Fluorescent detection has the advantage of great specificity and sensitivity. Optical filters are used to provide light of just the wavelength needed to excite the fluorochrome, and a dichroic mirror and an emission filter can then be used to provide an image arising just from the fluorescently emitted light. This has the effect of a high degree of discrimination against the illuminating wavelength and in favour of the fluorescent tag, resulting in a very good signal-to-background ratio. This selectivity is even greater in the nearly universal epi-illumination configuration; the imaging objective lens is also the illuminating condenser lens, and so in this arrangement the illuminating light which has not undergone absorption passes through the specimen and cannot contribute to the image background. A second major advantage is that light is relatively non-destructive to biological material, and so with care and skill optical microscopy can be used to image living cells, tissues, and organisms without too much damage. Many studies over the past few years have used fluorescence microscopy in living cells. In some cases fluorescent reporters can be simply perfused into living cells—for example, AM (acetoxymethyl) esters of calcium-sensitive dyes. Cellular esterases cleave the ester linkage, and prevent the dyes from diffusing out again. In many other cases microinjection has had to be used to introduce fluorescent probes. For example, a purified protein can be covalently labelled with a fluorochrome and then microinjected into cells. In favourable cases, the introduced protein will

P. J. Shaw behave in the same way as the native protein, allowing fluorescence microscopy to be used to determine its location and dynamic behaviour within the living cell. Most recently molecular genetic techniques have begun to be used to link green fluorescent protein (GFP)—a naturally fluorescent protein found in a jellyfish—to proteins of interest. A cDNA is constructed containing the target protein and the GFP sequence, under the control of a suitable promoter, and is then transformed either transiently or stably into the cells or organisms of interest. In a surprising number of cases the extra protein domain of the GFP does not seem to interfere with function and location of the target protein, and thus cells can be made which directly express a fluorescent analogue of the protein of interest. This approach is already beginning to bring about a revolution in the cell biological and developmental study of living organisms and cells.

2. The problem of out-of-focus light In spite of its advantages, conventional optical microscopy has some troublesome features. In principle if all the fluorescent light emitted from a specimen could be recorded, it should be possible to reconstruct a perfect map of the distribution of the fluorochromes which emitted the light, at least to the resolution limit specified by the wavelength. In practice this is not the case. The reason lies in the geometry of a practical microscope. The emitted light must be collected by passing through an objective lens, and this of necessity introduces an aperture into the system. The result is that some of the emitted light cannot be recorded, and the image reconstructed by the optical system has the resolution seriously degraded, particularly in the direction of the optical axis (usually denoted z). Furthermore the degradation depends on the level of image detail in a rather complicated way. Thus fine details, often called high spatial frequencies, are relatively little affected, whereas large structures, or low spatial frequencies, are greatly spread out in the z direction. This is often called the problem of 'out-of-focus' light. A small sphere ends up being imaged in three dimensions as a complicated elongated structure. This is discussed in more detail below. Some interesting approaches to this problem attempt to measure more of the sphere of light emission and thus to produce a reconstruction which suffers less from these defects (1), but for the purposes of practical biologists there are two feasible current methods to overcome the problem. The first one is confocal microscopy; the optical system is modified so as to minimize the problem by eliminating or at least reducing the contribution of the out-of-focus light. The second approach is to measure the conventional image accurately, out-of-focus light included, to measure also the properties of the imaging system in detail, and to use image processing techniques to remove the out-of-focus contribution from the image. This chapter will attempt to describe in as non-technical a way as possible the basis of confocal microscopy. There has been much published in recent years about 46

2: Introduction to confocal microscopy confocal microscopy. One of the best sources of information is the second edition of the Handbook of biological confocal microscopy (2).

2.1 The confocal principle: explanation by ray optics The most common way to explain the operation of a confocal microscope is by a ray diagram similar to the one shown in Figure 1. A light source, almost always a laser, is used to provide an effective point source illumination through a pin-hole. At the specimen this gives the diffraction limited image of a point. With a conventional light source this would be the Airy pattern. With a laser beam the focus is somewhat different, but may still be thought of as approximating an Airy pattern. Some of the emitted light which originates from this plane passes back up through the objective lens, through the dichroic mirror and emission filter, and through a second pin-hole, finally being detected by a photomultiplier behind the detector pin-hole. The two pin-holes—illuminating and detecting—are both located in planes conjugate to the plane of focus in

Figure 1. Ray diagram of confocal optical arrangement, showing how light rays originating away from the plane of focus are eliminated from the image by the detector aperture.

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P. /. Shaw the image ('confocal' is derived from a contraction of 'conjugate' and 'focal'). However, the out-of-focus problem arises because parts of the specimen above and below the plane of focus are also illuminated and also emit light. Now consider rays of light originating from above the plane of focus (the light shaded rays in Figure 1). Originating nearer to the objective lens, these rays are brought to a focus further away from the objective on the other side, behind the detector pin-hole. At the level of the detector pin-hole, these rays have not converged sufficiently to pass through the pin-hole, and so they are eliminated from the resulting image. A similar argument applies to light from parts of the specimen below the plane of focus. Therefore the confocal arrangement—the interaction of the illuminating and detecting pin-holes— has the effect of discriminating against light originating away from the plane of focus. So far, we have described confocal imaging of a single point on the specimen. For an imaging system, a method is needed to image successive points to build up an image. This is done in several different ways in different designs, and is one of the main differences between the different types of instrument. This is discussed in more detail later.

2.2 Linear, shift-invariant imaging and the point spread function The explanation given above uses a ray approximation, and while it explains in general terms the exclusion of out-of-focus light, it does not provide a very detailed idea of the image that a confocal microscope will produce. We give next a somewhat better approximation, although still simplified; a full explanation would inevitably be very mathematical. First, we will consider a microscope, confocal or conventional, in system engineering terms as a device which takes an input—the specimen—and transforms it to an output—the image. Now imagine cutting the specimen into two pieces, and imaging each piece separately. If adding these two images together produces the same result as imaging the whole specimen, the process is described as linear; the whole is the sum of the parts. If this condition holds, as it does for many types of imaging, including fluorescence microscopy, then the specimen can be imagined to be cut into smaller and smaller pieces. Ultimately the pieces can be made arbitrarily small and thus can be considered as points. Then the image of the specimen is simply the sum of the images of all the individual points. In principle the image of a single point therefore provides all the information needed to characterize the imaging process. This image of a point is usually called the point spread function (p.s.f.), and mathematically the operation of replacing each point by the point spread function and adding the resulting point images together is called a convolution. In summary, we can describe a linear image formation process as the convolution of the specimen with a point spread function. A further question is whether the 48

2: Introduction to confocal microscopy point spread function is the same for every point in the specimen. If it is the same everywhere the imaging is said to be shift-invariant. Whether this condition holds is not always an easy question to answer. It is certainly a good approximation for fluorescence imaging, but is discussed in more detail below. In fact this explanation is not limited to optical microscopy. It applies to many other types of microscopy, to astronomical imaging through telescopes, to various sorts of spectroscopy, and many other biomedical and physical imaging methods. Thus in conventional fluorescence microscopy each point of the specimen is replaced by a copy of the p.s.f., whose intensity is directly proportional to the intensity of the point. In a confocal microscope, the same argument shows that the illuminating pin-hole, effectively a point source, produces a light distribution at the specimen which is also given by the p.s.f. Light is detected from the specimen only if it passes through the detector pin-hole. In principle this is equivalent to a point detector, although in practice most confocal microscopes allow the detector pin-hole to be opened to admit more light, which causes a departure from true confocal imaging. Since light paths are reversible, this means light is detected from a region of the specimen corresponding to the image of the detector pin-hole at the specimen—again given by the p.s.f. Thus the overall effect is that the p.s.f. is applied twice in confocal imaging—once because of the illuminating pin-hole, and again because of the detector pin-hole. As far as the detected confocal image is concerned, applying the p.s.f. twice means that the effective confocal p.s.f. is the original, conventional p.s.f. squared. Since the p.s.f. comprises a central maximum with lower surrounding maxima, squaring it reinforces the central peak while weighting down the surrounding components. It is the subsidiary maxima that give rise to the out-of-focus contribution in conventional microscopy, and so decreasing their relative weight in the confocal p.s.f. has the effect of removing a large part of the out-of-focus light.

2.3 The shape of the point spread function So far, we have left an explanation of the actual shape of the p.s.f. somewhat vague. There are two ways to determine the form of the p.s.f.: we can derive it from optical theory or we can actually measure it by using a specimen which is as near as possible to a point. A full derivation of the p.s.f. requires physical and mathematical theory which is beyond the scope of this chapter, and in any case probably no theoretician has yet adequately taken account of all the various aberrations and other factors which are necessary to derive an accurate p.s.f. for a real microscope. On the other hand, an outline explanation may be helpful, so we shall give one; some measured p.s.f.s are shown below. Light can be described as waves. A simple, but very useful way of understanding the light originating from an object was first given by Huygens. According to this principle, each point on an object acts as a secondary point source for 49

P. J. Shaw light waves. In the case of fluorescence, each fluorochrome molecule will be such a point source. Thus the total light emission consists of many spherical waves expanding from the points. Within certain limitations, which need not concern us here, this sum of spherical waves can be described as the Fourier transform of the object. In fact, what is usually called the diffraction pattern is actually the intensity of the Fourier transform, and this is a simple way to understand Fourier transforms—a pattern derived from the image where low resolution components, often called low spatial frequencies, are near the origin at the centre of the pattern, and high resolution components, or high spatial frequencies, are furthest away from the centre towards the edges of the pattern. In order to change back the emitted waves to an image, an inverse Fourier transform is needed—this is exactly what an optical lens does. However, not all the light coming from the object enters the lens—there is inevitably an aperture within which light is passed, and outside of which light is excluded from image formation. This aperture, wherever it is actually located in a lens system, may be regarded as an aperture in the diffraction or Fourier transform plane. It is generally called the back focal aperture of the lens. The effective radius of this aperture depends, among other things, on the numerical aperture of the objective lens. Thus the subsequent image made by the lens—the Fourier transform of the diffraction plane—is modified by the aperture. Rather than show the effect of this mathematically, we refer interested readers elsewhere (3), and simply demonstrate this result by a computer simulation. Figure 2A shows a point, corresponding to the original object. The Fourier transform of a point is a flat, uniformly grey plane (Figure 25). We next pass only components within a circular aperture (Figure 2C), and then carry out a second Fourier transform of these components. This gives the pattern shown in Figure 2D. This is a circular central maximum, surrounded by annular regions of light and dark. This is in effect the Airy pattern, and it is produced by the circular aperture mask within the lens system. The three-dimensional p.s.f. is more complicated. The central disc of the Airy pattern becomes weaker away from the plane of focus, and the concentric rings become relatively stronger and increase in diameter. This is shown by images collected from small fluorescent beads in Figure 3A. (See Chapter 6, Protocol2 for a suitable specimen preparation method.) These out-of-focus bead images can be thought of as focal sections through a 3D p.s.f., in which the rings actually comprise a series of concentric cones. This is shown in Figure 3C in which a 3D p.s.f. has been resectioned so as to show a central section parallel to the optical axis (an x-z section). The diverging cones are seen as diagonal bright lines radiating out from the central maximum. The equivalent measured p.s.f. from a confocal microscope is shown in Figure 3B. The main difference is clearly that the subsidiary maxima have been much reduced in comparison to the central maximum as theory predicts. When resectioned as a central x-z section (Figure 3D) the overall pattern is much closer to an elongated ellipsoid, with little contribution from the subsidiary rings. 50

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Figure 2. Computer simulation of the effect of a circular aperture in the diffraction plane on the image of a point object. IA) A single point source is shown. (B) Its Fourier transform is a uniformly grey plane. (C) The finite numerical aperture of the objective lens gives rise to a circular aperture in the Fourier or diffraction plane. (D) The inverse Fourier transform carried out by the optical action of the objective lens then produces the Airy pattern of concentric rings surrounding a central maximum.

2.4 Aberrations and the limits to linear, shift-invariant imaging In the absence of aberrations, the p.s.f. should he circularly symmetric about the z axis, and should have reflection symmetry about the x-y plane, i.e. it should be the same above and below the plane of focus. It is clear that the measured p.s.f. shown in Figure 3A is different above and below the focal plane, and this is due to the presence of aberrations, mostly spherical aberration. It is very common in fluorescence microscopy to observe the presence of 51

P. J. Shaiv

Figure 3. Measured bead images of subresolution fluorescent beads to show the form of the 3D p.s.f, (A) Series of optical sections through a subresolution fluorescent bead, imaged using a cooled CCD camera and conventional fluorescence microscopy. Notice the concentric rings expanding away from the centre away from the in-focus image (which occurs on section nine in the series). The optical sections are spaced 0.4 urn apart. (B) Equivalent series of optical sections through a subresolution fluorescent bead collected by confocal microscopy. The out-of-focus rings are virtually absent. (C) The conventional fluorescence data shown in (A) has been enlarged and resampled in the x-z

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2: Introduction to confocal microscopy spherical aberration. To see this in any fluorescence microscope, simply find a small, bright structure on a fluorescence specimen and observe the way its image changes above and below the plane of focus. Almost invariably, the image will disappear much more quickly on one side of focus that the other. This is true even with high quality, well corrected objectives, and arises because the objectives are usually not used for real specimens under the conditions for which they were designed. For example, oil immersion objectives are usually designed to image objects immediately underneath a coverglass of the correct thickness (usually 0.17 mm), through the immersion oil. Generally, however, the interesting part of the specimen is further away, and so light also passes through the intervening mounting medium (usually water or a glycerol:water mixture) which has a different refractive index from the immersion oil. This refractive index mismatch introduces spherical aberration. It also causes an 'apparent depth' effect, so that the physical changes in the objective or stage position used for focusing are not the same as the changes in the plane of focus—usually the change in focal plane is 10-20% less than the physical focus movement. These effects are minimized if the immersion medium for the objective is the same as the mounting medium, and several companies are now selling water immersion objectives for imaging cells in water-based media. However, this is not a complete answer because cells and cellular constituents themselves have a range of different refractive indices—for example, nuclei, starch grains, and plant cell walls have high refractive indices. The effect of this is that the focal plane is not really flat, but has bumps in it corresponding to the different refractive indices in the structures. The spherical aberration caused by refractive index mismatch is likely to be dependent on the depth of the focal plane within the specimen. Therefore, the imaging is no longer shiftinvariant, since the p.s.f. differs for different z planes within a 3D image. Chromatic aberration can also have important effects, particularly in confocal imaging. In the presence of chromatic aberration, light of different wavelengths is brought to a focus at different focal planes, and in the case of light originating away from the central optical axis, at slightly different x-y positions. We have shown above that confocal image formation can be considered as the overlapping of the images of the excitation and detection pinholes. For fluorescence imaging, the excitation light is a shorter wavelength than the detected light, and if chromatic aberration is present, the two pinhole images are centred at slightly different places and so do not overlap fully. plane to show the 3D shape of the p.s.f. more clearly. The out-of-focus rings (or cones in 3D) can be seen to be much stronger one side of focus than the other, due to spherical aberration. (D) A similar x-z section of the confocal p.s.f. shown in (B). These images were all measured using a Leitz planapo x 63 oil immersion objective (NA 1.4). The microscope was a Zeiss photomicroscope, either linked to a Bio-Rad MRC600 confocal microscope, or to a Photometries series 200 cooled slow-scan CCD camera. The beads were labelled with fluorescein, and imaged with standard fluorescein filter sets in each case. Laser excitation wavelength 488 nm.

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P. J. Shaw This reduces the amount of light detected and can change the effective shape of the confocal p.s.f. Moreover these effects are position-dependent, the overlap becoming worse the deeper into the specimen or the further the imaging is from the optical centre. Thus, again, the p.s.f. is no longer shift-invariant. The significance of different types of aberration on confocal imaging is still a matter for research and debate. It is clear that the detected confocal image, even with visible wavelengths, becomes increasingly faint deep into thick specimens (say 100-200 mm). This effect is most marked for shorter wavelengths, and least for longer wavelengths. It is probably due to a combination of increasing aberrations and increasing light scattering. Chromatic aberration is of very great importance for UV confocal imaging, since virtually all achromatic or apochromatic objectives are corrected only for some visible wavelengths, and have considerable achromaticity in the UV (in fact very few objectives even transmit very well below 400 nm). This has meant that confocal microscope manufacturers have had to make special allowances for UV imaging. In the machine made by Leica (TCS UV), the illuminating pin-hole for the UV channel is placed in a different plane from that for the visible excitation channels. In the case of the Bio-Rad UV design (MRC 1024 UV), which has a single illuminating aperture for all wavelengths, a pre-focusing lens system is used in the UV light path, so as to bring the focal plane for the UV light to the same level as the other wavelengths. A similar compensation is used in the machine made by Zeiss (LSM 510-UV). (See Handbook of biological confocal microscopy (2), Chapter 27 for a detailed discussion of UV confocal microscopes.)

3. Practical implementation of confocal scanning systems We have shown how the confocal pin-holes combine to eliminate the out-offocus light from detection for a point on the specimen and thus improve the characteristics of the p.s.f. To complete the description of a practical confocal microscope, we now consider in outline how the image of a specimen plane is built up by a scanning mechanism. The third dimension (z) for a threedimensional image is produced by scanning 2D images at successive focal planes, either by moving the specimen stage or the objective through a series of small focus steps.

3.1 Point scanning In principle, the illuminated spot can be itself scanned across the specimen in a raster, in a manner very similar to the scanning electron beam in a TV screen, or the specimen can be moved through a stationary light path. The latter design has the advantage of a simple and accurate optical design, but suffers from lack of speed in scanning an image, particularly at low magnification. 54

2: Introduction to confocal microscopy Some of the first confocal microscopes built were of this stage scanning type. However, virtually all current biological confocal microscopes use a scanned beam design. An angular deflection of the light beam at a diffraction plane becomes transformed into a translation in the specimen/image planes. Thus, a system of two deflecting mirrors scanning back and forth about two axes positioned at or near a diffraction plane can be used to scan the light beam in a 2D raster across the specimen. The scan driving and measurement circuitry are interfaced together so that light intensity measurements are taken which cover the specimen area in a regular raster, and these intensities are digitized into a computer image framestore to produce a digital image. Generally image accumulation and averaging are provided, either frame by frame, or line by line, or both. Often, the scanning rate can also be changed. This principle is used in confocal microscopes made by several companies, including Bio-Rad, Leica, and Zeiss. With currently available mirror deflection systems, the maximum scanning rate is a few frames/second (i.e. well below 'real time' or video scanning rates). A detailed description of the various scanning systems and a discussion of their relative merits is given by Stelzer (Handbook of biological confocal microscopy (2), Chapter 9). We will not attempt to give a detailed discussion of the merits of these machines, since the models are still evolving and so such data would quickly be out of date. Most of the spot scanning confocal microscopes currently in use are from these three manufacturers, and anyone considering purchasing a confocal microscope is advised to test at least these machines. It is important to use a specimen which is stable and familiar for assessment, so that the user knows what sort of image to expect, and preferably can image the same specimen on all the contending machines. It is also a good idea to take images in stored digital form away from all the tested machines to compare with each other on a known computer and printer back at the home laboratory.

3.2 Slit scanning In order to increase the scanning to video frame rates or higher, slit scanning designs have been developed. Instead of a pin-hole aperture giving a diffraction limited spot, a narrow slit of light is scanned in a direction at right angles to its length across the specimen. This is achieved in a similar way to spot scanning by using one or more scanning mirrors and a stationary slit aperture. The emitted light is then passed through a narrow detector slit. This has the advantage that since only a one-dimensional scan is required, the scanning rate can be much faster than in a point scanning system. Furthermore, since a line of the specimen is imaged at one time, the rate of light accumulation from the specimen is much higher. However, the disadvantage is that a proportion of the out-of-focus light—that component which is distributed in the direction of the slit—is also detected, and so the optics are only partially confocal. This design, therefore, is inherently not capable of producing such clean optical sections as a point scanning system, but the fast scanning speed and bright 55

P. J. Shaw image produced mean that the image can be observed directly through an eyepiece, as in a conventional microscope. For many applications, particularly those where a single focal section relatively free from out-of-focus blur is required rather than a complete 3D image, this design provides a good compromise between conventional imaging and confocal imaging. Linked up with a high sensitivity CCD (charge coupled device) camera, it could be a good choice for dynamic studies of weak or light-sensitive specimens. Slit scanning microscopes for biology are currently made by Bio-Rad (ViewScan) and Meridian Instruments (Insight). Slit scanning and video rate systems are discussed in detail in the Handbook of biological confocal microscopy (2), Chapters 25 and 29. A design which is intermediate between a point and slit scanner is made by Noran. This machine, Odyssey, scans the incident light beam with a device called an acousto-optical modulator (AOM). AOMs work by using sound waves to set up a standing wave in a crystal, which then behaves in a very similar way to a diffraction grating, producing a diffracted beam at an angle to the incident beam. By modulating the sound frequency and thus the effective grating spacing, the beam can be deflected or scanned through different angles. The major problem is that since the deflection is wavelength-dependent, unlike a mirror system, the fluorescent light, at a longer wavelength than the excitation light, cannot be sent back along the same path through the AOM. In practice the detected light is passed through a slit aperture. Thus the optical arrangement is point scanning for excitation, slit scanning for detection. The design is capable of very high scanning rates.

3.3 Spinning disc The final confocal design we shall describe was actually the first to be built, by Petran (4). Instead of a single pin-hole and a point light source, this design uses an extended light source and an array of many pin-holes on a Nipkow disc which is placed in a conjugate image plane. In the original design the array of pin-holes was symmetric about the centre of the disc. The incident light passed through each pin-hole, and the reflected or fluorescent light passed through equivalent pin-holes on the opposite side of the disc. In a more recent design, the emitted light passes back through the same pin-holes. The condition necessary for confocal imaging is that no emitted light should pass through the 'wrong' pin-hole, and this means that the pin-holes must be spaced far apart relative to their diameters. This means that only a limited set of points are imaged by the disc in each position. The full image is obtained by spinning the disc rapidly, so that the pin-holes, which are usually arranged in a spiral pattern, scan across the whole image area. The main problem with this design is that the light source has to be spread out over the whole of the disc, and so is orders of magnitude less bright that the single pin-hole/laser arrangement. Furthermore, only a very small proportion of the available light passes 56

2: Introduction to confocal microscopy through the disc to illuminate the specimen, and more seriously, only a very small proportion of the reflected or emitted light passes back through the disc to be detected. The result is that it is difficult to record enough light for a satisfactory image, particularly in the case of fluorescence, and much of the available fluorescent light is wasted. This has substantially limited the use of this design in biological applications. Juskaitis et al. (5) have recently designed a variation of the spinning disc approach which overcomes the problem of poor efficiency to a large extent. A detailed description is beyond the scope of this chapter, but in essence they have shown that it is possible to design discs with 25% or more light transfer efficiency that produce an image which is the sum of the conventional and the confocal image. In the prototype machine, half the disc was of this design, the other half was transparent. The image produced therefore alternated between the conventional and the composite image. Digital image processing was used to subtract the conventional from the composite image, leaving the confocal image. It is hoped that this simple and elegant idea can be quickly commercialized; it promises to be a very cheap, powerful, and efficient system.

3.4 Two-photon imaging Two-photon imaging has attracted much attention recently. So far, several research machines have been constructed, and a commercial system is available from Bio-Rad. In conventional fluorescence microscopy, the specimen is illuminated with photons of the correct wavelength to raise an electron in the fluorophore to a higher energy level. However, if a high enough intensity of light at double the required wavelength can be used, then the fluorophore can absorb two photons almost simultaneously to produce the required energy level change. The combined probability of the double absorption depends on the square of the light intensity distribution. Thus, the illuminating p.s.f. is the square of the conventional p.s.f. (i.e. very similar to a confocal p.s.f.). In practical instruments, a detector pin-hole has also been used to give an improved p.s.f. The advantages of this method are that light should only be absorbed near the focal spot, and thus should only cause fading and photodamage at this position in the specimen. In normal confocal imaging, although light is only detected from near the focal plane, light is absorbed by the specimen at every plane, causing the associated photobleaching and other photodamage, in just the same way as in conventional, wide-field microscopy. Another potential advantage of two-photon imaging is that the long illuminating wavelengths used—about 1000 nm—are relatively good at penetrating into deep specimens. Also, UV fluorophores can be excited with visible wavelengths, which avoids the problems mentioned above with UV optics. The disadvantage is that a powerful and expensive pulsed laser system must be used. Currently, such laser systems cost in the region of £100000, but it is hoped that more affordable lasers will soon be available. Another potential problem 57

P. J. Shaw is damage to the specimen by the high intensity long wavelength incident light. Experience with different cells is so far limited; some living cells have been imaged without problem, but the presence of any cellular constituents which absorb in the far red/near IR region of the spectrum would have disastrous effects, and cause the specimen to be rapidly destroyed by heating.

4. Comparison of conventional, wide-field fluorescence imaging with confocal fluorescence imaging We refer readers elsewhere to a more detailed comparison of wide-field and confocal imaging (6), merely giving here a brief discussion of some of the more important aspects of this topic. A major limitation of most current confocal microscopes, apart from the spinning disc designs, is that they must use laser light sources, with consequent severe restrictions on the available wavelengths. While developments in laser technology will reduce these limitations, it is unlikely that they will disappear in the foreseeable future. In contrast, conventional, wide-field fluorescence microscopy can use virtually any wavelength from the visible spectrum and beyond.

4.1 Noise and resolution The major advantage of confocal microscopy over conventional, wide-field microscopy is the elimination of the out-of-focus light. This is broadly equivalent to an increase in resolution in the z direction—the optical axis. In principle, a confocal microscope is capable of better resolution in the image (x-y) plane as well, but this will only be realized if very small pin-holes are used, and if the signal-to-noise ratio is large enough. Figure 4 shows a diagram of the contrast transferred (in the x-y plane) as a function of resolution for a wide-field and for a confocal microscope (data redrawn approximately from ref. 7). It is clear that the confocal microscope has a better contrast transfer value, especially towards the resolution limit, than the wide-field microscope. However, the image detail actually observable will also depend on the noise level in the image; the higher the relative noise, the more the image components will be lost beneath it. Since the image contrast decreases at high resolution this means that increasing the noise level will have the effect of decreasing the resolution in the image; coarse, large scale structure remains above the noise, while fine detail is lost. In practice, the noise in a confocal image can easily be very much higher than the noise in a wide-field image as we will show below, and so it is entirely possible for the effective resolution to be worse for a confocal image. For the sake of this discussion we will assume that a wide-field image is recorded with a high quality, scientific grade cooled CCD camera, and compare it with the image from a spot scanning confocal microscope which uses a 58

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Figure 4. Diagram to show contrast transfer as a function of in-plane resolution. The confocal microscope has a better contrast transfer, especially towards the resolution limit, but this can easily be outweighed by poorer signal-to-noise characteristics.

photomultiplier detector (see refs 6, 8, and 9 for more detailed discussions). Leaving aside for the moment where in the specimen the photons making the image have come from, we will compare the two systems purely in terms of their image detection efficiency and noise. Noise in the image can be considered as coming from two sources: (a) Detector noise—noise introduced by the detection device and circuitry. (b) Poisson noise—noise arising from the statistical distribution of the recorded photons. In general terms, detector noise is most important at very low photon levels, Poisson noise at higher levels. In a typical cooled CCD camera, the measurement noise, mostly readout noise from the A/D converter in the camera, is about ten electrons (r.m.s.) per pixel. This means that if many measurements of exactly the same number of electrons were made, the standard deviation (root mean square deviation from the mean value) would be ten. So, if exactly 100 electrons had accumulated in a particular pixel in the array, there would be a probability of about 70% that the measured number would be between 90 and 110. The detective quantum efficiency (i.e. the fraction of incident photons which produce a detectable electron in the charge wells—DQE) varies between 20% and 80% depending on the wavelength and the CCD chip design. Let us assume 50%. This means that the detector noise is equivalent to about 20 incident photons. Poisson noise arises because the emission and detection of photons is a random process and fluctuates according to the Poisson distribution. So if many measurements of a 'uniform' flux of photons 59

P. J. Shawwere made using a perfect measurement device, then the standard deviation of these measurements would be given by the square root of the mean value. So, if the detected number of photons is 100, we can regard this as a sample of a probability distribution whose mean is 100 and whose standard deviation is 10. Thus at a signal level of 400 photons, the Poisson noise would be 20 photons—the same level as the detector noise. At this signal level we could expect an r.m.s. noise-to-signal ratio of around 7% (400 photons measured with 20 photons Poisson noise and the equivalent of 20 photons detector noise—since the two sources of noise are independent we can add their contributions as the square root of the sum of their squares). With a higher signal level, the Poisson noise would be greater than the (signal-independent) detector noise. The photomultipliers used in confocal microscopes, if they are kept cool, or at least not allowed to get warm, and if the detection circuitry is optimal, are capable of very low detector noise levels—probably one to two counts or lower. Thus, to obtain the same 7% signal-to-noise ratio as for the CCD camera we would only need to measure about 200 counts. (Only Poisson noise will contribute significantly and V200/200 is about 0.07.) However, the DQE is much lower—perhaps only 10% or less for a typical photomultiplier—so detecting 200 counts would require 2000 incident photons at this sort of efficiency level. Thus, purely in terms of image detection, the confocal microscope is clearly worse than the best scientific CCD cameras. But another problem for the confocal microscope comes when we consider how the photons arrive at the detectors. In the wide-field microscope, the whole specimen is illuminated at once and a 2D image is recorded at the camera. For typical biological fluorescent specimens, it is common for a one second exposure to give thousands of detected photons in the brightest pixels. On the other hand, the confocal microscope accumulates the image one pixel at a time, and each pixel is only exposed to the laser beam for a very short time (perhaps a microsecond or less). It is not easy to estimate the number of photons being measured, but it has been estimated that in a typical specimen, the brightest pixels may only represent 10-20 photons in a single scan. Therefore to obtain the numbers of photons in the image that a CCD camera records in a single second exposure may require hundreds of successive confocal scans to be accumulated. This is usually not feasible, and thus confocal images generally have a considerably higher noise level than wide-field images recorded by CCD cameras.

4.2 Out-of-focus light As has already been mentioned, the principle advantage of the confocal microscope is that it eliminates the unwanted out-of-focus light from measurement. This is entirely equivalent to the differences in the p.s.f.s between confocal and wide-field microscopy that have been described above. However, 60

2: Introduction to confocal microscopy the fact that the measured image can be described mathematically as the convolution of the object with the p.s.f. means that it is in principle possible to reverse this modification by the p.s.f. This process can be accomplished by various computer image processing methods. A detailed description is beyond the scope of this chapter, and the interested reader is referred elsewhere (6,10). This removal of the out-of-focus component in a wide-field image is usually called deconvolution, restoration, or simply deblurring. Deconvolution methods can also significantly improve confocal images (6). The faithfulness with which such a reconstruction can be achieved depends on the noise level in the image. With no noise present, a perfect reconstruction can be carried out trivially. But, as we have already described, all images are certain to contain noise, often at relatively high levels. In the presence of noise, image restoration becomes a technically difficult problem. Computer methods for handling this problem have been extensively developed, and suitable programs for optical microscopy are now commercially available (see ref. 10 for a discussion of this topic and a survey of currently available software). Thus, it is possible to collect wide-field images taking advantage of the excellent imaging characteristics of scientific grade CCD cameras, and using image processing to counteract the effects of the wide-field p.s.f. However, it should be realized that while it may be possible to eliminate or reduce the out-of-focus component from each focal plane by restoration methods, this unwanted light is present in the measured image, and contributes to the noise in the measured image. The more out-of-focus light there is in the wide-field image, the more noise it adds to the image, and the smaller is the fraction of the measured photons corresponding to the in-focus image. The confocal arrangement eliminates this light from measurement and, along with it, the associated noise. So the more out-of-focus light is present, the greater the benefit of eliminating it from measurement by the confocal arrangement. However, although the confocal optics eliminates the out-of-focus light from detection, it does not prevent parts of the specimen out of the plane of focus from being illuminated by the excitation light. Photodamage is therefore likely to be as great with confocal as with conventional imaging, other factors being equal. Two-photon and other related methods hold out the promise of improving on this situation by restricting the specimen excitation to the centre of the focal spot.

4.3 When should confocal microscopy be used? Although the main aim of this chapter is to explain the basis of confocal imaging, we shall briefly discuss some of the factors that should be considered in deciding whether confocal microscopy is the most appropriate technique for a particular imaging experiment. Since confocal microscopes do not currently compare favourably with high grade CCD cameras in detection sensitivity and speed of image capture, they are not well suited to imaging very weak or photosensitive specimens. It is 61

P. J. Shaw entirely possible that a confocal microscope will produce an image for a weak specimen that is actually worse than a conventional fluorescence microscope with an attached CCD camera. The only real advantage of a confocal microscope is that the out-of-focus light is eliminated. Therefore there is only any real point in using confocal microscopy where this advantage is important; in practice this means for specimens with substantial thickness. But it is not always obvious what 'substantial' means. Certainly we have obtained good confocal images from specimens several hundred micrometres in thickness in which the in-focus signal was virtually obscured by the out-of-focus light, making the conventional fluorescence image so poor as to be useless. On the other hand it is pointless using a confocal microscope to image thin microtome sections (a fraction of a micrometre in thickness). Better results will almost certainly be obtained by the better image detection capabilities of a good camera on a conventional microscope. However, most real specimens are somewhere in between these two extremes, and it is not always easy to predict what imaging method will perform best for a given specimen. To some extent what is used will depend on what facilities are available. If a very good cooled CCD camera, coupled with state-of-the-art deconvolution software is available, and, importantly, if there is access to expertise in using it, then it is likely that better images can be produced in this way in many cases, at least for specimens which are no more than one cell thick. There are good examples in the literature of outstanding 3D reconstructions produced in this way, and several integrated imaging and restoration packages are now commercially available. Nevertheless the expertise necessary to achieve such good results should not be underestimated. Most researchers involved in projects of which imaging is typically only one part, and who therefore cannot afford the time to learn to be experts, will probably find it quicker to learn how to use a current confocal microscope adequately than a CCD/image processing workstation system. However both types of system are rapidly improving in power, affordability, and user-friendliness. The ideal situation is to have access to both types of equipment and to use whichever provides the best imaging for the experiment in hand. There is a great deal of overlap between a confocal microscope and associated computer and a high quality CCD microscope imaging system, in both microscope and computer hardware, and in the software needed for image collection and subsequent analysis and processing. It may be hoped that one of the various manufacturers might take the step of integrating both types of imaging into a single instrument. Figure 5 shows an example of the same specimen—a Drosophila embryo labelled with rhodamine-phalloidin to show the actin distribution—imaged by both conventional fluorescence using a cooled CCD camera and by confocal fluorescence microscopy. Because this is a relatively thick specimen with a great deal of out-of-focus light arising from other planes than the plane of focus, the difference between the two methods of imaging is dramatic. 62

2: Introduction to confocal microscopy

Figure S. Comparison of conventional, wide-field and confocal imaging. (A) An unprocessed conventional fluorescence image, collected using a cooled CCD camera, of a Drosophila embryo (gastrula stage) in which the actin network has been labelled with rhodamine-phalloidin. The large amount of out-of-focus light masks much of the fine image detail. (8) A confocal image of the same specimen. The exclusion of the out-offocus light leaves the image detail easily visible. (Specimen courtesy of Richard Warn.) The confocal microscope, CCD camera, and microscope objective were all the same as in Figure 3, In this case the fluorochrome was rhodamine, and the standard filler sets for this fluorochrome were used. Laser excitation wavelength used was 568 nm.

5. Practical examples of specimen preparation for confocal imaging We conclude with two examples of specimen preparation for confocal imaging taken from our current research into plant nuclear organisation. The purpose is not to give an exhaustive set of protocols, which are necessarily spccialized and depend on the biological system and type of study, but rather to illustrate some general points and provide a link between the foregoing theory and practical biological imaging studies. Other chapters in this volume (e.g. Chapter 6) provide more specialised protocols for specimen preparation, data collection, and subsequent data handling. In general terms, confocal fluorescence microscopy simply requires a fluorescent specimen, which can be prepared in a similar way to any other lluorescently labelled specimen. However, the ability of the confocal microscope to produce clean optical sections from thick, thrcc-dimensionally well preserved specimens should not be wasted. In practice, this means that some care should be taken to preserve the three-dimensional structure of the specimen as well as possible. For living specimens, methods that keep the tissue or cells alive and active will almost certainly preserve three-dimensional structure well. For dead, fixed specimens the best fixative that is consistent with the 63

P. J. Shaw labelling method should be used. We nearly always use formaldehyde solutions (which should be freshly made by dissolving solid paraformaldehyde), sometimes with a small percentage of glutaraldehyde. Electron microscopy has shown that the bifunctional glutaraldehyde is a better fixative than formaldehyde, but it often interferes with penetration of antibodies and other probes. It also causes a high autofluorescent background, which can be alleviated to some extent by a subsequent treatment with sodium borohydride. Fixed tissues often also need extra permeabilization to allow penetration of probes. In the cases of plant material, partial digestion of the cell wall with cellulase and other cell wall degrading enzymes is usually needed. Producing good specimens usually depends on achieving a balance between preservation of the structures of interest and disrupting them so as to allow probes in to visualize the structures. In general, it is best to leave the physical form of tissues as unaltered as possible. So in one of the protocols below we describe how to label entire roots from the plant Arabidopsis. In many cases the tissue of interest is within a relatively large organism or organ and must be physically removed or sectioned to make labelling and imaging possible. This is the case with the large roots of pea seedlings described in the second protocol. For these cases we routinely use a vibratome, which can cut quite thick sections (50-100 |xm) from living or fixed roots. Finally, the specimen should be mounted with care, so that the coverglass does not squash the carefully preserved structure. For thin specimens we find that the thickness of the coating between the wells on multiwell slides is sufficient to protect the specimen from the coverglass. For thicker specimens it may be necessary to support the coverglass. Either use nail varnish or more coverglasses to make a platform to raise the coverglass away from the specimen. In principle any objective which can be used for conventional fluorescence can also be used for confocal fluorescence imaging. As with all fluorescence imaging the brightness of the image is strongly dependent on the numerical aperture (NA) of the objective and so objectives with the highest available NAs should generally be used. As discussed above, confocal imaging is more seriously degraded by aberrations than conventional imaging, and so it is preferable to use high quality planapochromat objectives if possible. However it should be noted that while all planapochromat objectives are suitable for fluorescence imaging with visible light (e.g. FITC, rhodamine, Texas Red, Cy3), the objectives of this type made by some manufacturers do not transmit the UV light required for excitation of the common DNA dyes like DAPI. In fact very few available objectives transmit the commonly used UV laser wavelengths from high power argon ion lasers efficiently (351 and 363 nm). We nearly always use oil immersion objectives for confocal imaging, in spite of the spherical aberration and refractive index mismatch problems described above. We have obtained optically very good results from a water immersion, 64

2: Introduction to confocal microscopy coverglass-free objective (Zeiss, X 63,1.2 NA). In fact the lower aberration in use of this objective means that optically better images are obtained with it than with the oil immersion objectives having higher numerical apertures (e.g. Nikon, X 60,1.4 NA, and Leitz, X 63, 1.4 NA). However, using these coverglass-free objectives poses several problems: the specimens are not well protected, and are difficult to keep; focusing the objective transfers forces to the specimen which tend to cause it to move around and often to break up; it is difficult to find a satisfactory water-based antifade mountant. A water immersion lens with a coverglass would be a better alternative. Several manufacturers are now making these objectives, but at the moment they are very expensive. Protocol 1. Whole mount immunofluorescence of Arabidopsis roots Reagents • Bovine serum albumin (BSA) (Sigma, ACellulase R10 (Yakult Honsha Co. Ltd.) 7030) Pectolyase (Sigma) Nonidet P-40 (NP-40) (Fluka Chemicals Ltd.) Driselase (Sigma) PEM buffer: 50 mM Pipes (Sigma), 5 mM -y-aminopropyl tri-ethoxy-silane (APTES) EGTA (Sigma), 5 mM MgSO4, pH 6.9 with (Sigma) KOH . PBS buffer: 130 mM NaCI (Sigma), 7 mM . Blocking solution: 3% BSA in PEM, 0.2% Na2HPO4 (Sigma), 3 mM NaH2P04 (Sigma) NP-40 pH 7.4 . 4,6-diamidino-2-phenylindole (DAPI) (Sigma): Paraformaldehyde (Sigma) 1 M-Q/ml in distilled water Glutaraldehyde: EM grade, 25% solution Vectashield mounting medium (Vector Lab(Agar Scientific Ltd.) oratories Inc.) Decon 90 (Decon Laboratories Ltd.)

Formaldehyde fixative Add 8% (w/v) paraformaldehyde to distilled water (e.g. 2 g to 25 ml) and heat with constant stirring to 60°C. Add one drop of 1 M NaOH (for 25 ml) and continue to stir at 60°C. The cloudy suspension should clear to a colourless solution within a minute or two. Cool to room temperature, add an equal volume of 2 x concentrated PEM buffer and check the final pH. (It is recommended that indicator paper or a dedicated pH meter is used for this, since fixatives can damage pH electrodes, mainly by cross-linking protein contaminants onto the permeable plug.) If the paraformaldehyde does not dissolve easily with this procedure (e.g. requires more NaOH to be added) it is recommended that it should be discarded and a new batch obtained. In our experience more failures in labelling experiments can be traced to problems with fixation than to any other cause. We regularly replace stocks of paraformaldehyde, at intervals of six months to one year, and store it in small, separate aliquots at 4°C. Once opened, an aliquot is used or discarded after a few days. 65

P. J. Shaw Protocol 1. Continued APTES treatment of slides To ensure that sections adhere firmly to slides, we routinely pre-treat the slides with APTES, which we find more effective than the commonly used treatment with poly-L-lysine. Wash slides with ethanol or with 2% (v/v) Decon 90 or similar detergent, for a minimum of 1 h, then rinse thoroughly with several changes of distilled water. Place slides in a freshly prepared solution of 2% (v/v) APTES in acetone for 10 sec, then transfer to acetone alone to wash, and allow to air dry. The slides can then be stored in dustfree conditions for up to six months, but gradually lose their effectiveness. Just before they are needed, the slides are placed in a 2.5% (v/v) solution of glutaraldehyde in PBS for at least 30 min, then rinsed in distilled water, and air dried. Method 1. Fix roots in 4% freshly made formaldehyde in PEM, 0.2% NP-40 for 1 h. 2. Wash in PEM, 0.2% NP-40 and then in PEM alone. 3. Dry onto APTES treated multiwell slides. 4. Treat with 0.05% cellulase, 0.025% pectolyase, 1% driselase in PEM for 10 min. 5. Wash in PEM, 0.2% NP-40 and allow to air dry. 6. Block with blocking solution for 90 min. 7. Incubate in primary antibody overnight at 4°C. 8. Wash in blocking solution for 1 h. 9. Incubate in secondary antibody for 2 h at 37°C. 10. Wash in PEM, 0.2% NP-40 for two days. 11. Counterstain with DAPI if required and mount in Vectashield or other antifade mountant. If a coverglass is used, it should be the correct thickness for the objective lens that is intended to be used—this is usually 0.17 mm and corresponds to number 1 1/2. The imaging will generally be worse with any other coverglass thickness, whether greater or less. It is important that the coverglass does not squash and flatten the specimen. If necessary, the coverglass should be supported by small fragments of broken coverglass placed on the slide. The coverglass should be secured in position by nail varnish at the edge.

Figure 6 shows part of an Arabidopsis root labelled with an antibody to the spliceosomal protein U2B", which is located in the nuclei both in the interchromatin region and prominently in the coiled bodies. 66

2: Introduction to confocal microscopy

Figure 6. An Arabidopsis root labelled with antibody to the U2-associated spliceosomal protein U2B". The most prominent structures fabelled by this antibody are small, spherical bodies within the nuclei, ranging from one or two to several per nucleus. We have shown that these bodies correspond to structures called coiled bodies by electron microscopists (because the EM ultrastructure often displays an apparent substructure of coiled fibres), A confocal section showing files of epidermal cells is shown. The files which will develop root hairs (e.g. the central file) contain markedly more coiled bodies than the files which will not make root hairs (e.g. the two files either side of the central file). A fluorescein labelled secondary antibody was used, and imaging was carried out on a BioRad MRC600 confocal microscope as in Figures 3 and 5, using 488 nm excitation laser light, and a Leitz planapo X 63 oil immersion objective (NA 1.4). Bar = 10 [sm. (Courtesy of Kurt Boudonck.)

Protocol 2. In vitro transcription on vibrato me sections of plant roots Equipment and reagents Vibratome (e.g. Series 1000 Vibratome from TAAB Laboratories Equipment Ltd.) Multiwell slides: glass 8-well multitest slides from ICN Biomedicals, Inc. PB buffer: 100 mM potassium acetate (Sigma), 20 mM KCI (Sigma), 20 mM Hopes (Sigma), pH 7.4 with KOH 1 mM MgCl2 (Sigma) 1 mM ATP (disodium salt, from Sigma) 1% (v/v)thiodiglycol (Sigma) 2 ng/ml aprotinin (Sigma) Triton X-100 (Sigma)

0.5 mM PMSF (phenylmethylsulfonyl fluoride, Sigma): make up as 100 mM stock solution in ethanol, store at -20oC, and add to buffer just before use Transcription mix: 50-500 jJV! CTP (sodium salt, Pharmacia), 50-500 (iM GTP (sodium salt, Pharmacia), 25-250 iiM BrUTP (sodium salt, Sigma), 125 ^M MgCl2, 100 U/ml RNA Guard (Pharmacia), made up in PB TBS: 25 mM Tris-HCI pH 7,4 (Sigma), 140 mM MaCI, 3 mM KCI Hexylene glycol (2-methyl 2,4-pentanediol, Sigma)

67

P. J. Shaw Protocol 2.

Continued

Method 1. Excise the first 3-4 rr»M of a freshly germinated root tip and mount on the vibratome. Cut sections (40-50 \i.m) in PB containing 1 M hexylene glycol and transfer to tissue handling device (11). 2. Incubate in 0.05% Triton X-100 in PB for 1 min, then wash in PB alone, three times over ~ 30 sec. 3. Incubate in transcription mix for 5 min or other times, then wash again in PB alone, three times over ~ 30 sec. The lower concentration of NTPs given is the minimum that will support transcription in our experience, the higher concentration routinely gives high levels of transcription. 4. Fix in 4% freshly made formaldehyde in PEM (see Protocol 1) for 1 h. 5. Wash in TBS for 10 min, then H20 for 10 min. 6. Remove from tissue handling device and allow to air dry onto APTES coated multiwell slides (see Protocol 1). 7. Treat sections with 2% cellulase (Onozuka R10) in TBS for 1 h, then wash in TBS, three changes over 15 min. 8. Incubate for 1 h with primary antibody (mouse anti-BrdU; Boehringer), diluted 1:20 in PBS, 3% BSA. 9. Wash thoroughly with TBS, then incubate for 1 h with secondary fluorescent antibody (Cy3-conjugated anti-mouse; Jackson Immunoresearch). 10. Mount in Vectashield or other antifade mountant.

Figure 7 shows a section of pea root labelled in this way to show the sites of RNA transcription. The nucleoli in the centre of the nuclei are most strongly labelled and correspond to incorporation by RNA polymerase I, with many dispersed transcription sites corresponding to RNA polymerase II and III throughout the rest of the nucleus. Protocol 3. Collection of confocal images This is not really a protocol so much as a set of procedures that should be used in optimizing the parameters for collection of confocal images. In principle it should apply to any confocal microscope, but the details will differ on different machines. The settings described in steps 1-4 all interact with each other, and must often be iterated to produce the best image. What constitutes the 'best' image also depends on the imaging experiment being undertaken. Ideally, for multiprobe imaging of several different 68

2: Introduction to confocal microscopy fluorochromes, steps 1-4 should be optimized for each fluorescent probe in turn, although this is not always possible. Method 1. Optimize the laser illumination intensity. This depends on the specimen and the fluorescent labelling. In general the highest intensity that will not cause problems should be used. Problems from too much illumination light include fading of the fluorochrome and photodamage to living cells. It should be remembered that the very high laser light levels used can easily cause a complete population inversion of the fluorescent molecules to the excited state; any further excitation will only cause photodamage. Often it is better to use a lower illuminating light level, and average the image over a longer time. 2. Optimize the detector pin-hole diameter. The smaller the pin-hole, the better the resolution—primarily the better the exclusion of out-offocus light, but the less light is detected and the poorer the statistical properties of the resulting image. It is often better to open up the pinhole for a weak specimen, and compromise on the confocality of the image. 3. Optimize the setting of the photomultiplier amplifier circuitry, usually provided as a gain and black-level setting. In general this should be set so that maximum and minimum detected signals in the image correspond to the highest and lowest digitized level respectively—i.e. usually 255 and 0 for an 8-bit analogue to digital conversion. It is often useful to arrange for these values in the image to be flagged as different display colours, and the computer software will often provide a special look-up table (LUT) for this purpose. If the minimum and maximum are set too low and high, then the image grey levels will not span the available range and will be compressed into a smaller grey level range than necessary. This could result in the loss of image information. If the minimum is set too high or the maximum is set too low, this will cause artefactual thresholding of the image at the set minimum or maximum, and again may result in the loss of image information or the introduction of artefacts. 4. Optimize the image averaging. Single image scans usually show poor image statistics—i.e. high levels of image noise—simply because of Poisson noise given the low numbers of photons detected in a single scan. The solution is to accumulate many scans of the image, or to scan at a slower rate, or both. The image signal-to-noise ratio increases as the square root of the number of scans accumulated. The disadvantage to accumulating many scans is the time taken and the increased light dosage, causing fluorochrome fading, photodamage, etc. In the case of living cells, dynamic changes may occur and be blurred by 69

P. /. Shaw Protocol 3.

Continued

long accumulation series. The requirement for dynamic information and good image signal-to-noise must be balanced against each other. 5. Set up the other data collection parameters, such as for focal series (z series), time series, etc., and collect the image data.

Figure 7. A pea root section labelled by BrUTP incorporation. The strongest labelling is from incorporation by RNA polymerase I into pre-rRNA in the nucleolus, but all cells also show labelling in the nucleoplasm in many disperses foci, which must be due to incorporation by RNA polymerases II and III. The confocal microscope used was an MRC600 (as in the previous figures). An argon/krypton mixed gas laser was used In the confocal microscope, and an excitation wavelength of 568 nm was used to excite the fluorochrome (Cy3). Bar = 10 um, (Courtesy of Alison Beven.)

References 1. Stelzer, E. H. K.. Lindek, S., Albrecht, S., Pick. R., Ritter. G., Salmon, N. J., et al. (1995),J, Microsc.,179, I. 2. Pawley, J. B. (ed.) (1995). Handbook of biological confocal microscopy. Plenum Press, New York and London. 3. Castleman, K. R. (1979). Digital image processing. Prentice-Hall, 4. Petran, M., Hadravsky, M., Egger. M. D., and Galambos, R. (1968), J. Opt. Soc. Am.,58.66l. 5. Juskaitis, R., Wilson, T.,Neil, M. A. A., and Kozubek, M. (1996). Nature, 383, 804. 70

2: Introduction to confocal microscopy 6. Shaw, P. J. (1995). In Handbook of biological confocal microscopy (ed. J. B. Pawley), p. 373. Plenum Press, New York and London. 7. Inoue, S. (1995). In Handbook of biological confocal microscopy (ed. J. B. Pawley), p. 1. Plenum Press, New York and London. 8. Pawley, J. B. (1995). In Handbook of biological confocal microscopy (ed. J. B. Pawley), p. 19. Plenum Press, New York and London. 9. Sheppard, C. J. R., Gan, X., Gu, M., and Roy, M. (1995). In Handbook of biological confocal microscopy (ed. J. B. Pawley), p. 363. Plenum Press, New York and London. 10. Shaw, P. J. (1997). In Cell biology: a laboratory handbook (ed. J. Celis), Vol. 3, p. 206. Academic Press, New York and London. 11. Wells, B. (1985). Micron Microsc. Acta, 16,49.

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3 Video microscopy DIETER G. WEISS, WILLI MAILE, ROBERT A. WICK, and WALTER STEFFEN

1. Video microscopy and the equipment required 1.1 Introduction A new quality of microscopy, called video microscopy, emerges, if one observes the specimen, instead of with the human eye, with a video camera connected to video processing equipment working at real time. Video microscopy is, therefore, much more than just adding a camera and monitor to the microscope to share the images with a larger audience. More recently, electronic devices other than video cameras, such as high sensitivity charge coupled device (CCD) cameras and scanning light detector systems for confocal microscopy have been added to microscopes. The three fields (i) video-enhanced contrast microscopy for highest resolution work, (ii) video-intensified microscopy for low light applications, and (iii) electronic scanning microscopy for confocal microscopy and 3D imaging differ in the type of device generating the electronic image, but all three use basically the same types of analogue and digital image processors. While all these techniques are generally defined as electronic light microscopy, this chapter, video microscopy, deals with the first two techniques that involve CCD and video cameras as imaging devices. Video microscopy has produced a revolution in light microscopy of biological samples equivalent to that of the development of the immunofluorescence technique. It has once more made the traditional light microscope a powerful tool for those working on dynamic aspects of small biological systems, for example biochemists, molecular and cell biologists. It has given further resolving power to the light microscope enabling the observation of particles which bridge the size range between those normally studied by electron microscopy and those which are already well known to light microscopists as a whole, with the added advantage in that specimens can be examined alive. As well as allowing small particles to be resolved, the technique has the capacity to clean up the image, so allowing greater visibility. Also, changes of such parameters as amounts, concentrations, transport, or metabolism of specific molecules in both tune and space can be quantitatively determined. The improvement in resolution is achieved because a microscope equipped

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen

Figure 1. A possible circuit diagram for video microscopy containing equipment for image acquisition, analogue and digital image processing, image display and recording. At least one B/W monitor is required, but one monitor showing the unprocessed live image is recommended for comparison and focusing. ALU = arithmetic logic unit; A/DC = analogue/digital converter; D/AC = digital/analogue converter; LUT = look-up table; ODR = optical disk recorder; RGB = red green blue; Sync = synchronization signal; VTR = video tape recorder.

with an electronic camera, analogue and digital image processors, and electronic display, recording, and printing devices (Figure 1) is able to detect differences in intensity which are far smaller than those detectable in a conventional microscope with the human eye. For the same reason, weakly self-luminous objects can be found and characterized as images. The greatest improvements in microscope imaging are only possible on the basis of very precise optical microscopy arising from a thorough knowledge of the principles of light microscopy, for example of Kohler illumination and conjugate optical planes. However, it should also be said that, on occasion, the less skilled microscopist might be able to considerably improve the quality of the images when using video microscopy. Another major advantage of the technique is that it provides easy recording of information on video tape or disk for later quantitative analysis. Finally, the ability to store and post-process images with PC-based software packages enables microscopists to re-examine their previously obtained data, compare them with more recently produced images, and optimize and finally arrange them for publication. In video microscopy we deal with images that contain the intensity information encoded in either analogue or digital form: 74

3: Video microscopy (a) Analogue: the brightness at each point of the optical microscope image is converted into a voltage signal by the camera. The analogue signal is a continuous signal where 0.4 V represents black and 1 V white. It is interrupted by synchronization signals defining the end of lines and fields. Normally, one frame consists of 576 visible lines (European standard, CCIR) divided into two fields (half frames) containing either all even numbered or all odd numbered lines ('interlaced mode'). (b) Digital: by the use of an analogue-to-digital converter (A/DC) the continuous voltage signal is converted into discrete numbers which are assigned to an array of picture elements (pixels). A common format is 768 X 576, i.e. 768 pixels per line and 576 lines. If an 8-bit conversion is used, one obtains images with 256 grey levels where 0 represents black and 255 white. In analogue image processing contrast can be amplified electronically up to several thousand-fold. In digital image processing contrast is enhanced numerically. The upper limit of useful digital enhancement is only about threefold; however, image quality can be further improved considerably by a large number of additional algorithms. Video microscopy, as dealt with here, involves the generation or improvement of microscopic images in three basic ways (Sections 1.1.1-1.1.3). 1.1.1 Video enhancement Video enhancement is the procedure of increasing contrast electronically in low contrast or 'flat' images. This process not only clarifies images containing details visible to the eye, but renders visible structures 5-20 times smaller than could be detected by vision alone or in photomicrographs (Figure 2). Video-enhanced contrast (VEC) microscopy was developed in the laboratories of S. Inoue (1, 2) and R. D. Allen (3-5). Allen et al. (3, 4) found that the use of VEC microscopy with polarized light methods allowed the introduction of additional bias retardation, which, after offset adjustment and analogue enhancement, permitted much better visualization of minute objects. AVEC (Allen video-enhanced contrast) microscopy is the term used to describe this technique. 1.1.2 Video intensification Video intensification is the procedure for making visible low light level objects and scenes generating too few photons to be seen by the naked eye (Figure 2). Video-intensified microscopy (VIM) amplifies low light images so that very weak fluorescence and luminescence can be visualized (6,7). This is especially important in biology because living specimens benefit from the sparing application of potentially hazardous vital dyes or excessive illumination. VIM and VEC microscopy differ mainly with respect to the type of camera used; i.e. 75

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen

Figure 2. Video microscopy works beyond the former limits of light microscopy. New applications are opened (dotted areas) both in low light situations (VIM) and when working with very small objects (VEC microscopy). All borders are meant to be approximate.

low light level cameras for VIM and high spatial resolution cameras for VEC microscopy. 1.1.3 Digital image processing Microscopic images that have been picked up by a video camera can be converted to a digital signal allowing digital image processing to be performed. Image processing can be used to reduce image noise by digital filtering or averaging, to subtract undesired background patterns, to further enhance contrast digitally, or to perform measurements in the image (e.g. intensity, size, speed, or form of objects). Since the development of procedures for noise reduction and contrast enhancement in real time, that is at video frequency, the microscopist is able to generate electronically optimized pictures while working at the microscope. The procedures to be applied here are generally very similar to those used with other types of electronic imaging with video cameras or photomultiplier tubes, such as confocal microscopy (see Chapter 2) or scanning electron microscopy (SEM). 76

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1.2 General strategies of electronic image improvement 1.2.1 For photon-limited situations In photon-limited situations, especially with fluorescent, luminescent, and most dark-field specimens, video intensification is required. VIM techniques require the use of low light level cameras which, unfortunately, are usually not high resolution cameras. It may be possible, however, to visualize highly fluorescent objects considerably below the limit of resolution if they are well separated from one another (e.g. a diluted suspension of fluorescent actin filaments). The procedure in low light level microscopy would usually require the full range of video-microscopic techniques, that is the use of a videointensification camera, analogue enhancement, and a variety of digital image processing steps such as real time background subtraction (to compensate the uneven sensitivity of some VIM cameras), averaging to reduce noise, and digital enhancement (see Section 3). VIM images with their smooth transitions (low spatial frequencies) are usually well suited for digital image analysis of intensities and intensity changes (see Section 4.2). 1.2.2 For high image fidelity and detail If high image fidelity and detail are desired and the smallest objects of interest are larger than the limit of resolution of the microscope (~ 200 nm), then high resolution cameras are appropriate. They require a fair amount of light but can be used with differential interference contrast (DIC) and all brighter techniques. In such a case we would use VEC microscopy image improvement, but moderate analogue enhancement may suffice. If uneven shading occurs in the image then an analogue shading correction is usually sufficient at this level of magnification. Background correction by digital processing, or digital enhancement is usually unnecessary. Typical applications include observations of whole cells for studies of cell form, cell division, or movement of large organelles. Also, studies at low or intermediate magnifications of small organisms, embryos, or developing eggs, making use of the optical sectioning capacity of DIC microscopy, will benefit considerably. 1.2.3 For visualizing the smallest objects possible If it is desired to visualize the smallest objects possible, one would also use VEC microscopy, preferentially employing DIC or anaxial illumination techniques. Visualization of microtubules (25 nm in diameter) or vesicles with diameters of 50 nm or less can be achieved (8, 9). For this purpose we would need the following functions of electronic image improvement: high analogue enhancement, high performance polarized light microscopy [DIC or POL microscopy according to Allen (3, 4) or Inoue (1)], and digital image processing including real time background subtraction, and digital enhancement. If the resulting image is noisy, real time averaging over two or four frames or real time digital filtering might be employed. 77

Dieter G, Weiss, Willi Maile, Robert A. Wick, and Walter Steffen For most users VEC and VIM microscopy are complementary since the former reveals the intracellular structures while the latter, with the use of fluorescent tags such as dyes or fluorescent antibodies, is needed to determine the identity of the objects depicted. For the microscopist it is most important that the equipment operates rapidly, that is the processor must operate sufficiently fast to display changes in the image in real time.

1.3 The different video-microscopic techniques 1.3.1 Video-intensified microscopy Video-intensified microscopy has greatly extended the capabilities of the light microscope (Figure 2) and has provided the technical vehicle for the development of several important new techniques. VIM has made possible the observation and recording of images too weak to be seen by direct viewing or film recording. Furthermore, it has provided a mechanism to study living cells for extended periods without disrupting normal metabolic activity or bleaching photosensitive molecules. Examples of naturally occurring low light phenomena, such as autofluorescence or bioluminescence, are widespread. Moreover, the use of exogenous luminescent and fluorescent molecules (e.g. fluorescent antibodies or genetically modified proteins tagged with green fluorescent protein) as probes of cellular structure and function has become an important tool in almost all areas of biological research. The application of VIM in these research areas has been reviewed (6,10,11). Video-intensified microscopy is particularly useful in the following situations: (a) Where the total number of photons available for imaging is limited by the nature of the event, as in bioluminescence or in fluorescence where the number of labelled molecules is small. (b) Where low intensity illumination is required to avoid interfering with the biological process(es) under investigation or to avoid phototoxic effects. (c) Where there are rapid changes and the amount of light available in the time interval studied are small. (d) Where the long exposure times necessary for photography could prevent the recording of dynamic processes, e.g. fluorescence from cytoplasmic components in living cells. (e) Where fluorescence excitation needs to be minimized to reduce photobleaching. (f) Where labelling is intentionally limited to avoid biological interference from, for example, toxic dyes. While the procedures of analogue contrast enhancement (see Section 1.3.2) may not be required routinely, digital image processing is still very helpful, especially for noise reduction. 78

3: Video microscopy 1.3.2 Video contrast enhancement Video-enhanced contrast (VEC) microscopy, enables one to increase contrast and magnification to an extent that positions and movements of biological objects as small as 15-20 nm can be analysed in the living state. VEC microscopy is especially useful to the cell biologist, biochemist, and the molecular biologist because: (a) Objects beyond the limit of resolution of conventional light microscopy can be visualized (e.g. microtubules with a diameter of 25 nm). (b) It enables one to visualize cell organelles and supramolecular aggregates in living cells. (c) Under certain circumstances, it allows quantitative measurements of amount, concentration, transport activity, or metabolism of specific molecules. 1.3.3 Analogue contrast enhancement The introduction of analogue enhancement resulted in the remarkable breakthrough which led to a new level of performance of light microscopy. Digital enhancement, as discussed later, is often useful as an addition but it must be emphasized that it cannot replace analogue enhancement. An important basic rule is that only optimized analogue signals should be digitized and processed further. Understanding the image manipulations required for analogue contrast enhancement is complicated and a basic understanding of straylight, contrast, and resolution is required. A short review of each of these aspects is therefore included here. i. Straylight Light distributed evenly over the image and not contributing to image detail is called straylight. Some of its origins are summarized in Table 1. In many cases straylight prevents the use of otherwise optimal settings of the microscope. For example, the resolution achievable is often sacrificed by closing the condenser diaphragm, thus reducing the numerical aperture (NA), in order to avoid too bright an illumination. When polarized light is used there is usually an annoying contribution of unpolarized straylight, even at the highest extinction settings of the polarizers or prisms. In the video image the effect of such straylight can be removed electronically by applying a negative DC voltage to the video signal, called the offset or pedestal voltage. By applying suitable gain to the camera signal the contrast is enhanced, by using a variable offset the camera signal is shifted to the appropriate region of grey levels (brightness) for best visibility on the video screen (Figures 3b and 4). In Figure 3 the improvements achievable by image processing are shown for each step of the technique. 79

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen Table 1. The various sources of straylight which may be removed by applying offset Bright-field microscopy Excessive condenser aperture Uncoated lens surfaces Reflected light from tube inner surfaces Polarized light and interference microscopy Optical rotation at lens surfaces Strain birefringence in lenses Light scatter due to dust, lens cement, etc. Surface imperfections in lenses Defects (holes) in polarizing filters Submaximal compensation Fluorescence microscopy Autofluorescence of any material in the light path Non-specific localization of fluorochromes Incomplete removal of excitation light Bleed-through of one fluorescence in double labelling experiments due to imperfect filters

21. Contrast The brightness at each point of the optical microscope image is converted into a voltage signal by the television camera. Contrast (C) for the eye is perceived approximately as the absolute value of the difference between the intensity (or brightness) of the background (IB) and that of the specimen (IS), divided by the intensity of the background:

Contrast can be amplified, within a factor of 100 or more by the gain applied to the camera signal, provided the proper offset setting is used. 111. Contrast manipulation The manipulation of contrast may be applied to the images in any mode of optical microscopy. With polarized light, considerable additional contrast can be achieved by adjusting the compensator to a higher bias retardation (AVEC microscopy, see Sections 2.1 and 2.3). The resulting images are usually of inadequate visual contrast because the denominator of Equation 1 is too high due to excessive straylight (see Sections 2.1, 2.3.4, and 2.3.5). However, in the analogue image, the offset voltage applied to the video signal acts in a manner analogous to a 'negative brightness or intensity' (Iv), which is added to the 80

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Figure3. Processing of poor contrast image. The specimen, an unstained electron microscope (EM) thin section of striated muscle, was viewed by differential interference contrast microscopy. Video microscopy was intentionally carried out prior to proper cleaning of the optics to demonstrate the procedure in the presence of unusually heavy mottle, (a) In-focus, not enhanced, (b) In-focus, analogue-enhanced, (c) Out-of-focus, with mottle, (d) Out-of-focus, mottle subtracted, (e) In-focus, mottle subtracted, (f) Digitally enhanced. Microscope, Zeiss IM 405, Plan Neofluar, x 63, NA 1,4, x 16 eyepiece, 63 mm camera lens, processor ARGUS, Hamamatsu Photonics. Frame width = 42 um.

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Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen denominator. Video contrast (Cv) is then expressed as Equation 2, where A is the electronic amplification obtained by setting the gain.

Once the straylight has been compensated for by the offset (pedestal) voltage, the analogue gain of the camera can be adjusted once more to utilize the full range of grey scales in the unprocessed image. iv. Contrast enhancement and resolution of objects The gain in resolution is about twofold. Using the best lenses, where aberrations are negligible, resolution, i.e. the smallest distinguishable distance between two points, is only limited by the size of the spreading of the image points due to diffraction, that is by the size of the Airy disk. One reason why resolution can be somewhat increased by contrast enhancement is that the Rayleigh criterion of resolution, that is defined as a 15% drop between the two peaks that the eye can perceive (Figure 4a), is replaced by the Sparrow criterion (12) (Figure 4c and d) which the video camera can detect. This is applicable to electronic images because they can be enhanced, so that even a slight trough in the intensity distribution of the two unprocessed images (Figure 4c) can be enhanced to give good separation (Figure 4d). v. Contrast enhancement and visualization of objects Objects smaller than the limit of resolution create a blurred image or diffraction pattern known as the Airy pattern whose amplitude (intensity) is very

Figure 4. Improvement of resolution by VEC microscopy. The diffraction pattern (Airy pattern) of a very small object is characterized by a central zero order maximum and smaller maxima of first, second, and higher orders, (a) The overlapping images of two closely adjacent objects (pin-holes) with their summed intensity distribution (dashed) are shown. The two objects are resolved according to Rayleigh's criterion since the central depression is sufficiently deep to be perceivable, (b) A much improved image is obtained by redefining the low intensity (black) end at the position indicated by the horizontal line by applying offset, and subsequently amplifying the signal by applying gain, (c) The same objects are somewhat closer so that they are not resolved according to Rayleigh's criterion, (d) However, if contrast is enhanced as for (b), even in this situation an image can be obtained which shows the two objects separated. Sparrow's limit of resolution is reached when there is no trough between the two peaks.

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Figure 5. Improvement of visualization of subresolution size objects by VEC microscopy. Positive objects (slits) that are larger than (a), equal in size (b), and much smaller (c) than the limit of resolution are imaged by transmitted light (arrows) using an ideal diffractionlimited optical system represented schematically by a single lens. The top panels show the resulting intensity distributions across the images (diffraction disks, Airy disks). Before digitization, this corresponds to the voltage of the analogue video signal, i.e. brightness, along a video scan line. The subresolution size object (c) yields a very low contrast 'image' which cannot normally be distinguished from surrounding noise and therefore remains indiscernible by eye. However, its contrast can be enhanced by applying offset and gain, i.e. applying a negative DC voltage of a magnitude indicated by the dashed line and subsequent electronic amplification. This results in the definition of a new black level (intensity zero) and a higher signal, as seen in (d). As a result of such analogue contrast enhancement, objects much smaller than the limit of resolution (c) can be clearly visualized (d). However, their real size and shape cannot necessarily be inferred from the size or shape of their 'images', such as image (d) of object (c), which is inflated by diffraction to be equal in size to image (b). (Reproduced with permission from ref. 13.)

small, but their size cannot be reduced further (Figure 5c). Usually the Airy rings around larger objects are negligible. If, however, the size of the object becomes smaller than the wavelength of the light used, the diffraction pattern may be larger than the object. In the best lenses aberrations have been made negligible and image quality is only limited by the spreading of the Airy pattern due to diffraction. By applying video enhancement such normally invisibly weak Airy patterns can be visualized. However, if two objects are separated by a distance less than the limit of resolution their diffraction images will merge. Hence, by using contrast enhancement such objects can be visualized (Figure 5d) although they can not be resolved (Figure 4). Using Nomarski-DIC and VEC microscopy, biological structures of 15-20 nm can be visualized, while inorganic materials, such as colloidal gold particles, can be visualized down to sizes of 5 nm and less (see Chapter 12). In DIC microscopy the shadowcast diffraction patterns cancel out for many very small objects located at distances less than the limit of resolution and thus remain 83

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen invisible. This situation is met in many types of small cells and nerve endings densely packed with organelles. vi. Advantages of analogue contrast enhancement (a) Straylight is removable in the video situation by offset (IV). (b) The practical resolution is increased by a factor of about two over conventional microscopy. This is partly because it becomes now possible to use the maximal condenser numerical aperture since the resulting excessive image brightness due to Straylight can be suppressed electronically with offset, and partly because Rayleigh's criterion of the limit of resolution is replaced by Sparrow's criterion. (c) The gain in contrast is sufficient to visualize structures in living cells that are about one order of magnitude smaller than could be detected previously under the same conditions (Figure 2). (d) The AVEC conditions reduce the diffraction anomalies, caused by depolarization at lens surfaces or by residual strain birefringence in the lenses, that produce spurious detail and contrast in conventional polarized light-based techniques (see Section 2.1). vii. Limitations of analogue contrast enhancement (a) Electronic noise is amplified along with the video signal in the enhancement process and may have to be subsequently reduced (see Section 2.3). (b) If the optical system (including the slide and coverglass) contains dust, dirt, or manufacturing imperfections, these will create a fixed pattern of mottle that is enhanced along with the image. This can only be removed by digital processing (Figure 3e) (see Section 2.3). (c) If the illuminating system is poorly designed or incorrectly adjusted for Kohler illumination, the field may be unevenly illuminated, since slight unevenness of the illumination will also be enhanced considerably. Therefore, the requirement of even illumination is much more stringent for VEC microscopy than for conventional photomicroscopy. Note, however, that within certain limits, uneven illumination can also be treated as fixed pattern noise and removed by digital subtraction (Figure 6) (see Section 2.3). 1.3.4 Digital image processing Digital image processing such as digital filtering, background subtraction, or averaging may be performed once the video image has been converted into a digital signal. Many of the digital image processing routines were available long before their value was recognized by microscopists (5, 14). With the rapid development of faster computers many of these routines became available at video frequency. The principles of digital image processing and some of the procedures to be employed will be described below. In VIM and VEC 84

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Figure 6. Correction of uneven illumination (shading) in analogue-enhanced images by background subtraction. If after fixing potential flaws in the optics (see Section 2.3.2) and possibly correcting the shading in analogue mode (see Section 1.3.3) an image as in (a) appears, that shows an annoying hot spot after analogue enhancement (b), subtraction of a specimen-free mottle image (c) will result in an evenly illuminated image (d). This sequence also demonstrates the usefulness of a calibratable scale bar, the timer bar (months to 1/100 sec), and the intensity measurement along a line (see Section 4.2). Specimen is the test diatom Amphipleura pellucida with a known line spacing of 250 nm. Microscope, Zeiss Axiophot, Planapochromat x100, NA 1.25, Hamamatsu Photonics Microscopy System,

microscopy they arc used for rapid pre-processing, that is improvement of the images prior to their storage on tape or disk. It should be noted, however, that individual frames, once stored, can also he subjected to subsequent digital processing and image analysis employing essentially the techniques and image processors discussed in Chapter 8. Following analogue contrast enhancement, the analogue TV signal (a temporal pattern of voltage changes), is digitized so that it can he manipulated by the arithmetic logic unit (ALU) in an image processor. In the most suitable processors, the instructions necessary to carry out a number of different 85

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen arithmetic manipulations are controlled by firmware or by software packages so that an operator can easily process images combining the most suitable procedures for his/her particular operation. The operations themselves are carried out in 'real time', that is repetitively during the intervals between consecutive frames (i.e. 25 or 30 times per second depending on the TV standard used). During digitization the image is subdivided into pixels assuming one out of 256 (8-bit) grey levels. Depending on the complexity of the operations to be performed, one or more frame memories (usually 512 X 512 or 768 X 576 pixels) are required. If averaging or other multiframe procedures are to be used the ALU must be capable of 16-bit operations to avoid truncation errors. The spatial resolution is much poorer than that of a photograph that may have a resolution of approx. 5000 line pairs for 35 mm film. In principle, comparable resolution may be achieved by image analysis systems capable of handling images having a spatial resolution of 4096 X 4096 pixels, but these, if existing, would be extremely expensive. Therefore, additional optical magnification must be applied for highest resolution video microscopy (see Sections 1.5 and 1.6). i. Rolling average or jumping average The rolling average function computes the average of the last incoming image and the previously stored average. This procedure results in an exponentially weighted average with the most recent frames dominating (recursive filtering). In jumping average mode a time average is computed from a pre-defined number of frames and this is displayed for the duration of the accumulation of the next set of frames. Both modes diminish electronic noise in the video signal by the square root of the number of frames averaged. The former smears and de-emphasizes any motion present, while the latter accentuates slow motion. Both are generally used for VIM where photon fluctuations and electronic noise from the imaging device often present severe problems and they are advisable for VEC microscopy when high enhancement is used, that is when electronic noise becomes annoying. The Kalman filter (15) represents another, often better way to produce a continuously displayed image with reduced noise. ii. Mottle subtraction Patterns of image imperfections (mottle) remain in the analogue image when the specimen is defocused or moved out of the field of view (Figure 3c). Consequently, mottle can be stored in a video frame memory and then subtracted from each frame of the incoming video signal (Figure 3d). This operation (mottle or background subtraction) results in a 'clean' image lacking mottle (Figure 3e). The same operation also eliminates inhomogeneities in background brightness (Figure 6), if their contrast does not exceed the range of grey levels, which can be handled by the processor (usually 256). 86

3: Video microscopy Hi. Digital contrast enhancement The analogue-enhanced, mottle -subtracted image may not have sufficient contrast. In this case the image can be enhanced further digitally, e.g. by stretching the histogram of grey levels (Figure 3f). The procedure is analogous to analogue enhancement but the selection is made digitally by choosing that restricted region of the grey levels containing the image information and expanding it to stretch the entire distance from black to white, that is 256 grey levels. This is done by assigning new grey levels to the original ones through the use of an output look-up table (LUT). Please note that analogue enhancement cannot be replaced by digital enhancement. iv. Enhancement of motion by sequential subtraction The analogue-enhanced image can be subjected to sequential subtraction in order to observe and detect only moving elements. This is done by freezing a reference image without taking the specimen out of the field or out-of-focus and then subtracting it from all incoming frames. Subtraction of a (stored) image from very similar subsequent (i.e. the incoming live) images results in blank images in which only moving elements cause image differences and so make their presence known (Figure 7). This is an extremely sensitive means of

Figure7. Selective visualization of moving objects by sequential subtraction, (a) When the reference image taken at time 29:40 sec is subtracted from the incoming video image at time 29:50 sec, i.e. 0.1 sec or five frames later, almost no contrast is visible because all objects remained close to their original location, (b) 2 sec later (31:50 sec) the moving organelles became visible. Each organelle is depicted twice, once appearing as a depression and once in positive contrast. The depression marks the organelle's location at commencement, i.e. the locus where the organelle is now missing, while on the video screen the actually moving objects become visible in positive contrast. In this sequence in a bundle of pike olfactory nerve axons of 0.25 um diameter the movement of mitochondria (elongate) and lysosome-like organelles (round) is observed. Most organelles except one mitochondrion near the centre can be seen moving to the lower right. Microscope, Polyvar 1, Leica/Reichert/Cambridge Instruments, Planapochromat x 100, NA 1.32, Hamamatsu Photonics Microscopy System, Scale bar = 2,7 (um.

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Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen motion detection, but only works satisfactorily with a very stable microscope stand under good temperature control. Any drift in focus or pressure applied to the stage or microscope body may result in a distorted image. This mode gives both the position of the moving object at time zero (frozen and in negative contrast, i.e. a 'missing object') and the live position of the moving object (Figure 7). Distance measurements for velocity calculations can be very conveniently obtained by this technique. v. Interval subtraction This method is an alternative mode of sequential subtraction that is programmed to refresh after predetermined intervals, i.e. a new 'background' image to be subtracted from incoming video images is automatically chosen and stored after a certain, pre-selectable number of frames. vi. Pseudocolour display This process allows the user to systematically or arbitrarily assign colours to various grey levels. This can be very helpful in discerning patterns or small differences in intensity since the human eye distinguishes only about 70-90 grey shades but a multiple of colour shades. Additional digital functions which prove particularly useful for VIM include the following: vii. Frame (or beam) blanking This is another method for improving signal-to-noise ratio (S/N) during image acquisition. Here, the charge pattern on the faceplate of the camera is allowed to accumulate for an extended period of time rather than being 'read out' every 1/25 or 1/30 of a second (video rate). This greatly reduces the amount of readout noise. For example, a typical 16-frame averaging uses 16 images, each containing a certain percentage of readout noise. If, however, the image is allowed to integrate on the camera target for a period equivalent to 16 frames before being read out the amount of readout noise is reduced to l/16th. viii. Spatial filtering By applying various convolution operations the image can be 'filtered' in the spatial domain. This process allows for suppression of noise or the accentuation of high frequency information as in edge detection or image sharpening (see Chapter 8 for details). ix. Arithmetic operations This permits the application of the basic arithmetic operations to a single image or between multiple images. Ratio imaging, where one image is divided by another, is a typical example (see Section 3.3.2). x. Image superimposition The ability to superimpose one image upon another is very useful in many applications. Examples include combining a pseudocoloured fluorescence 88

3: Video microscopy image with the corresponding black and white transmitted light image or combining two images, each of which represents a different fluorescent label, to evaluate their relationship to one another. It is clear that a multitude of additional procedures to accentuate specific features have been developed and are now accessible with programmable digital image processors (see ref. 14 for overview).

1.4 Electronic equipment for video microscopy Once you have determined the functions which will be essential for your research (see Section 1.2) you need to select the appropriate type of videomicroscopical equipment. A generalized scheme is set out in Figure 1. Cameras and image processors are discussed in this section, while comments on suitable ancillary equipment such as recorders and monitors will be found in Section 5. VIM and VEC microscopy differ mainly in the types of cameras appropriate for each technique. A publication by the Centre for Light Microscopy and Imaging and Biotechnology (at Carnegie Mellon University, Pittsburgh, PA, USA) will provide additional useful information for choosing the appropriate camera (16). 1.4.1 Cameras for video-enhanced microscopy For VEC microscopy historically exclusively high resolution vidicon cameras were used, mainly of the Chalnicon, Newvicon, or Pasecon type. If special spectral requirements such as high sensitivity in the infrared, red, or UV range, or extremely low lag are desired, suitable cameras may be selected from various manufacturers, such as Hamamatsu Photonics, DAGE-MTI, Inc., COHU, Inc., and others. Over the last decade CCD (charge coupled device) cameras have improved substantially in terms of resolution and sensitivity. Black and white (B/W) tube cameras with 3/4 or 1 inch tubes are still often used, if images are to be processed in analogue mode. CCD cameras with 1/3 or 1/2 inch chip size are progressively replacing the tube cameras. CCD cameras are offered by a number of manufacturers such as Diagnostic Instruments, Inc., Hamamatsu Photonics Deutschland GmbH, Javelin Systems, Kappa MeBtechnik GmbH, PCO Computer Optics GmbH, Polaroid Export Europe, Princeton Instruments, Inc., Proxitronic GmbH, PTI, Inc., Sony Corp., or Theta Systems GmbH. Colour cameras with either one or three separate chips for red, green, and blue (RGB signal) which are useful for dye discrimination in histology are rarely useful for video microscopy. When selecting a camera make sure that it can be used without automatic gain control (AGC) because gain and offset (pedestal) must be set by the user manually (see Section 2.3, Table 2, step 5). Only if this is possible, can analogue contrast enhancement be performed. The non-uniformity of response across the camera target is called shading. In some low light level cameras this effect can be as high as 20% from one 89

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter S t e f f e n

Figure8. Cameras and camera control units for video microscopy, (a) B/W vidicon camera C 2400-77E 2/3" (Hamamatsu Photonics) and its control unit with gain, offset, and analogue shading correction capability, (b) B/W intensified (CCD camera C2400-81 Hamamatsu Photonics) and its control unit, (c) Chilled B/W (CCD camera C5985 Hamamatsu Photonics) with control unit (courtesy of Hamamatsu Photonics Corp., USA).

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3: Video microscopy region to another. A number of commercially available cameras offer 'shading correction' which allows the user to introduce various combinations of linear and parabolic waveforms to compensate for this non-uniformity. Examples of cameras and controls are shown in Figure 8. 1.4.2 Camera systems for video-intensified microscopy (low light level cameras) Historically, a diverse number of camera technologies have been applied to low light level imaging. For a variety of reasons most of these technologies have given way to two general classes of low light detectors: intensified video cameras and cooled solid state cameras. From an application standpoint these detectors are differentiated from one another by the fact that the former, being intensified, is capable of viewing dynamic specimens; while the solid state device, being an integration-type detector, is, at very low light level situations, suitable only for very slowly moving or static samples. i. Intensified video cameras These devices, as implied by the name, consist of two separate functional components—an image intensifier and a video camera. The image intensifier serves to detect the image, amplify its intensity, and present the resulting image to the video camera so that it may be 'readout' in a systematic format. The most common low light level camera currently utilized in microscopy is the silicon intensifier target (SIT) camera. This design combines an electrostatically focused image intensifier with a silicon target camera tube within a common glass envelope. SIT cameras can provide sensitivities up to 100 times greater than the silicon target camera alone, which itself is considered a sensitive camera. The price to be paid is, however, a considerably lower spatial resolution. The SIT camera is also available in a double intensified configuration known as the ISIT (intensified silicon intensifier target). This camera utilizes an additional intensifier which is fibre-optically coupled to the photocathode of the SIT. This combination provides sensitivity approximately 20-30 times greater than the SIT camera and allows for operation very close to the limits of human vision. Both the SIT and ISIT normally employ a multialkali photocathode which provides spectral sensitivity from 300-850 nm (Figure 9) (11, 17). An alternative approach to the SIT/ISIT design is to couple optically an image intensifier to a video camera. In contrast to the SIT, most image intensifiers employ a phosphor window as the output element, thereby reconverting the electron image back to an optical image for viewing by the video camera. By way of lenses or a fibre-optic coupling the image at the phosphor is focused onto the faceplate of the video camera. A major practical advantage of this design is that it provides flexibility in selecting image intensifiers and video cameras with performance characteristics for specific applications (Figure 8b). 91

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Figures, (a) Sensitivity range of vidicons and low light level cameras relative to incident light intensity. Very sensitive film is included for comparison as well as illuminance levels of moonlit and starlit scenes. Photon counting imager type 1 is with phosphor screen output of the first stage while type 2 is with semiconductor-based position-sensitive detector output, fc, foot candles or lumens per square foot; lux, lumens per square metre, (b) Spectral responses of the three most common types of photocathodes. B, bialkali type; M, multialkali type (S20); S, S1 type.

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3: Video microscopy The major differences among image intensifies revolve around the focusing mechanisms employed and the method of amplification. The simplest configuration is the wafer, or biplanar, type. Its greatest attributes are small size and lack of distortion. Higher performance in terms of gain and image quality can be obtained by incorporating a focusing mechanism and both electrostatic and electromagnetic focusing systems have been employed. The electrostatically focused type is more compact, lightweight, and less expensive. For extremely high intensification requirements these tubes can be configured so that the output phosphor of one is optically coupled to the input photocathode of another. Such cascaded intensifiers can realize luminous gains in excess of 106, when constructed of three or four stages. High gain can also be achieved by providing for electronic amplification within the image tube itself (17). This is made possible by placing a microchannel plate (MCP) between the photocathode and output phosphor. A single MCP can provide electron gains of 103 and multiple MCPs may be used for higher gain requirements. MCP image intensifiers offer similar performance to multiple stage electrostatic focused systems with smaller size, lower distortion, and decreased power requirements (Figure 9a). Combinations of intensified cameras and tube cameras or CCD cameras are available for applications where both highest sensitivity and video rate capacity are required such as for high time resolution Ca2+-imaging (Figure 8b). Such combinations reach extremely high sensitivities down to single photon counting range (Figure 9a) sometimes with extremely wide dynamic ranges (up to nine orders of magnitude). Manufacturers of SIT cameras, intensified tube cameras, and intensified SIT cameras, would include Cohu, Inc., DAGE-MTI, Inc., Hamamatsu Photonics Deutschland GmbH, and Proxitronic GmbH. ii. Cooled solid state detectors Recent advances in the development of CCDs have been very promising with regard to their application in low light microscopy. While technically not an intensified camera, the CCD camera has the ability to obtain images at low light levels. Being inherently high quantum efficiency devices with respect to the ability to convert photons into electrons, high sensitivity is achieved primarily by cooling and slow readout. Cooling the CCD (Peltier cooling to around 0°C or external cooling to - 125°C) dramatically reduces the dark current as a noise component. The slow readout further reduces the noise associated with high bandwidth electronics. Low light images are integrated directly on the chip in much the same way as an extended photographic exposure. High quality slow-scan CCDs offer excellent geometry, photometric accuracy, and large dynamic range and are, therefore, the cameras of choice in microscopy for the quantitative imaging of static low light samples. These cooled CCD detectors require their individual special imaging board to guarantee precise readout of the CCD chip which is achieved only at relatively slow rates. While formerly one image required up to several seconds, 5-10 Hz cameras are 93

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen presently available and video rate is expected to be possible in the near future. Hamamatsu has recently developed cooled CCD cameras (e.g. Hamamatsu C5985) achieving readout rates of seven images per second (Figure 8c). This camera type could, therefore, provide a good alternative to SIT cameras currently used in dynamic VIM. Manufacturers of suitable slow-scan CCD cameras are Diagnostic Instruments, Inc., Hamamatsu Photonics Deutschland GmbH, Javelin Systems, Kappa Messtechnik GmbH, Optronics Engineering, PCO Computer Optics GmbH, Photometries Ltd., Photonic Science Ltd., Princeton Instruments, Inc., PTI, Inc., Theta Systems GmbH, and others. For further information about cooled CCD cameras see refs 11, 16, and 18. Because cooled CCD cameras are controlled via a specific computer interface (image acquisition board), one should first decide on the computer platform (Macintosh-, PC-, or Unix-based) to be used with the camera system. Most manufacturers will offer solutions for both Macintosh and PC computers. By deciding on a particular computer platform one should, however, only consider computers with a fast processor and sufficient RAM (minimum of 64 MB) and hard drive memory (minimum of 2 GB). Image acquisition requires dedicated software packages compatible with a variety of slow-scan cameras and their specific interface boards, such as MetaMorph (Universal Imaging Corp.), IPLab Spectrum (Scanalytics, Inc.), NIHImage (public domain from NIH, Bethesda), AxioVison (Carl Zeiss GmbH), or the packages obtainable from Data Translation, Inc. or Vaytec, Inc. Some of these software packages also control video recorders in a microscope workstation and can be used to improve image quality. PhotoShop (Adobe Systems, Inc.) and Paint Shop Pro (Jasc Software, Inc.) are general software programs to improve image quality, but are not sufficient to control sophisticated cameras such as cooled slow-scan CCDs. A detailed comparison between cooled CCD and SIT cameras is provided in ref. 11. Hi. Practical considerations in choosing a low light level camera system (a) Sensitivity. Clearly the most important consideration in selecting a camera for low light imaging is sensitivity. A wide variety of cameras is available with sensitivities ranging from applications in DIC and phasecontrast microscopy down to the single photon level. Sensitivity is typically measured by illuminating the camera target with a known quantity of tungsten light and measuring the resulting output current of the camera. The data is expressed in amperes/lumens and is plotted in a log-log fashion as the 'light transfer characteristic'. This provides a useful way to compare various systems but this definition can be misleading when applied to low light cameras. It does not take into account the noise component of the output signal and, being based on tungsten light, is heavily biased toward red-sensitive photocathodes. Figure 9a illustrates the typical sensitivity range of a number of low light level cameras. 94

3: Video microscopy (b) Spectral response. The spectral response of intensified cameras is determined by the window material and the type of photocathodes. The most common photocathode is the multialkali type (S20) which has a peak sensitivity at 420 nm and provides usable sensitivity to approximately 800 nm. The spectral responses of three common photocathodes, multialkali, bialkali, and SI, are shown in Figure 9b. It should be noted that the extended red-sensitivity of the S20 and S1 types is accompanied by higher noise levels (thermal noise). While not a problem in most situations, at very low light levels one is advised to select the bialkali type unless this extended sensitivity is required. (c) Resolution. As noted, high resolution and high sensitivity tend to be mutually exclusive characteristics. This relationship is due to noise. Noise in low light level systems can generally be classified as signal-independent or signal-dependent. Signal-independent noise is essentially present in a fixed amount, in both the absence or presence of light. It arises from thermal noise of the intensifier, the camera target, the video-amplifier, etc. This is the predominating type of noise at relatively high light levels. As light levels decrease signal-dependent noise becomes the dominant component. At low photocathode illumination levels, resolution is primarily limited by the finite number of photoelectrons released at the photocathode—the so-called photoelectron, or quantum, noise. It is this latter noise which accounts for the decrease in resolution as light levels decline. Resolution, therefore, should be evaluated as a function of light intensity and preferably at a specific wavelength when comparing for possible photocathode differences. (d) Lag. Lag, or dynamic response, describes the camera's ability to respond temporally to changes in light intensity. If the system is to be used for the imaging of rapidly changing specimens, a camera with low lag characteristics should be selected. Lag in an intensified camera is due primarily to the readout video camera, therefore utilizing a low lag camera such as a saticon or solid state will greatly improve this characteristic. One should be aware that certain camera types retain a considerable part of the information (up to 30%) from one frame to the next. For more details see ref. 17. Microscope/camera combinations suitable for both VIM and VEC microscopy are depicted in Figure 10. 1.4.3 Analogue image processors Analogue image enhancement is the step in video microscopy whereby most of the possible image improvement is gained. Analogue functions (Figure 1) are either incorporated into the camera control units (Figure 8) or into the frame grabber boards. Essential for video microscopy is the capability to manually set gain and offset within a wide range. Cameras with automatic gain 95

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steff en

3: Video microscopy Figure 10. (a) Video microscopy set-up based on a Zeiss Axiophot. The monitor for the processed image is seen behind the Hamamatsu control unit C2400 for SIT camera (front) or Newvicon camera (back). The control unit can be used for either one of the two cameras unless simultaneous use of both cameras is needed (courtesy of Hamamatsu Photonics Corp. USA), (b) Multi-purpose video microscopy workstation using a Nikon Diaphot 300. (1) Newvicon Camera (Hamamatsu C2400-08); (2) fan heating system (Nikon) for 37°C box around the stage; (3) mercury HBO100 lamp house; (4) Xenon XBO100 lamp house with remotely controlled shutter (Uniblitz); (5) low light camera SIT (Hamamatsu C2400-07); (6) slow-scan camera (Photometries SenSys); (7) control unit for SIT camera with ARGUS 10 and control unit for Newvicon camera with ARGUS 20 (Hamamatsu Photonics); (8) video monitor, Uniblitz control unit and time lapse video recorder (Panasonic AG6730 SVHS); (9) colour printer (Tektronix Phaser 350); computer (Macintosh PowerMac). The set-up can be used simultaneously for VIM and VEC microscopy because the high resolution (5) and low light (6) cameras are connected to a Nikon multi-image module containing a proper dichroic beam splitter.

control (AGC) only or gain and offset adjustable only in a narrow range or by small set screws on the back of the camera are, therefore, unsuitable. Some controllers provide additional analogue processing features for image preprocessing such as shading correction, intensity line scan, and others. As a rule, cameras should fulfil the requirements mentioned in Section 1.4.1 especially when offered for microscopy applications. Suitable equipment is provided by Colorado Video, Inc., DAGE-MTI, Inc., For-A Company Ltd., Hamamatsu Photonics Deutschland GmbH, or Optronics Engineering. 1.4.4 Compact digital image processors Digital image processors for video microscopy may come as (i) compact stand-alone systems, which also may be (ii) externally driven together with other microscopy devices (stage motor, filter wheels, etc.) by a PC, or (iii) fully PC-based systems (see Section 1.4.5). Compact digital image processors are the minimal requirement available at reasonable cost for those who need digital image processing in real time in order to obtain the desired image quality, but do not intend to do further digital image analysis (e.g. morphometry) of single frames with the same system. Be aware, however, that some of these systems do not include the analogue enhancement feature, which is indispensable, if cameras without their own analogue enhancement capability are to be used. The processors most suitable for our purpose should include digital enhancement, real time subtraction with simultaneous frame averaging functions, plus additional potentially useful features. Within this group ARGUS 10 and 20 (Hamamatsu Photonics) are the most widely used device (Figure 11). A few dedicated systems are available which allow all these essential real time functions and provide the user in addition with a fair number of measurement functions and some flexibility to manipulate and analyse a set of stored images. To this end these stand-alone systems may be connected to an external PC and thus allow to access software libraries for specific applications. These 97

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Figure 11. Compact image processor for analogue contrast enhancement and digital processing, (a) Example of the mouse-driven on-screen menu of the Hamamatsu ARGUS 20 (b) that permits real time image improvement by a great variety of both analogue and digital functions together with additional measuring functions (bottom).

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3: Video microscopy systems can provide software for real time image processing (e.g. for ratio imaging) and for controlling additional hardware such as video recorders, filter wheels, or special cameras. In combination with remotely controlled high speed shutters (e.g. Uniblitz) and video recorders important features arise: time lapse recording (each n-th frame), or alternate recording from two cameras, such as continuous DIC recording with one and fluorescence with a second camera. Generally, previously recorded sequences played back from a recorder can equally be processed with these systems. Examples in this category are the ARGUS 20 as a stand-alone system or the PC-driven ARGUS 50 series of Hamamatsu Photonics and the systems from Optronics Engeneering or Datacube, Inc. 1.4.5 PC-based digital image processors The improving speed and storage capacity of PCs now in use allows them to be used with appropriate analogue-to-digital converters (A/DC), which are often integrated in sophisticated frame grabber boards, to perform the required steps for video microscopy. A whole industry offers image processing boards and software packages that perform both image acquisition and processing not only with single frames but also at video rate. Examples of companies producing frame grabber cards are Datacube, Inc., Data Translation, Inc., Fast Electronic GmbH, HaSoTec GmbH, Imaging Technology, Inc., Matrox Electronic Systems Ltd., Scion Corp., or Silicon Graphics, Inc. For high performance tasks such as pixel point processing in single frames (digital spatial filters, Fourier transforms) or two- and more-frame processing (ratio imaging, 3D display) dedicated on-board processors are widely used, although the rapid progress in computer performance makes fully software-based systems of astounding capabilities already possible. A dedicated system would have to include all the above mentioned processing functions and the capacity of controlling ancillary devices such as a variety of commonly used cameras including slow-scan CCD cameras, focus motor, shutters, stage drive, and motorized filter wheels or filter sliders. Very important is the capability to synchronize one or two recorders with cameras and shutters in order to allow parallel recording and to protect the fluorescent specimens from photobleaching by recording only a few frames after every n-th minute in the fluorescence channel. Such regimes can be very helpful because they permit long-term experiments without extensive photobleaching. The system should also offer a number of software routines needed for biological video microscopy, such as for ratio imaging, organelle tracking, or multiple fluorescence. The display functions should include playback of stored video sequences in real time and in time lapse mode, i.e. selecting each n-th frame and assembling it into video sequences (video-clip). For post-processing the availability of point, kernel, and frame operations for further image improvement and extraction of quantitative data (data analysis, morphometry) are desirable. 99

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen Several companies offer fast or video rate image processing systems working on PC-, VME-, or UNIX-based platforms that combine almost all of the above mentioned features (e.g. Bitplane AG, Improvision, Inc., Inovision Corp., Media Cybernetics, NIH-RSB, Scanalytics, Inc., Silicon Graphics, Inc., Stemmer Imaging GmbH, Universal Imaging, Inc.). Some companies also offer software packages capable in addition of very useful device control (video recorder, light shutter, etc.) and microscope automation features (Carl Zeiss GmbH, Inovision Corp., Scanalytics, Inc., Universal Imaging Corp., VayTec, Inc.). A high-end IBM compatible PC system is that developed by Universal Imaging Corp. using their software package MetaMorph (19). Alternatively a Macintosh-based system as that depicted in Figure 10b could be assembled from available components. In our case this consists of a Nikon Diaphot 300 microscope with DIC optics and epifluorescence equipped with four different cameras attached to front, side, and two upper ports. We are currently using a 35 mm Nikon F-601 camera (front port), a SIT camera (Hamamatsu C2400-08) with controller connected to an ARGUS 10 image processor (side port), a NIKON multiimage module (upper port) with two cameras, a Newvicon camera (Hamamatsu C2400-07) with controller connected to an ARGUS 20 image processor, and a SenSys cooled digital CCD camera (Photometries Ltd.) with LG-3 computer interface card (Scion Corp.). The multi-image module allows by using an appropriate dichroic mirror to use epifluorescence and DIC simultaneously (20). In our set-up two ARGUS image processors (ARGUS 10 for SIT camera and ARGUS 20 for Newvicon camera) are used for digital enhancement, subtraction, and frame averaging. Image acquisition is performed from the ARGUS processors either by a Panasonic S-VHS time lapse video recorder or directly by a Power Macintosh 9600 (350 MHz/128 Mbyte RAM/3 GB HD) equipped with the LG-3 frame grabber (Scion Corp.), which captures greyscale images and provides four input channels that can be used for image acquisition from different cameras. The S-VHS time lapse video recorder (Panasonic, model AG-6730) is controlled via a RS-232C interface adapter from the computer serial port. This combination is able to count and access individual frames' positions on the tape. The image acquisition software used is IPLab Spectrum (Scanalytics, Inc.) which supports, in our case, (i) the LG-3 frame grabber, (ii) the slow-scan camera (SenSys, Photometries Ltd) interface hardware, (iii) the video tape recorder control, and (iv) the shutter control (Uniblitz). Using the IPLab Spectrum software in combination with the appropriate hardware it is possible to control several video cameras and video recorders and capture images onto the computer. Using a microscope set up with a Newvicon camera and a SIT or cooled CCD camera one can record DIC images and fluorescence images simultaneously. If time lapse series of fluorescence images have to be recorded it is highly recommended that a Uniblitz 100

3: Video microscopy shutter be included to protect fluorescent specimens from photobleaching during periods of no recording. All of these features can be controlled with the IPLab Spectrum software in combination with the LG-3 frame grabber card (Scion Corp.). Examples for studies that can be performed with this set-up include: (a) Tracking of organelle movement along fluorescent actin filaments in vitro. Fluorescent actin filaments labelled and stabilized with rhodaminephalloidin are visible in the fluorescence channel and moving organelles in the transmitted light DIC channel (21). (b) Tracking of organelle movement along fluorescent microtubules in living cells. Microtubules labelled by microinjection of rhodamine tubulin into the cells are visible in the fluorescence channel and moving organelles in the transmitted light DIC channel (see also refs 7 and 20). 1.4.6 Single function processors While the above mentioned devices are more or less multifunctional, sometimes only one or two functions may be required. In this case it may be cheaper to obtain unifunctional, hard-wired instruments for analogue enhancement, distance measurement, perimeter measurement, time/date generation, or intensity measurement along a scan line etc. Such devices can be obtained from, for example Colorado Video, Inc., For-A Company Ltd., Hamamatsu Photonics, or Optronics Engeneering.

1.5 Considerations on the microscope For all types of video microscopy, a high performance research microscope should form the basis of the optical imaging system. Due to the unusually high magnifications used for video microscopy special attention has to be paid to bright illumination, optimized light throughput, and high mechanical microscope stability. A heavy stand for the microscope is recommended in order to reduce vibrations and internal movements resulting from temperature changes. This is especially important when heavy cameras are used like those designed for VIM and when using highest magnifications. It should be noted that a 1 um displacement in the specimen is registered on the video screen at typical magnifications for VEC microscopy as 1 cm. In some cases vibration isolation tables may therefore be required. 1.5.1 Illumination Video cameras must work near the saturation end of their dynamic range in order to enable one to utilize their contrast enhancement advantages. When planning high magnification VEC microscopy xenon or mercury arc lamps are required, in most cases, to provide enough light, so that the cameras work near the saturation end. Furthermore, it may be essential that all unnecessary 101

Dieter G. Weiss, Willi Mails, Robert A. Wick, and Walter Steffen components such as filters and diffusers are removed from the light path and the lamp is optimally adjusted. Only at lower and intermediate magnifications or in some microscopes that are optimized for very high light transmission 100 W halogen or tungsten lamps may be sufficient even for VEC-DIC. We have seen the best results with the HBO 50W DC mercury lamp, which has a short intense arc, in combination with the Axiomat and the Axioskop (Carl Zeiss GmbH), the inverted microscopes IM and ICM (Carl Zeiss GmbH) and the Polyvars (Reichert/Cambridge Instruments). The 100 W or 200 W mercury arc lamps also worked well in most cases, although sometimes the former may have a too small bright centre, thus producing a central hot spot (Figure 6), while the latter may not be bright enough because the light is too much spread. Great care should be taken to set the illumination so that it is uniform and filling the NA of the condenser. If this is not possible with the arc lamp used, a fibre-optical scrambler may be inserted (see ref. 2, p. 127). Due to the fact that after contrast enhancement minute changes in lamp intensity may result in transitions from well modulated to bright white or black images, stabilized DC power supplies may be unavoidable. With arc lamps it is advisable to use a narrow-band green interference filter (e.g. the Hg line 546 ± 10 nm for mercury lamps) for optimal DIG results and to protect cells from blue light. In this case it is also necessary to protect the interference filter, the polarizers, and the cells from heat and UV light with at least one piece of each of the following filters: a UV filter, a heat-reflecting filter, and a heat-absorbing filter. 1.5.2 Optimized light gathering and throughput All VIM and most of the high magnification VEC microscopy applications are characterized by the limited amount of light available. Therefore, by far the two most important considerations involve optimizing the microscope for light gathering ability and the efficient transmission of light through the optical path. The light gathering ability of the microscope is a direct function of the numerical aperture (NA) of the objective lens. While factors such as an objective's working distance often necessitate using a lens with lower than maximal aperture for a specific magnification, whenever possible the highest available numerical aperture should be utilized. The importance of this point is clearly appreciated when one considers the relationship of magnification, numerical aperture, and light intensity. In a system where the illuminating aperture is equivalent to the objective's acceptance aperture (such as in epifluorescence), intensity (/) is proportional to the fourth power of the numerical aperture. Further, light intensity is inversely proportional to the square of the magnification:

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In comparing two X 40 objectives with apertures of 0.9 and 0.5, one sees that the higher numerical aperture objective captures over ten times more light than its lower numerical aperture counterpart. The demand of high NA objectives necessitates the use of oil or water immersion objectives and condensers whenever possible. It should be clear to the reader that the ultimate performance of VEC-DIC (e.g. visualization of microtubules) will require a X 63 or X 100 objective with NA 1.3 or 1.4 and a oil condenser with NA 1.4. As a general rule, in lenses of similar design, the numerical aperture tends to increase with magnification. The above relationship clearly dictates that when using additional magnifying optics, such as optovars, relay optics, or projection eyepieces, one should maximize the numerical aperture and magnification of the objectives, and minimize the magnification of the intermediate optics, since these optics contribute nothing to the light gathering ability of the system. Therefore, if faced with the choice of using a X 40 objective in combination with a X 10 camera relay lens versus a X 25 objective and X 16 relay lens, the former combination will prove far more efficient despite the fact that both systems deliver a final magnification of X 400. Regarding the second major consideration, we have to remember that the efficiency of light transmission within the optical system is primarily a function of the transmission properties of the objective lens and the number of optical elements in the system. Interestingly, there are often significant differences in the transmission characteristics of objectives of different designs (e.g. fluorite versus apochromatic) and from different manufacturers, even when the numerical aperture and magnification are identical. If possible, it is recommended that a number of lenses be evaluated with regard to the specific wavelength(s) that will be utilized. Of equal, if not more, importance, is the issue of intermediate optical elements. Uncoated lens surfaces typically reflect 4—5% of the light incident upon them. And while it is true that research quality microscopes generally employ anti-reflection coatings, these cannot completely eliminate losses due to reflection and the cumulative loss associated with the complex light paths can be substantial. Therefore, all unnecessary optical elements should be removed from the light path. If this is not possible, the use of a microscope with a simple light path should be considered. When planning fluorescence work, make sure the optics transmit the short wavelength light which is often required for the excitation of dyes, for example 340 nm for Fura-2 used for video measurements of Ca2+ concentrations. The requirement of the highest quality optics is sometimes even less stringent, since in most cases monochromatic light is recommended anyway, and since 103

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen often only the central area of the field (~ 1/3 to 1/2) is picked up by the video camera. Wide-field optics are usually of no particular advantage. 1.5.3 Special considerations for AVEC microscopy For AVEC microscopy (see Section 2.1) two special considerations arc important: (a) De Senarmont compensator. In order to set the recommended bias retardation of 1/9 of a wavelength (546 nm)a de Senarmont compensator set-up is required (see Section 2.3.4). Most biological DIC and POL microscopes have only the Brace-Kohler type or other compensators or analysers that allow only the introduction of bias retardations of 1/100 or 1/50 of a wavelength but not 1/9. Nikon is the only manufacturer to offer a dc Senarmont compensator for AVEC microscopy routinely with their Eclipse microscopes. It consists of an additional quarter wave plate (y/4 plate) which converts the polarizer or the analyser into a compensator (22). The \/4 plate should match the wavelength used, preferably the green Hg line (546 nm). Other microscope manufactures will, however, have the proper parts in their mineralogical programmes (Figure 12).

Figure 12. Parts for AVEC-DIC microscopy with de Senarmont compensation as built into a Zeiss Axiophot microscope. (1) Objective Wollaston prism; (2) 1/4 wave plate; (3) X 1.6 magnification optovar instead of slider for the fluorescence filter blocks; (4) rotatable calibrated polarizer used as analyser; (5) additional magnification changer x 2.5. With this setting an empty camera attachment tube can be used. Alternatively, a fixed x 4 camera adapter tube can be used instead of the two optovars to obtain the additional magnification. The polarizer and Wollaston prism in the condenser are not visible,

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3: Video microscopy (b) Short shear Wollaston prism. In DIC microscopy visual contrast generation depends on the shear introduced by the pair of Wollaston prisms. If, however, the shear is too large, i.e. larger than the resolution, not only the bright and dark shadow-cast margins that make objects visible at high contrast, but also annoying slightly shifted double images will appear. For work with the eye best contrast is achieved at a shear just below its limit of resolution (i.e. 200 or 220 nm). For video microscopy that is higher in resolution, so that 200-220 nm are truly resolvable, shorter shear prisms are required to make use of the gain in resolution. Therefore, shorter shear sets of Wollaston prisms are desirable to fully utilize the advantages of AVEC microscopy. Nikon is the first manufacturer to acknowledge the importance of special 'short shear' prisms for VEC microscopy as it offers two sets of prisms with their Eclipse microscopes: 'short shear' prisms with low visual contrast but higher resolution with video enhancement and 'medium shear' prisms for high contrast observation by eye but lower resolution with VEC techniques. 1.5.4 Additional magnification For video microscopy we need considerable additional optical magnification. This is due to the fact that because of the relative small number of video lines the video image has a spatial resolution which is much lower compared to that of a photomicrograph or that of the microscope optics. In order to fully utilize the resolution of the microscope we have to make sure that it is the limiting component rather than the video system. This means that we have to magnify the specimen onto the target of the video camera far beyond the magnification which is normally considered useful in conventional light microscopy. For a detailed discussion of object resolution matching the CCD pixel size see ref. 23. Whenever possible, this should be reached by the use of higher power objectives, but when subresolution objects are to be visualized with the X 63 and X 100 oil immersion objectives, additional magnification of X 4 to X 6.3 is required. This can be achieved either by a zoom system as in the Zeiss Axiomat and the Zeiss Axio series, or by a X 2 or more additional magnification changer (optovar-type) and/or a high photo eyepiece (X 16 or X 25) plus projective lens (50-63 mm) as in most other microscopes, e.g. IM, ICM, and Axio series (Carl Zeiss GmbH), Orthoplan and Aristoplan (Leitz), and Polyvar (Reichert/Cambridge Instruments). Alternatively a combination of two X 2 or one X 4 extension tubes may be used (Zeiss, Leica, Nikon). As a rule of thumb, when objects at or below the limit of resolution are to be observed on a medium-sized monitor, the desirable maximal useful magnification should be such that the field of view portrayed on the monitor is 15-30 u,m wide, or the final magnification is around X 8000 to X 15000. 1.5.5 How to interface microscope and camera One should ensure that 100% of the light can be directed to the camera. If the microscope has a fixed 'split' (80/20% or 50/50%) between the binocular and 105

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen camera port, it may be advisable to have a 100/0% reflecting mirror installed on the slider in place of the existing one. The system projecting the images onto the camera target classically consisted of the ocular and the camera objective lens in close contact. Both consist of several lens elements and often some surfaces are close enough to the image plane to introduce very disturbing mottle from dust, grease, and nonperfect coatings. Oculars or projecting oculars are, however, indispensable in all cases where part of the correction of optical aberrations is taken care of in the objective lens and part in the ocular. Several manufacturers have switched to totally internally corrected objective lenses, so that microscopists have much more freedom to use alternative arrangements. In such optical systems the tube lens directly projects an intermediate image onto the TV target through an empty connecting tube. This yields very good images with little mottle. For highest magnifications, this arrangement requires that the additional magnification of at least X 4 is reached by a Galilean-type telescope in the parallel, infinity corrected, light path, or by an additional magnification changer ('optovar'), or by a combination of both. Another connection, which creates less mottle but requires considerable bench space, is to mount the camera to a lateral exit with a high power eyepiece and to project the intermediate image directly onto the camera target. The desired magnification is adjusted by sliding the camera to and from the microscope at a 10-30 cm distance. For biological microscopy it is very important to note that monochromatic DIC and fluorescence images of different wavelengths can be sent to two different cameras simultaneously by using a proper dichroic beam splitter (20). Using the multi-image module (Nikon) (Figure 10b) one can increase the number of detectors mounted simultaneously. By using an appropriate dichroic mirror it is possible to direct all the emitted light from the epifluorescence to a SIT or slow-scan camera (wavelength 1) while all the light from transmitted light DIC goes for simultaneous imaging of fine detail to a Newvicon camera (wavelength 2). Intermediate magnification lenses (X 1.25, X 1.5, X 2.0) inside the multi-image module and the C-mount zooming adapter (Figure 10b, upper left) allow the fluorescence and transmitted light images to be adjusted to the same magnification independent of camera type and target size.

2. High resolution: video-enhanced contrast microscopy 2.1 Different types of video-enhanced contrast microscopy Video contrast enhancement of microscopic images using bright-field, darkfield, anaxial illumination, or fluorescence techniques is very straightforward 106

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Figure 13. Dependence of intensity (image brightness) and contrast (see Section 1.3.3) of DIC images of phase objects on phase retardation. Phase retardation is introduced and varied by laterally displacing a Wollaston prism or by setting the de Senarmont compensator away from extinction. This converts positive and negative phase gradients of specimens to contrast, thus producing highlights (/H) and shadows (/s) relative to a neutral grey background (/B). A phase shift of n/2 radians corresponds to \/4, i.e. 1/4 of a wavelength. It can be seen that theoretically the image contrast is highest at the position n/2 (vertical arrow). Due to straylight the image background (/B) is of considerable intensity at this setting and has to be compensated for by the addition of a negative voltage (offset). For reasons discussed in the text, smaller retardations (1/9 of wavelength) are recommended. (Reproduced with permission from ref. 4.)

and can generally be described by the term 'VEC 'microscopy'. Allen (3, 4) and Inoue (1) simultaneously described procedures of video contrast enhancement for polarized light techniques which differed considerably in their approach but yielded very similar results. There is a need to distinguish clearly between the two strategies in order to avoid confusion. Allen named his techniques 'Allen video-enhanced contrast' differential interference contrast and polarization (AVEC-DIC and AVEC-POL, respectively) microscopy. These techniques involve setting the analyser and polarizer far away from extinction in order to gain a high specimen signal IS (see Section 1.3.3). Allen suggested the use of a de Senarmont compensator set-up (3, 4, 22) which is comprised of a 1/4 wave plate (specific for the wavelength used) in front of a rotatable analyser (Figure 12). He recommended a bias retardation of 1/4 to 1/9 of a wavelength away from extinction with 1/9 as the best compromise between high signal and minimal diffraction anomaly of the Airy pattern. The enormous amount of straylight (IB) introduced at this setting (Figure 13} is removed by adding the appropriate negative offset voltage. 107

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen The technique recommended by Inoue, which in this chapter is called IVEC microscopy for the purpose of distinction, aims to reduce straylight and diffraction anomaly arising mainly from curved lens surfaces (Table -1) by employing extremely strain-free objectives and the rectifying lenses developed by Inoue (24). The latter were commercially available some time ago and only for a few microscopes, such as some lines of Nikon. Inoue's microscope (2, 24) is optimized in such a way and works at a polarizer setting very close to extinction, which cannot be used for VEC microscopy with most other instruments because insufficient light for near-saturation is passed to the camera. In IVEC microscopy, straylight is not admitted since the polarizers stay close to extinction and the special rectified optics further reduce the straylight. Consequently, filters to reduce the brightness are not required. On the contrary, a bright arc lamp, ideally with a fibre-optic illuminator, is necessary to saturate the camera. The AVEC technique electronically improves optical images of low contrast and high straylight content due to 'non-optimal' optical arrangement, while in IVEC microscopy no compromise is made regarding the optics and consequently less demanding electronic steps are required to rescue the image. The AVEC technique is, however, the only one which can be used with any good research microscope equipped with commercial film polarizers. The proper compensator setting can be experimentally evaluated between 1/100 and 1/4 of a wavelength within the ability of ones illuminating system to nearly saturate the camera. Best resolution is achieved on theoretical grounds at 1/9 of a wavelength (25) and best visualization (highest contrast) has been reported for 1/15 of a wavelength in some microscopes (26). However, according to Allen the dimensions of an object imaged at the latter setting may differ considerably, if it is orientated in different directions due to the diffraction anomaly of the Airy pattern (3, 4).

2.2 Sample preparation Samples used in conventional light microscopy can also be used for video microscopy. Live cells from tissue cultures should preferentially be grown on a coverglass. The specimen's region of interest should be close to the coverglass surface, where the best image is obtained. If highest magnifications are intended, it may be found that the optics can be adjusted for Kohler illumination only at this surface and about 20 um below (upright microscope), since high magnification objectives are usually designed for optimum imaging of objects at a distance of 170 nm from the front element (No. 1 glasses, approximately 170 um thick). To identify objectives for which this distance and the presence of a 170 um thick coverglass is critical, look for the '0.17' engraved or refer to the manufacturer's data sheet. If thick cells or other extended specimens must be observed with such objectives, thinner glass slides (0.9 instead of 1 mm) may have to be used instead of the regular ones to allow the setting 108

3: Video microscopy

Figure 14. Microscope flow chamber suitable for highest magnification VEC microscopy with upright and inverted microscopes and for superfusion of live cells or suspension specimens. (1) Metal frame of the size of regular microscope slides made of brass or aluminium about 2 mm thick and absolutely flat with an insert to hold a 24 x 50 mm coverglass (2), The coverglass can be secured to the metal frame by placing small drops of VALAP (3) at the corners of the insert. (4) Small coverglasses or thin adhesive tape can be used as spacers (e.g. double sticky Scotch TapeR). (5) Coverglass (e.g. No. 1; 24 x 24 mm). (6) Filter paper wick to induce medium flow (arrow).

up of Kohler illumination with oil immersion of both condenser and objective lens. If necessary, No. 0 coverglasses (80-120 um thick) can be used (both glass slides and coverglasses are available from O. Kindler GmbH or Clay Adams Co.), but it should be noted that under the latter condition image quality may be impaired. The use of a slide preparation made of two coverglasses mounted in a frame as depicted in Figure 14 would serve the same purpose. For imaging deep within an aqueous sample, water immersion objectives must be used. Special high NA objectives for working with aqueous specimens have recently become available from manufacturers for confocal microscopes. These lenses combine high numerical aperture (e.g. x 60, NA 1.2) with correction for 'extreme' working distances of up to 220 um below the coverglass (27). Note, however, that, if regular oil immersion objectives are focused through water (the specimen) rather than through glass and oil only, spherical aberration will be introduced. The use of monochromatic illumination or of immersion oil of different refractive indices (R. P. Cargille Laboratories) is recommended to overcome this problem. Aqueous samples have to be prevented from drying out by completely scaling the coverglass to the slide. Nail polish may be used or, if live or solventsensitive specimens, such as microtubules, extruded cytoplasm, or cultured cells are observed, VALAP is to be used. This consists of equal parts by weight of vaseline, lanolin, and paraffin (melting point 51-53 °C); it liquefies at around 65°C and is applied around the coverglass with a cotton tip applicator. If the specimen is in suspension (e.g. organelles), no more than 5-10 ul aliquots should be used with 22 X 22 mm coverglasses in order to produce very thin specimens (~ 10 um) for best image quality. 109

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen If working with an inverted microscope, the slide has to be inverted with the coverglass underneath. With most microscope stages this will interfere with the VALAP sealant and flat positioning of the slide will not be possible. It is recommended instead to use a metal frame of the size of a regular slide and 0.8-1 mm thick. A larger coverglass is attached with VALAP so that it covers the opening. After the sample has been applied, the top coverglass of regular size and thickness is added with or without spacers made of coverglass or adhesive tape and sealed (Figure 14). If thick specimens such as tissue slices, vibratome sections, or nerve bundles are to be observed at high magnifications that require immersion, only contrasting techniques that allow optical sectioning such as DIC, Hoffman modulation contrast, or anaxial illumination (28) (including Varel contrast) are recommended. Only the first 10-20 (um behind the coverglass will yield perfect images. However, even if expensive long working distance water immersion objectives are not available, part of the loss of quality can be compensated for by video enhancement with good results down to a depth of 100 (um or more, with respect to both visibility and contrast, sometimes also to resolution (28). The opacity of live tissue such as brain slices is greatly reduced when light of 700-800 nm is used because they are transparent for near-infrared or infrared light (29).

2.3 Procedure for image generation Because VEC microscopy requires some steps which are different from conventional microscopy, image generation is discussed here in some detail (Table 2). Steps 1-5 of the procedure yield the image if only analogue enhancement is required. The continuation leads to highest resolution and visualization of submicroscopic objects. The procedure chosen is basically that used for AVEC-DIC and AVEC-POL, but if these are not required step 4 can be disregarded. 2.3.1 Focusing the specimen: step 1 Find the specimen preferably by looking through the oculars or, alternatively, by looking at reduced magnification at the monitor. If the entire specimen consists of subresolution size material (density gradient fractions, microtubule suspensions, unstained EM sections) (see Figure 3) it will be difficult to find the specimen plane. Use a relatively dark setting of the condenser diaphragm and/or polarizers or prisms and look for brightly shining contaminating particles. If there are none, routinely apply a fingerprint to one corner of the specimen side of the coverglass and use this for focusing. 2.3.2 Adjusting Kohler illumination: step 2 After finding a coarse setting for the illumination, the desired plane for the specimen is selected exactly. Then the condenser is finely adjusted, but now in relation to the image on the monitor (make sure the light is reduced to avoid 110

3: Video microscopy Table 2. Steps in AVEC-DIC microscopy Step

Equipment

Manipulation

Result

1a 2

Microscope Microscope

Image appears Image improves

3

Microscope

Focus specimen Adjust microscope correctly for Kohler illumination Open condenser diaphragm fully

4

Microscope

5

Camera

6

Microscope

7

Processor

8

Microscope

9

Processor

10

Processor

11

Processor

Set compensator up to 20° from extinctionb Analogue-enhance by manually adjusting gain and offset

Optical image becomes too bright Optical image worsens

High-contrast video image with often disturbing mottle pattern appears Object disappears, mottle Defocus or move specimen laterally out of field of view remains Average and store mottle image, then Absolutely homogeneous, subtract mottle image from incoming light grey ('empty') image appears video images Clear image appears; if Return specimen to focal plane contrast is weak go to step 9 Contrast becomes optimal; Contrast-enhance digitally if pixel noise is high go to (histogram stretching) step 10 Use rolling or jumping averaging or Clear, low noise, and high digital filtering contrast image appears Spatial filtering using various masks Sharpened image or highlighting special aspects of image contents

aBefore step 1, one should set the 'brightness' and 'contrast' controls of the monitor showing the processed image to their intermediate positions because the degree of enhancement will not be adequate in the recorded sequence if the monitor had been adjusted to an extreme setting. To use VEC microscopy to its full extent make sure that the microscope objective and condenser front lenses are absolutely clean (check at least once daily) and that the lamp is always optimally adjusted and centred. * Particles need to appear in DIC microscopy images as if illuminated from above, i.e. with their bright part up, while vacuoles have the opposite shadows. If this is not the case, the camera has to be rotated 180° or the compensator or Wollaston prism has to be set to the opposite side with respect to the extinction position. Modified from ref. 30.

damage of the camera!). The field diaphragm must be centred on the monitor and opened until it becomes just invisible. If the field diaphragm is opened too much, most microscope-camera adapter tubes or high power projectives and oculars will create a very annoying central hot spot (Figure 6). If this problem persists after the adjustment for proper illumination, closing the projective diaphragm, or inserting a self-made diaphragm to cut the peripheral light at the microscope exit usually helps. Note that at higher magnifications and numerical apertures, Kohler illumination has to be readjusted once the focus is changed more than a few micrometres. As we will apply extreme contrast enhancement later, we have to start out 111

Dieter C. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen with as even an illumination setting as possible. Proper centring of the lamp and setting of the collector lens are, therefore, important. Also, as much light as possible needs to be collected at high magnifications. Some workers have used critical illumination that is focusing the light source onto the specimen plane instead of Kohler illumination (26). This is counter to good microscopical practice and can lead to very uneven illumination as the filament or arc will be superimposed onto the image of the specimen and subsequently, has to be subtracted digitally by mottle subtraction. Critical illumination might be useful, however, in those cases where the illuminating light has been made extremely homogeneous by light scrambling with a light fibre device (2,26,31). Generally, it is necessary in video microscopy to provide enough light for near-saturation of the video camera prior to applying analogue or digital enhancement. Manufacturers of suitable equipment have red and green control lamps built into their camera control units to indicate this (e.g. Hamamatsu Photonics). If there is insufficient light, the following measures are recommended: (a) Use brighter lamp types (mercury or xenon lamp). (b) Redo the illumination adjustments such as setting Kohler illumination and centring the filament or arc, possibly while observing the image on the monitor to improve. (c) Remove ground glass diffusers from the light path. (d) Make sure that the video exit port receives 100% of the light. (e) Reduce magnification slightly. (f) Replace the otherwise highly recommended narrow band pass filter by a wider one. (g) In AVEC microscopy near-saturation should be reached at a retardation setting of about one-ninth of a wave (20°), as this provides the best resolution (25). Further opening of the crossed polarizers far beyond 20° will rarely improve the image further, but it may introduce amounts of straylight no longer manageable by offset and it will lead to a bright-field type of image. In the case of excessive light reaching the camera the following steps are recommended: (a) Apply weak neutral density grey filters. (b) Employ high power light sources that can be attenuated, e.g. the AttoArc system (Zeiss) or metal halide burners (e.g. from Nikon). (c) Increase magnification slightly. (d) Reducing light intensity by closing the aperture diaphragm is not recommended as it reduces resolution. 112

3: Video microscopy (e) AVEC microscopy: if the one-ninth of a wavelength setting yields too much light for the camera, this could be reduced by setting compensator or Wollaston prism to a position of less than 20°. Although widely used, this does somewhat compromise image quality especially if settings of less than 15° or 10° are used (3, 4, 25). 2.3.3 Full numerical aperture for highest resolution: step 3 Open the condenser diaphragm fully in order to utilize the highest possible numerical aperture to obtain highest resolution. Also any iris diaphragm of the objective should be fully opened. Be careful to protect the camera from high light intensity prior to this step. The result of opening the condenser diaphragm will usually be that the optical image worsens because it becomes too bright and flat for the eye. This setting will result in a small depth of focus, especially with DIC (optical sections of 0.3 (um or less with x 100, NA 1.4 oil objectives). If the collection of specimen information from a larger depth of focus is desired and resolution can be sacrificed (e.g. when viewing dilute suspensions of organelles or bacteria), the condenser diaphragm may be closed down as desired. 2.3.4 Setting the compensator: step 4 (polarized light techniques only) Set the compensator or polarizer (AVEC-POL) or the main prism (AVECDIC) to about 1/9 y. The optical image, that is the one seen in the oculars, will disappear due to excessive straylight. If you have the accessories for de Senarmont compensation as recommended by Allen et al. (3, 4) (Figure 12) this is done by setting them at 20° off extinction. The basic set-up of de Senarmont compensation is done as follows: (a) (b) (c) (d)

Remove both Wollaston prisms and 1/4 wave plate from the light path. Set analyser and polarizer to the maximal extinction. Insert a 1/4 wave plate at 0° (maximal extinction). Insert Wollaston prisms and set the adjustable one to the best symmetrical extinction (if possible, check with a phase telescope for symmetry of the pattern in the back focal plane). (e) Use the rotatable analyser as compensator and set it as desired (1/9 of a wave is 20°). If you do not have such a calibrated system, first determine the distance between extinction (0°) and maximum brightness (90° or 1/2 X.) by moving the adjustable polarizer, then estimate and select the 1/9 of a wave or 20° position (Figure 13). If this is not possible, one can try to find empirically a suitable setting by shifting the movable Wollaston prism away from extinction. Many microscopes equipped with DIC for biological applications do not allow a 113

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen phase shift of 90° and some may not even allow 20° since for observation by eye phase shifts of a few degrees yield good contrast. Microscope manufacturers will, however, have the proper parts in their mineralogical programmes. At this point you have again to make sure that the camera is protected from excessive light but receives enough light to work near its saturation end. Ideally this should be near the 20° setting. Light adjustment should be done as explained for step 2. In IVEC microscopy straylight is not admitted, that is the polarizers stay close to extinction and the special rectifying optics further reduce the straylight. Filters to reduce brightness will not be required, but much brighter lamps will most probably be necessary to saturate the camera. 2.3.5 Analogue enhancement: step 5 First, increase the gain on the camera to obtain good contrast. Then apply offset (pedestal). Always stop before you lose parts of the image that become too dark or too bright. Repeat this procedure several times, if necessary and helpful. Make sure that the monitor for watching the changes is not set to extreme contrast or brightness, and is terminated properly (75 O) (see Section 5.1.6). Analogue enhancement improves the image contrast of the specimen but unfortunately also emphasizes dust particles, uneven illumination, and optical imperfections. These artefacts, called 'mottle', are superimposed on the image of the specimen and may in some cases totally obscure it (Figures 3 and 6). Disturbing contributions from fixed pattern noise (mottle) or excessive degrees of uneven illumination can be tolerated if digital enhancement is performed later (Figure 3). If digital processing is not possible stop enhancement just before the mottle or uneven illumination becomes annoying. Apply analogue shading correction and other types of analogue image improvement if your camera control unit offers these features (see Section 1.4.1 and Figure 8). Optimal adjustment of the lamp and thorough cleaning of the inner optical surfaces of the microscope, especially the surfaces in the projecting system to the camera (ocular, camera lens), usually results in images which allow the application of considerably higher analogue contrast enhancement. Finding dust. When the imaged dust particles or the mottle pattern rotate when the camera head is rotated, they are in the optical path before the camera, while immobile dust is to be found on the camera target. Rotate the ocular or camera objective lens to find dust located there. Dust should be removed with a low-pressure air gun or an optical cleaning brush. If this does not help use lens paper or a fat-free cotton tip applicator (wooden stick, not plastic) with ethanol or ether (in the fume-hood only!). Work from the centre to the periphery in a circular fashion while carefully avoiding to apply any pressure. Dust or mottle that is defocused together with the specimen is part of the specimen. 114

3: Video microscopy 2.3.6 Find background scene: step 6 Try removing the specimen laterally out of the field of view or (when using DIC) defocus to render it just invisible (preferably towards the coverglass). The result is an image containing only the imperfections of your microscope system (mottle pattern) (Figures 3c and 6c). This step will not be satisfactory, however, with such techniques as phase-contrast or bright-field. 2.3.7 Background (mottle) subtraction: step 7 Freeze, i.e. store the mottle image, preferably averaged over several frames (e.g. 64), and subtract it from all incoming video frames. 2.3.8 Return specimen to focal plane: step 8 When returning to the focal plane, you should see an absolutely even and clean image, which may, however, be weak in contrast. If there are 'missing' regions that are grey and flat, there is too much contrast in the mottle of the raw image to be subtractable properly. Reduce gain, adjust offset, and repeat the procedure (Figures 3d, e, and 6d). 2.3.9 Digital enhancement: step 9 Perform digital enhancement in a similar manner to step 5, that is alternate between stretching a selected range of grey levels (setting 'width') and shifting the image obtained up and down the scale of grey levels (setting 'level' or 'lower level') until a pleasing result is found. If available on your equipment, display the grey level histogram and select the upper and lower limits which are to be defined as bright white and saturated black, respectively. If the image is noisy (pixel noise) go to step 10 or 11. 2.3.10 Temporal filtering, frame averaging: step 10 Use an averaging function in a rolling (recursive filtering) or jumping mode over two or four frames. This will still allow the observation of movements in your specimen, but very fast motions and noise due to pixel fluctuations will be averaged out. Averaging over longer periods of time will filter out all undesired motion, e.g. distracting Brownian motion of small particles in suspension. The image will then contain the immobile parts of the objects, exclusively. Please note that not all image processors are capable of performing background subtraction and the rolling average function simultaneously. In this case, averaging generally yields the better image improvement for VIM, while background subtraction is more advantageous in VEC microscopy, although this should be determined experimentally. Alternatively, background subtracted or averaged scenes (plus empty scene for later background) can be stored on video tape or disk and then subsequently be played back into the processor and used for further processing. 115

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen

2.3.11 Spatial filtering: step 11 A number of procedures for spatial filtering is available, which can be used to reduce noise, to enhance edges of objects, or to reduce shading. These have been described for the analysis of single images in Chapter 8 but some image processors offer such filters working at video rate so that live sequences can be accentuated by filtering prior to recording.

2.4 Interpretation Unlike in EM images, which truly resolve the submicroscopic objects depicted (Figure 15a), the sizes of objects seen by AVEC-DIC microscopy may not necessarily reflect their real size. Objects smaller than the limit of resolution, that is 180-250 nm, depending on the optics and the wavelength of light used, are inflated by diffraction to the size of the resolution limit, the Airy disk diameter. The orientation of linear objects relative to the direction of DIC shear may also somewhat affect their apparent thickness and contrast, if they are oriented at angles very close to 45° or 135" (Figure I5b).Whereas the size of the image does not allow a decision on whether one or several objects of a size smaller than the limit of resolution are present, the degree of contrast sometimes permits this judgement to be made, A pair of adjacent micro-

Figure 15. Schematic representation of visualization of differently sized cellular components of a cytoplasm extract sample using various kinds of microscopy. (a) In transmission electron microscopy (TEM) all membraneous and cytoskeletal elements are visualized and represented in their true shapes and size relationships, i.e. they are resolved. TEM can, however, only be used with fixed (physically or chemically) material. (b) AVEC-DIC microscopy permits visualization of objects smaller than the limit of resolution of conventional light microscopy but larger than 20 nm. Many objects of 20-200 nm can be detected. These objects would, however, not appear at their real size, but inflated by diffraction to about the size of the resolution limit (~ 200 nm). Linear objects such as microtutaules may lose their shadowcast appearance at orientations close to 45" or 135° (c) In conventional differential interference contrast (DIC) microscopy objects appear also in their typical shadowcast manner. The smallest objects visible are of an apparent size in the order of the theoretical limit of resolution: M, mitochondrion larger than 500 nm; MT, microtubule of 25 nm in diameter; NF, neurofllaments of 10 nm in diameter; sV, axoplasmic or synaptic vesicles of about 50 nm in diameter; V, vesicles larger than 200 nm.

116

3: Video microscopy tubules would, for example, appear to have the same thickness as a single one, but the contrast would be about twice as high. If large numbers of subresolution objects are crowded together and separated by distances less than 200 nm from one another (e.g. vesicles in a synaptic nerve ending), they will remain invisible, because of the overlap of their Airy disk images. However, they will be clearly visualized, if they were separated by more than the resolution limit. Also remember that, if in-focus subtraction (Figure 7) or averaging over time has been used to create the image, all immobile or all moving parts of the specimen, respectively, will be completely missing in recorded video scenes.

2.5 Typical applications and limitations 2.5.1 Bright-field microscopy i. Advantages. Low intrinsic contrast due to low concentrations of natural chromophores or especially vital stains can be greatly enhanced by VEC microscopy. This can be of great advantage, since toxicity of a dye is often less of a problem at lower concentrations. Observation in monochromatic light at the wavelength of maximum dye absorption further increases contrast. By using appropriate cameras with quartz windows, even UV microscopy which yields high resolution and high contrast images (32) is feasible to some extent, because the reduction in illumination light intensity allows a considerably longer survival of the specimen and longer observation time. Colloidal gold, especially when coupled to antibodies, is an important tool in immunocytochemistry at the EM level (33). Colloidal gold in sizes down to 5-10 nm can also be detected using bright-field VEC microscopy (nanovid microscopy, see Chapter 12). Different sizes of colloidal gold will appear the same size (0.1-0.2 um depending on focus), but occupy different grey levels. By pseudocolour conversion of these grey levels, they can, however, be differentiated into size classes. Thus, double labelling with different sizes of colloidal gold is feasible (see also Section 2.5.7). ii. Problems with enhanced bright-field. Phase objects exhibit minimal contrast in-focus, and show opposite contrast above and below focus. It is, therefore, often very difficult to store the out-of-focus mottle image without contributions from the specimen. This limits the use of high analogue contrast enhancement in bright-field microscopy. 2.5.2 Dark-field microscopy Dark-field microscopy uses the light scattered from obliquely illuminated objects which then behave as self-luminous objects. These have been visualized, but not truly resolved, down to sizes of a few nanometres, if sufficiently powerful illumination was used (up to 1000 W mercury lamps) (see Figure 2) even without video contrast enhancement. i. Advantages. Dark-field images provide very high contrast, and this can be further enhanced by applying gain to the video signal. Genuine straylight 117

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen from nearby or out-of-focus scatterers can be subtracted by using offset to a limited extent, thereby increasing contrast and visibility of weak objects next to larger ones. It may be noted that dark-field images can also be subjected to video-intensified microscopy. ii. Disadvantages. The specimen should be thin and contain relatively few scatterers. The most annoying out-of-focus scatterers do not produce true straylight because the scattered light is not evenly distributed over the image. Because for transmitted light dark-field microscopy the objective numerical aperture must be limited, dark-field is not a very high resolution method. Nevertheless, it can detect the presence of structures in the nanometre range in thin preparations. In epi-illumination dark-field mode it is possible, however, to work at maximum numerical aperture and resolution. 2.5.3 Anaxial illumination In anaxial (or oblique) illumination methods the condenser aperture is unevenly illuminated to produce a differential (shadowcast) image of increased true resolution. In Abbe's original method, the condenser diaphragm is moved to one side of the front focal plane of the condenser. In Hoffman's modulation contrast method (34) (Modulation Optics, Inc.) undiffracted light is partially excluded by a trizonal plate in the condenser, which also serves to accentuate high spatial frequencies. The optional rotatable polarizer makes it possible to vary the eccentricity of the undiffracted light. The objective single sideband imaging technique (31) also yields very good results with VEC microscopy. It is also possible, using a tungsten filament lamp with a hemispherical mirror, to displace the mirror to illuminate only half of the aperture (28). For video-enhanced contrast microscopy, these high resolution methods work very well, because their former limitation of providing low contrast images can be overcome by contrast enhancement. i. Advantages. Phase details are observed in directionally shadowed differential images (similar to Nomarski-DIC). High specimen birefringence does not interfere as in DIC microscopy. For example, myelinated vertebrate axons can be better seen with enhanced anaxial illumination than with DIC, because their highly birefringent myelin sheaths create excessive contrast in DIC. The equipment is cheap, especially when only a bright-field microscope is required. It has been reported that observation in deeper layers of tissue is still satisfactory where the quality of DIC images would be much poorer (28). In principle, anaxial illumination does not reduce working aperture or resolution as long as more than half the entrance pupil is illuminated, but can increase true resolution, if higher order maxima of the specimen's diffraction pattern are captured. ii. Disadvantages. Anaxial illumination is sometimes less sensitive than Nomarski-DIC, and the resulting image may have some anisotropy (for test 118

3: Video microscopy images see ref. 36). The optical sectioning is also thicker and less precise than with DIC. 2.5.4 Phase-contrast Phase-contrast is an adequate method for viewing very thin, unstained specimens and very small phase objects, especially living cells. Video contrast enhancement is often of great value, especially in thin specimens and if analogue enhancement is sufficient. Enhancement is limited to cases where the bright halos around objects are negligible, that is with the use of the highest magnification objectives only. i. Disadvantage. Thick specimens imaged in phase-contrast cannot be optically sectioned, because each image plane contains opposite contrast information contributed from details in the levels above and below the focus, and because halos are seen around phase details. Enhancement rapidly raises a very disturbing mottle pattern. It can sometimes be subtracted from a plane approximately 0.25 um above or below the plane of interest, especially, if an area free of specimen is used. In most cases out-of-focus mottle images of the specimen for background subtraction cannot be obtained due to the strong contributions from the out-of-focus levels of the specimen. Even extreme cleaning of the projecting optics is usually not sufficient to remove all the annoying mottle pattern. 2.5.5 Polarization microscopy For visual observations of most biological objects that are only weakly birefringent (spindles, organelles, cytoplasm), the microscope equipped with simple plastic polarizers is rather inadequate. The image is flooded with straylight owing to a combination of depolarization at lens surfaces, lower extinction of the polarizers, and strain birefringence in one or more lens elements (Table 1). For sensitive visualization or photomicrography of weakly birefringement structures, it was previously only possible to use a polarizing microscope with a rectifier (Nikon) that eliminated not only straylight due to depolarization at lens surfaces, but also the disturbing diffraction anomaly that can lead to spurious contrast and resolution (4, 37). The rectifier consists of a zero power meniscus lens and a properly orientated half-wave plate (2, 24). Using video-enhanced contrast polarization microscopy (VEC-POL), however, relatively sensitive observations can be made with a simple polarizing microscope even if the lenses are slightly strained. One can, in fact, in most instances convert a bright-field microscope into a video equivalent of a relatively high extinction microscope by adding inexpensive film polarizers (e.g. Polaroid HN 22 polarizing material which is available in sheets and can be cut down to any desired size; Polaroid, Inc.). In an analogy to what has been said concerning AVEC-DIC (see Section 2.1 and Figure 13), at 1/9 X. bias retardation (AVEC-POL), the anomalous clover leaf Airy disk diffraction pattern 119

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen normally seen at extinction (zero retardation and crossed polars) is converted into a normal Airy disk, resulting in the absence of spurious resolution or contrast. In addition, the diffraction anomaly due to weak strain birefringence in any of the lens components also disappears (3, 4). The use of the highest quality optical components for polarized light microscopy together with video enhancement (AVEC-POL and IVEC-POL, see Section 2.1) allows visualization of extremely weak birefringent objects, such as individual microtubules, bacterial flagella, etc. (2, 36). 2.5.6 Differential interference contrast microscopy In DIC microscopy the illuminating beam of plane-polarized light is split by the first Wollaston prism into two parallel beams of polarized light passing through the specimen oriented at right angles to each other and very close together. The separation (or shear) needs to be smaller than the limit of resolution to prevent the appearance of double images but large enough to create a satisfying shadowing effect. Sets of Wollaston prisms with very high shear, e.g. a X 100 objective and a 200-250 nm shear, create superb contrast for visual observation (such as some older lines from Zeiss Jena, e.g. JENAVAL), but at the increased true resolution of VEC microscopy one sees very annoying double images. Generally, all medium shear pairs of Wollaston prisms are suitable for VEC-DIC and AVEC-DIC. However, Nikon offers a special 'short shear' pair of Wollaston prisms with their Eclipse series, that are especially suitable for VEC microscopy. i. Advantage. Differential interference contrast is the optical technique, which is, in most applications, best suited for video contrast enhancement. The Nomarski-DIC method gives in-focus, high contrast, shadowcast images of phase details in which the detection of shadowing is opposite for phase advancing and retarding details (37). The generation of contrast at high working aperture is limited to a very thin depth of field, with the result that this technique is unique in its ability to render high contrast optical sections of only 0.3 um in thickness; i.e. thinner than those obtainable by confocal microscopy (Chapter 2). Amplitude contrast can also be obtained (37). With the AVECor IVEC-DIC method the sensitivity of detection is increased to the level that transparent phase objects as small as 15-20 nm can be detected under optimal conditions. Increasing the bias retardation (Figure 13) causes a disproportionate increase in the contrast due to small phase retardations in comparison to large retardations. For example, if low contrast details of cytoplasmic organelles are obscured by bright structures, such as, for example starch grains, the contrast due to the latter can be diminished by using a dimmer light source and increasing the bias retardation more towards 1/4 A. (Figure 13). ii. Disadvantage. Perhaps the only disadvantage of this method is that contrast due to refraction in some highly birefringent objects (e.g. striated muscle or 120

3: Video microscopy myelinated axons) can be masked by the contrast due to birefringence. In this case anaxial illumination is recommended as a substitute. Because in DIC contrast is directional, the specimen should be mounted on a rotatable stage and examined at more than one orientation to avoid missing weak linear features aligned parallel to the direction of shear (see Figure 15). Note that DIC microscopy uses polarized light and any birefringent material other than the object will show up bright at certain orientations. Strain birefringence in the optics, in lens cement or any plastic material, makes DIC microscopy impossible. Therefore, optic elements certified 'DIC' or 'POL' are required. Working on plastic slides or plastic Petri dishes with their unavoidable birefringence is impossible. If working with these is unavoidable, Hoffman contrast or anaxial illumination contrast need to be used. 2.5.7 Reflection interference contrast microscopy Surface reflection interference from epi-illuminated specimens is used to image zones of cellular attachment to glass (38). To reduce the inevitable straylight caused by reflections from lens elements, the use of antiflex lenses designed for metallurgical microscopes containing a 1/4 wave plate on the front surface of the objective was introduced. Using video enhancement, simple epireflection microscopy with a 50% mirror cube but without polarizing elements or special antiflex lenses can be performed for several purposes. (a) To enhance the observation of focal contacts of cell attachment to glass. (Surface reflection interference at high illumination aperture.) (b) To enhance the observation of contour-mapping interference fringes to document the shapes and heights of attached cells or the topology of surfaces of cells above a flat glass surface. (Surface reflection interference at low illuminating aperture.) (c) To render visible colloidal gold or other metals down to a diameter of 5 run as bright objects. Epipolarization even further improves the visibility (39, 40). In these applications the use of antiflex objectives with 1/4 X plate (Carl Zeiss GmbH) as in the original technique (38) will further improve image quality. For further applications see Chapter 7. 2.5.8 Fluorescence microscopy With fluorescence optics images are produced that are characterized by relatively high contrast and very low intensity. Therefore, fluorescence images usually need intensification, especially if the level of illumination has to be kept low to delay bleaching. Any straylight arising from residual autofluorescence in lenses or the mounting medium can be removed by offset. However, light from out-of-focus fluorescent structures usually cannot be treated as straylight because of its non-random distribution. Often fluorescent details 121

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen can be seen better when their contrast has been reversed by inverting the television signal (negative image). Fluorescent specimens have usually to be viewed with VIM but the use of most types of low light cameras will inevitably lead to reduced resolution when compared to images obtained by VEC microscopy at the same magnification. With some applications, especially in the case of strong fluorescence it may pay to try the use of a high resolution, high sensitivity camera (Newvicon or Ultricon) in the rolling average mode (over one or more seconds). Similarly accumulation of images for a few seconds may bring about a useful image. In fluorescence microscopy self-luminous objects are depicted, images contain, therefore, a strong contribution from out-of-focus objects. Images can, however, be limited to a depth of focus of about 1 um by digital blur-deconvolution (41, 42), using suitable software packages or to a depth of focus of about 0.6 nm by the use of confocal microscopes (see Chapter 2). Suitable software packages are provided by, for example, Bitplane AG, Carl Zeiss GmbH, ImprovisionTechnology, Inc., Inovision Corp., Scanalytics, Inc., VayTec, Inc., or Vital Images. 2.5.9 Examples of biological and biochemical applications Specimens which are extremely weak in contrast or even invisible by conventional microscopy are best suited for AVEC-DIC microscopy. Examples in this class are micelles, liposomes and single or double layer membraneous material, colloids (43), live, actively transcribing rDNA genes (44), synaptic and other small cytoplasmic vesicles (8, 9), artificial latex particles of 50 nm and smaller, and cytoskeletal elements such as microtubules (9, 36) and actin bundles (21, 45). The process of microtubule gliding was discovered by AVEC-DIC microscopy (8), its ATP-dependence and the motor enzymes were discovered (46) using video-microscopic motility assays (8), and even molecular events such as microtubule subunit assembly and disassembly can be measured (47). Stained material or other objects, which already have high contrast are less well suited. An overview of what can be seen and analysed by this method in the living cell has been published elsewhere (30). The AVEC-POL technique can visualize very weakly birefringent objects such as individual microtubules (36). When applied to bright-field or epipolarization microscopy, the VEC technique visualizes 5-20 nm diameter colloidal gold particles (39, 40) and it can be used to screen gold labelled EM specimens quickly in the light microscope. The same is true for semi-thin and thin, unstained, plastic-embedded EM sections (Figure 3). Techniques to improve images of moving objects can be generated with the aid of digital processors. The 'trace' operation adds frames at predetermined intervals to the frame memory thereby generating images showing multiple positions of moving objects. Jumping averaging over several seconds visualizes processes too slow to be detected otherwise, such as cell growth, chromosome movements, and cell locomotion. Rolling averaging can be used as a 122

3: Video microscopy filter to remove velocities greater than a certain pre-selected velocity from the images. Conversely, subtraction of sequential in-focus images and scenes can be used to view moving objects only, while stationary ones are absent from the image (Figure 7). The interval subtraction mode (see Section 1.3.4.v) can be used to de-emphasize especially the slowly moving objects. If the interval before the next background image is grabbed is made longer, slower movements, which are otherwise excluded, will become part of the image.

3. Low light: video-intensified microscopy 3.1 Introduction This technique requires the especially sensitive cameras such as cooled CCD cameras or the image intensifier units mentioned in Section 1.4.2. Once the image has been analogue enhanced and is picked up then the digital techniques are used as in VEC microscopy (see Section 2.3). First and most importantly, they improve the S/N during image acquisition by, for example integration, averaging, or digital filtering. Secondly, they remove any 'fixed pattern noise' such as mottle, uneven illumination, background fluorescence, or any pattern emanating from the camera or the digitizing process (background/mottle subtraction). Thirdly, they can be used for digital contrast manipulations, while enhancement or suppression of motion is the fourth feature of VIM. The major difference relative to VEC microscopy in the application of these functions to VIM is that generally only static or slowly moving specimens can be examined. These require the processing described in Sections 3.2.2 and 3.2.4, while a dynamic specimen (see Section 3.2.3) can be treated similarly to those viewed by VEC microscopy.

3.2 Procedure for image generation 3.2.1 Microscope considerations There are no special or unconventional adjustments of the microscope necessary to perform VIM. One should, of course, adhere to good microscopic practices such as cleanliness of the optics, careful alignment of the illumination system, and of the camera to the light path. There are, however, a number of practical issues which will help ensure the best results when performing VIM. (a) If using a camera with extended red-sensitivity (e.g. multialkali photocathode) for viewing in visible wavelengths it is suggested to use at least one high quality IR cut-off filter in the camera's light path. While the human eye and photographic film are not sensitive to these longer wavelengths, they can seriously degrade image quality. This is true even if good fluorescence filters are in place since these are notorious for passing through light over 700 nm. (b) The camera image should be as parfocal with the eyepieces as possible. 123

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(c)

(d)

(e)

(f)

(g) (h)

When working with very low intensity images it can be very difficult to accurately assess exact focus from the live image on the monitor. Provisions should be made for working in a dark-room or establishing some method to shield the objective lenses from all extraneous light. Highly intensified cameras easily pick up this light and image quality will consequently suffer. It is necessary to cover the eyepiece of the microscope with, for example a black cloth or slide the shutter provided by some manufacturers to the in-position. The illuminator should be equipped with a regulated power supply if low intensity illumination will be utilized (e.g. to reduce photobleaching in fluorescence) or if quantitative measurements are planned. Small fluctuations in line current are greatly magnified by intensified cameras so that stabilized power supplies are recommended. It is advisable to equip the microscope in such a way that a good optical image (bright-field or DIC) can be obtained for positioning and focusing the sample prior to directing the low intensity light to the camera. For applications in fluorescence it is important to work with non-excitatory light before moving into the fluorescence mode to minimize photobleaching effects. Since integration or averaging may be required over relatively long periods of time, the microscope should be as stable as possible to prevent any image blurring. Additionally, because many low light cameras are relatively large, it is often good practice to stabilize the camera by some means in addition to the C-mount. Care must be taken to obtain slides, coverglass, immersion liquids, etc. that are free of, or extremely low in, autofluorescence. The microscope should be equipped with a series of neutral density filters. These are used to reduce illumination of the specimen as well as to protect the camera from excessive light.

3.2.2 Acquisition of static images (video cameras) (a) Focus the specimen through the eyepiece ensuring that no light is directed to the camera. (b) With the camera sensitivity reduced to its minimum setting, direct the light to the camera. If light to the camera is excessive immediately direct the light away from the camera or block the illumination path. It will be necessary to insert neutral density filters into the light path before trying again. (c) With light to the camera and an image on the monitor, focus the specimen, if necessary, and increase camera sensitivity until some portion of the image begins to saturate. Now reduce sensitivity to just eliminate any saturation. This procedure ensures that the video signal is of full amplitude 124

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(d) (e) (f)

(g)

(h)

(1V peak-to-peak) and will provide the best S/N. An alternative approach to this is to increase the light level rather than the sensitivity. Obviously, this is not possible in those cases where light is limiting or where light levels are intentionally kept low in order to reduce photobleaching, phototoxicity, phototropic effects, etc. Adjust the monitor for best picture quality. If the image is to be recorded, intermediate settings for contrast and brightness are strongly recommended. If the camera is equipped with gain and offset, adjust these parameters for the best overall image quality or until that part of the specimen which is of interest displays the most information (see Section 2.3.5). Integrate the image for a suitable number of frames. Depending on the quality of the 'raw' or analogue image this can vary anywhere from 2 to 512 frames. The value chosen should be based on the user's qualitative judgement or some established criteria such as S/N ratio. As S/N is proportional to the square root of the number of frames, a point of diminishing returns is reached between 128-256 frames. It should be pointed out that the maximum number of frames integrated may be restricted by the 'bit depth' of the digital image memory. Systems for low light imaging should be equipped with memories at least 12-bits and preferably 16-bits deep to allow for extended integration. Establish and maintain a background image. Two possibilities exist for this background image. The best is to find a suitable background area adjacent to the specimen. This area will contain all the components of the image which should ideally be removed. This includes any shading contributions or fixed pattern noise of the camera, illumination irregularities, optical defects, autofluorescence in the system, and any digital noise. If such an area is not available, the second alternative is to block light to the camera and use this 'dark image' as the background. This will correct for any camera-related phenomena. In almost no case should an out-offocus image of the specimen be used as the background image. With few exceptions (e.g. DIC) defocusing cannot eliminate all specimen-based image information and one runs the risk of erroneously subtracting non-background information. Now, with all settings identical (camera sensitivity, illumination, etc.) integrate a background image into a different digital memory than that of the specimen and subtract it from the stored specimen image analogously to Section 2.3.7. Continue with Sections 2.3.9 to 2.3.11 to manipulate contrast for the most pleasing or informational image.

3.2.3 Acquisition of dynamic images (video cameras) (a-e) Proceed as in Section 3.2.2. (f) Acquire a background image as in Section 3.2.2(g). Ideally, this background 125

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen image should be integrated or averaged for the same period as the rolling average utilized in the next step. This image should be placed in a memory which can be subtracted from each incoming video frame as in VEC microscopy. (g) Return to the specimen image and begin video rate background subtraction as in Section 2.3.7. If background is high, this subtraction process may result in the image losing intensity. If so, add digital offset to the image to restore intensity or brightness. (h) Apply a 'rolling' or 'exponentially weighted moving' average to the background subtracted image as in Section 2.3.10. This type of average offers similar S/N improvement [S/N = ±(2n - 1); n = number of frames] as integration, which is used for static scenes (see Section 3.2.2, step f), but offers the advantage of being applicable to dynamic specimens. Additionally, unlike image integration, in averaging the image is normalized, therefore the digital memory does not overflow or saturate. (i) Digitally improve image contrast as in Sections 2.3.9 and 2.3.11. 3.2.4 The use of cooled, slow-scan CCD cameras to record images SIT and cooled CCD cameras allow the recording of signals at very low light level. With a SIT camera low light signals can be recorded at video rate thereby allowing the detection of dynamic changes in real time. However, high sensitivity and high spatial resolution tend to be mutually exclusive features of these cameras due to signal noise. In contrast, cooled CCD cameras provide high sensitivity while maintaining a high spatial resolution. To ensure this, images must be recorded at a slower rate. In the following paragraphs we will discuss some considerations on the use of slow-scanning, cooled CCD cameras. i. Installing a cooled CCD camera on a microscope The CCD camera can be linked to the microscope using the C-mount connector at the camera tube. The cooled CCD camera should be turned on always after the power supply for the arc lamp (e.g. high pressure mercury lamp) had been ignited to avoid damage to the CCD chip by a possible power surge. As part of the set-up protocol one should assure that the focal plane of the camera is properly lined up with that of the eyepieces. The software provided with the camera will usually have a submenu to focus the specimen onto the camera chip. The focus mode will display only a small portion of the image at a higher refresh rate. After the focus plane of the camera has been adjusted the mounting of the camera should be secured tightly. While the cooled CCD cameras can be operated at very short exposure times (< 1 sec), prolonged focusing on a fluorescent sample will still cause photo damage to the sample. It is, therefore, recommended to use the eyepieces for faster selecting the appropriate focal plane rather than the monitor. 126

3: Video microscopy ii. Taking images Most high resolution cooled CCD cameras will record images with a dynamic range of 12-bit (4096 grey levels) instead of 8-bit (256 grey levels). The high dynamic range of these cameras allows, therefore, to record fluorescence images. When collecting images with a high dynamic range of grey levels one should, however, be aware that most image processing software packages such as PhotoShop (Adobe Systems, Inc.) or Paint Shop Pro (Jasc Software, Inc.) can only handle 8-bit grey level and 24-bit colour images. From advances in computer technology more sophisticated software can be expected that will be able to handle 12-bit grey and 36-bit colour images. Keeping in mind the limitation of the current image processing software, particular care should be taken when choosing the proper exposure time. It is recommended to take several images with different exposure times. Hi. Obtaining colour images Multicolour fluorescence images can be generated from specimens stained with multiple fluorescent dyes. In this case the correct exposure of the individual images is particularly important. First, single images for each fluorescent signal are generated using the cooled CCD camera. It should be noted here that the original images should always be saved uncompressed and in a file format (e.g. .TIF) which is universally accessible by the various types of image processing software. The individual images are saved as 8-bit grey images in TIF file format. Colour images are then composed in, for example, PhotoShop (Adobe Systems, Inc.) or similar programs using the 24-bit RGB option of the software. Before generating a colour image the intensity range and the level of all images to be merged should be adjusted, so that the whole range of intensities available for that colour is used (24-bit for all three colours) (Figure 16). The colour image can then be generated by opening a new image file of the appropriate size (matching that of the images to be merged) in the RGB mode and copying the single B/W images in either one of the red, green, or blue image layers.

3.3 Typical applications The detailed information on spectral properties and the potential application of most dyes for the studies mentioned below are compiled in ref. 48. The dyes can be obtained from companies such as Eastman-Kodak Company, Sigma Chemicals, and especially from Molecular Probes, Inc. which specializes in fluorescent compounds for microscopy. 3.3.1 Fluorescent analogue cytochemistry In fluorescent analogue cytochemistry (FAC), the molecule or organelle of interest is isolated, purified, fluorescently labelled, and reintroduced into the living cell (Chapter 10). Ideally, images of these 'analogues' can reveal the 127

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Figure 16. Combined VIM and VEC microscopy of the periphery of a neuronal precursor cell (hNT PF) in culture, (a) DIG image showing organelles and cytoplasmic extensions of the cell (Newvicon camera C24-00-07, Hamamatsu Photonics). Microtubules (b) and actin filaments (c) were detected by immunofluorescence microscopy using a cooled slowscan CCD camera (SenSys, Photometries Ltd.). A Nikon Diaphot 300 inverted microscope equipped with oil immersion condenser (NA 1.4), x 100 PlanApo DIC oil objective (NA 1.4), a Xenon lamp (XBO 100), and a Mercury lamp (HBO 100) were employed for acquir ing the fluorescence and DIC images, respectively. (d) Colour merge of (b) and (c) performed in PhotoShop (Adobe Systems, Inc.) in RGB mode.

distribution and organization of the antilogous native molecule or organelle and how they changc over lime. For example, the rate and polarity of the incorporation of actin and tubulin analogues i n t o in vivo structures has been examined using FAC as well as the changes in the cyloplasmic distribution of a variety of molecules (50). Fluorescence labels can also be introduced into 128

3: Video microscopy cells as caged compounds which can be activated locally by focusing the beam of long wavelength UV light in a particular area (49). 3.3.2 Ratio imaging Ratio imaging is a powerful technique in analysing the spatial and temporal dynamics of ions, molecules, and organelles in living cells (see Chapter 6). By generating the ratio or quotient of two images, this technique normalizes the cell for pathlength and accessible volume, thereby allowing for quantitative evaluations of subcellular concentrations. 3.3.3 Fluorescence recovery after photobleaching (FRAP) Studies of translational diffusion and lateral motion in membranes have benefited greatly from the application of video-intensified imaging. With this method, an area of cytoplasm or membrane containing a fluorescent probe is bleached using a laser or other strong illumination source. The rate and pattern of the reappearance of fluorescence in the bleached region can be used to analyse the contribution of isotropic diffusion, anisotropic diffusion, and bulk flow to lateral transport phenomena in living cells (51, 52). 3.3.4 Molecular imaging A number of macromolecules, including DNA and actin, have been labelled with fluorescent groups and studied using VIM (45, 53). The visualization of single DNA molecules in solution has allowed the study of changes in molecular conformation. Chromatin structure in isolated nuclei and intact cells has also been studied. Similarly, the elastic properties of single actin filaments and their movement on immobilized myosin have been studied (45, 54). 3.3.5 Video microspectrofluorometry The use of intensified cameras as an alternative to photomultiplier tubes offers the advantages of whole image, spatially resolved spectral analysis (55). Since many of the fluorochromes utilized to monitor physiological changes in living cells exhibit measurable changes in quantum yield, spectral shape, and spectral moments, this technique shows great promise for in vivo analyses. 3.3.6 Luminescence The recent commercial availability of cameras with single photon sensitivity has made it possible to image the extremely low intensity emission associated with several luminescent systems. Calcium transients during fertilization have been visualized using aequorin luminescence (56). Another, particularly exciting, application of VIM in this area has involved the direct visualization of gene expression in living cells using green fluorescent proteins or the lux operon as reporter genes. The lux gene, which codes for the enzyme luciferase, can be incorporated into cells in such a way that its transcription is controlled by the promoter of the gene under investigation; hence, activation of the promoter results in the simultaneous emission of light (10, 57). 129

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen 3.3.7 Neurobiology In neurobiology, a number of VIM applications have opened unexpected experimental approaches. By selecting especially non-toxic and non-metabolizable dyes mammalian nerve cells and the stability of their connections can be monitored over time in living animals. Appropriate dyes stain, for example neuromuscular end plates for several months, so that the innervation of superficial muscle can be reinspected during small surgery (58). Voltagesensitive fluorescent dyes can be used at low magnification to monitor the electrical activity of the vertebrate brain with much better spatial resolution than was possible with EEG analyses (59). At high magnification one yields information on membrane potentials in tissue sections and single cells (60, 61). Brain slices are a standard preparation for neurophysiological experiments. The difficulty to visualize individual neurones in standard thick slices has been a major drawback. This problem has, however, been overcome by the use of infrared VEC-DIC microscopy. Neurones in slices can now be visualized in great details, and furthermore, neuronal processes can be patch-clamped under direct visual control. A further development of infrared video microscopy enables one to visualize the spread of excitation in slices making video microscopy a tool for the direct investigation of neuronal function (29).

4. Image analysis: video-based techniques for measurements in living cells Extracting quantitative data out of microscope images or image sequences was a relatively tedious process before the advent of video technology. Digitization of an image and processing its numerical representation in a digital image processor has made a great number of quantitative parameters accessible with relative ease. Video microscopy has an additional advantage that images are already in video format, either live, on tape, or optical disk and thus can be analysed by analogue devices (see Section 1.4.6) for determining, for example, the intensity distribution along a given line (Figure 6) or distance information. Such analogue devices are, however, far less versatile than digital image analysers. Most digital processors for video microscopy are considered real time image pre-processors, that is their output is an analogue video signal destined to a monitor or recorder. If one wants to get access to the enormous variety of techniques for digital image analysis, usually special equipment or special PCbased software packages will be required where the analogue video signal is directly used as input signal. Most of the processors capable of real time video microscopy (see Section 1.4.5) are also able to perform most of the functions required for image analysis with their own processor. As a first level of analysis, a single frame can be taken up and analysed. 130

3: Video microscopy However, since video microscopy has newly opened the field of vital microscopy we want to be able to analyse changes in sequences of video images. Depending on the complexity of the algorithms required to extract the desired features it may or may not be possible to achieve this at video rate, that is 25 or 30 times a second. In the latter case the life images from the video microscope or from tape are sampled such that the image analyser grabs only every second, fourth, or n-th image, extracts the desired detail, and stores the information for later display as a function or plot. An introduction to digital image processing is found in refs 14 and 62.

4.1 Spatial measurements and motion analysis It is easy to extract such parameters as size, length, width, area, perimeter, or the coordinates of the centre of gravity for one or more objects which can be differentiated according to their specific grey shade. This is achieved by setting an upper and lower threshold (binarization of the image) and can be done automatically and in real time by most systems. In addition to conventional measurements, this makes accessible the analysis of motion, such as growth of objects (nuclei, cells, microtubules) or movement of individual organelles in intact cells or along free microtubules (8, 9, 39, 40, 63-65). Motion analysis of moving organelles or of microtubule ends for analysis of subunit assembly/disassembly is performed by extracting the X and y coordinates as follows (65, 66). (a) Store scenes of AVEC-DIC microscopy on video tape: storage of sequences of images in a computer memory would be possible for a limited duration (one second of 512 X 512 X 8-bit B/W images takes up 6 Mbyte of memory). (b) Play back the sequences through an X,Y-tracker or a suitable software program which detect, by thresholding, a bright object in a small userdefined and -positioned frame (region of interest). (c) Coordinates are stored at video rate or more slowly, as desired. (d) Further analyse the time series of positional data in the personal computer using specific motion analysis software (65) or general time series analysis software. (e) Plot the X coordinate against the y coordinate to obtain the trace of the moving object. (f) Plot position against time to get the velocity function. Similarly acceleration behaviour and directional changes can be displayed as functions of location or time. (g) Analyse the movement for intrinsic regular features by the standard techniques of time series analysis, such as autocorrelation and fast Fourier transform for oscillations, or cross-correlation for similarity of the motion of different organelles. 131

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen (h) If the object is not easily distinguishable by contrast, but visible by eye, perform an interactive frame-by-frame analysis. To this end, use a tape recorder with a 'single frame advance' feature. From each or each n-th frame extract the coordinates manually by moving a cursor with the particle or the feature of interest (65). Simultaneous analysis of a multitude of moving objects such as organelles, swimming micro-organisms, or sperm cells may be achieved by sophisticated dedicated equipment for motion analysis (e.g. from DataCube, Inc., HaSoTec GmbH, Inovision Corp., Mitec GmbH, Motion Analysis, Inc., or Universal Imaging Corp.).

4.2 Intensity measurements The intensity of each digital picture element is represented by a number, in images digitized to 8-bit accuracy, between 0 and 255. Intensity in a microscope image means absorption, phase retardation, fluorescence intensity, or birefringence, depending on the technique used. If we ensure that only one of these contributes to the image, we can quantify these properties in the different objects comprising the image. This is easily done by using specific fluorochromes in fluorescence and by applying monochromatic light in bright-field (absorption image), while the other parameters are more difficult to isolate. It should be made clear that photometers for work with cuvettes or built into microscopes use photomultipliers to measure integrated light intensities in a given slit or diaphragm with high accuracy, typically at a resolution of several thousand intensity levels, but without containing spatial information. Measuring intensities in a video image, on the other hand, although giving usually only a resolution of 256 grey levels, has the advantage of providing this information spatially resolved over 512 X 512 pixels. Intensity in the final video image is the result of many steps and cannot necessarily be assumed to still be linear or to follow Lambert-Beer's law. It is, therefore, essential, if absolute quantification is intended, to calibrate the system with known standard samples using the same optical and electronic settings (55, 67). This difficulty and the relatively low photometric resolution are the two grave drawbacks of video techniques. Cooled CCD array cameras have some advantages for quantification (68). The distribution of absorbing and fluorescent compounds in biological samples are accessible both in the spatial and temporal domain. Hardware and software devices to read out the intensity of a given pixel or along a line (Figure 6) or in a region of interest are available. If video sequences are analysed in the spatial domain such measurements provide information on diffusion and transport of endogenous molecules or of compounds which have either been taken up by cells, or have been microinjected. If light is shone on the specimen, or an area of it, this can be used to measure photobleaching or FRAP of fluorescent compounds (51, 52). 132

3: Video microscopy While most measurements give only estimates of amounts of the compounds in the light path, it is possible in some cases to measure concentrations. This is if images are divided by each other, such as in ratio imaging which is used to determine the Ca2+ or the H+ concentration (pH) (69-71) in living cells (see Chapter 6). These techniques, which aim towards a 'biochemistry with the microscope', that is assaying amounts and concentrations of specific molecules and ultimately also enzymes, are presently in the process of rapid development. Hundreds of fluorescent markers are available for specific organelles, cells, enzymes, and membranes, which can be used in living cells to detect and analyse specific features (48) (available from, for example, Molecular Probes, Inc.). The discussion of the special applications for different fields of biology are beyond the scope of this chapter so that the reader is referred to the publications mentioned in Section 3.3 and in ref. 30.

5. Documentation and presentation of video microscopy data Only the expert use of analogue and digital video technology allows profitable working of the video microscopy laboratory. It is the aim of this section to provide the beginner with the necessary basic knowledge of this technology. As the images generated or improved by video microscopy cannot be seen in or photographed from the microscope directly, we have to learn how to properly record and archive tapes and images and then how to obtain copies of tape, hard copies or photographs of single images, or how to edit and present video sequences (video clips) to larger audiences. Traditionally documentation and presentation of video microscopy data had been carried out by analogue methods, either by photography or making movie films, by analogue video recording, analogue editing, and playback on monitors. With advancements in digital technologies we are presently witnessing the transition to digital techniques. Therefore, it is now possible to record, print, archive, edit, and present video sequences in addition fully digitally. While recording and archiving is still dominated by analogue techniques, editing and presenting video sequences is done today by both technologies, but all single image techniques are performed almost exclusively by digital technologies.

5.1 Video recording Video recorders are indispensable for the storage of video-microscopic sequences. They provide a comfortable way of storing the enormous amount of information, because of their easy handling and because they offer the possibility of recording one or two (up to six) hours on one tape. The recordings can be played back and examined immediately without processing. Before 133

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen purchasing video equipment one should know about the standards and formats available and weigh up the advantages and disadvantages of particular systems. Furthermore, one should consider new storage devices such as the DVD (digital video device), which might very well replace the standard video tape recorders in the near future. 5.1.1 Video standards There are several colour television standards in the world. Many of the European countries, Australia, and many African countries use the German PAL (phase alternating line) system. France, some African, and most Eastern European countries use the French SECAM system (SECAM = sequentielle couleur a memoire). The NTSC standard (NTSC = National Television Systems Committee), developed in the USA, is the video standard in North America, many countries of Middle and South America, and Japan. Complete lists of the TV standards of all nations are available in video shops or in ref. 2. The American standard differs from the European ones mainly in the scan rate. The NTSC standard displays 30 images per second; each is scanned by 525 horizontal lines. Because each frame is dissected into two interlaced fields ('half pictures'), one containing the odd numbered lines, one the even numbered lines, the frequency is 30 frames/sec (f.p.s.) or 60 fields/sec. Therefore, the NTSC scan rate is defined as 525/60. The two European standards have a scan rate of 625/50, that is 25 f.p.s. scanned by 625 lines. It is important that every part of the video equipment is compatible. This means that all parts of the sets must have the same TV standard. The different scanning rates of PAL and NTSC imply that it is usually impossible to copy a PAL recording onto NTSC equipment or vice versa. While multi-standard video players are common, the actual transformation, that is accepting for example PAL format and recording it in NTSC format, can be done only with a few types of special all standard video recorders. The terms PAL, SECAM, and NTSC characterize especially the colour modes of the standards. However, they are fully compatible with the corresponding black and white (B/W) standards, namely PAL and SECAM with CCIR (50 Hz) and NTSC with EIA (60 Hz). The acquisition of colour equipment (monitors and recorders) is recommended, because this allows one to record B/W sequences and false colour (pseudocolour) images (see Section 5.1.7) which are often very useful in video microscopy. 5.1.2 Obtaining a correct video signal The peak-to-peak voltage of the composite video signal is standardized to a maximum of 1 V (the image information occupies the range from 0.3-1 V and the synchronization signal (SYNC) from 0-0.3 V). In the pulse-cross mode the four corners of the picture and the blanked region of the frame (black) which normally is hidden can be displayed on a monitor (see Figure 17). If the signal is deteriorated after playing back a recorded video sequence, we are able to 134

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Figure 17. Demonstrating the video SYNC signal with the pulse-cross mode of a monitor, (a) Correct pulse-cross display derived from a TV camera. Due to their low voltage the SYNC pulses appear as dark stripes. Vertical on the monitor, H SYNC pulses; horizontal, V SYNC pulses, (b) Pulse-cross display of a video signal from a video tape recorder. The vertical SYNC pulse shows a distortion, (c) (d) Photograph of a time lapse scene and the corresponding pulse-cross display. The 'flagging' in the upper part of the picture is caused by a jammed SYNC signal.

control and try to improve the quality of the video signal using this mode. Figure I7a and b show a correct video signal from a TV camera and one from a video tape recorder with a discordant time base for the vertical SYNC signal. Often one can improve the signal slightly by turning the TRACKING control of the video tape recorder or sliding the SKEW lever. The SKEW control adjusts tension of the tape while the TRACKING control minimizes the tracking variances between different recorders. Altering the TRACKING control may reduce even the strong distortion (flagging) often seen in time lapse scenes in the upper part of the screen (Figure 17c and d). This may also improve the image if the video level of the original tape is inadvertently more than 1 V. Normally (especially during recording) the TRACKING control has to be set to the 'fixed' position (automatically done in most recorders). If these measures are not resulting in a satisfying image signal the generally advisable steps to improve the working of delicate electronic equipment

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Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen should be taken, such as proper grounding, connecting all parts of the set to the same power circuit breaker, etc. 5.1.3 Video tape formats For use in scientific laboratories two different video tape formats are practical. These are 1/2 inch (1/2") and 2/3 inch (2/3") tapes. i. The 1/2 inch format The 1/2" format is the well-known home video standard. Cassettes of the VHS format are not compatible with devices for Beta format, which is likewise obtainable. The VHS format has become the 1/2" standard in the world. The advantage of the 1/2" format is the low price for recorders and tapes. The tape, however, is thinner and narrower than 3/4" tapes, resulting in more dropouts (small white flashing spots) and noisy jams after repeated recordings than occur with 3/4" or Super-VHS tapes. Most of the high quality 1/2" video tape recorders offer the possibility of assembly editing. This means that one can add scenes consecutively in a very easy way without interference between the scenes. Some high grade recorders even offer insert editing. A considerably unproved VHS system called Super-VHS (S-VHS) has been established as a new standard. The S-VHS uses the standard VHS cassette size but offers much better resolution. Because the quality of S-VHS surpasses that of VHS and is comparable to the 3/4" formats, it is highly recommended for video microscopy. Digital-VHS (D-VHS) is the newest type of video format available. Video tape recorders for D-VHS accept VHS and the S-VHS tapes but not vice versa. D-VHS recorders can be used to record and play NTSC as well as PAL standard. Furthermore, the D-VHS is capable of bit-stream-recording and thereby allowing recording of compressed digital data sets. If connected to a serial computer interface, the system can record computer data, audio data, as well as video sequences. ii. The 3/4 inch format Video tape recorders of the 3/4" format using U-matic cassettes were widespread in scientific laboratories and among other semi-professional users. The U-matic format has now been substituted by the manufacturers by the Betacam system. Both the U-matic video tape recorders and the cassettes are about five times as expensive as the 1/2" ones, while the Betacam system is even more expensive. Their value is in the higher quality of the recordings on the thick or, more stable tape material. This is important if the user often needs still frames or search mode, because these modes put a severe strain on the tape. The 3/4" format offers a vertical resolution of approximately 340 lines instead of only the 250-300 lines of 1/2" format (BAV mode). There is no difficulty in copying from one format to another provided that the two video tape recorders have the same video standard (e.g. NTSC). 136

3: Video microscopy However, bear in mind that you will lose quality when copying from 3/4", SVHS, or D-VHS onto 1/2" VHS while transferring from 1/2" onto the better format tapes will not gain any further resolution. While the 3/4" Betacam cassette format is in use for professional recording, the U-matic format has been replaced by the S-VHS format in most video microscopy laboratories. iii. Video tape quality There are very different qualities of tape material obtainable. For video microscopy where high density and low noise recording is required only high quality tapes should be used. In addition, there are special tapes designed for still-frame operation, a technique very often used in video microscopy. The maximum recording time per VHS or S-VHS cassettes is six hours but the stronger one or two hour tapes are recommended. 5.1.4 Video tape recorders Video tape recorders used in video microscopy should have several special features. For taking photographs off the monitor a good still-frame capability is required. To add comments later to a video film without erasing the recordings you need a video tape recorder with AUDIO DUB capability. Most of these recorders have two audio channels which can be played back separately or synchronously. It is also convenient to have a remote control, especially if the video tape recorder is out of reach. 5.1.5 Time lapse recording Various microscopic specimens such as, for example slow particle motions or the progress of cell division, require time lapse recording, as they become apparent only after speeded up play back. A time lapse recorder may be used in parallel with a normal speed video tape recorder so you have the same sequences both in real time and time lapse for comparison. Previously recorded real time scenes may also be speeded up later with a time lapse recorder. There are several time lapse video tape recorders available which are designed to record for up to 400 hours or more onto a regular two hour tape for observation and surveillance applications. Note that when played back at normal speed the recordings of most of the cheaper surveillance video tape recorders show considerable interference and noise bars so that high grade time lapse recorders are needed. However, clear still-frames can be obtained from all time lapse recorders. Time lapse recorders use regular 1/2" or 3/4" cassettes but recorded tapes cannot usually be played back on a standard speed video tape recorder of the same format. During time lapse recording the tape moves very slowly past the incessantly rotating video head, thus straining the tape. Therefore, only new and best quality tapes should be used. Sometimes animation control units (e.g. from EOS Electronics AV Ltd., or from AVT GmbH), are alternatives to time lapse recorders suited for the 137

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen documentation of very slow movements such as cell growth, locomotion, or division (72). These units are connected to an edit video tape recorder and permit recording of one or several frames at intervals of 10-999 seconds. Played back in real time such recordings show at least a X 100 time lapse effect, but they are free of image distortions. 5.1.6 Recording video-microscopic sequences Prior to recording it is important to check the equipment, ensuring all plugs and switches are in their proper positions. A suggestion as to how to connect the system components is shown in Figure 1. The video tape recorder should receive the signal directly from the video processor and not 'at second-hand' from a monitor. For this the signal has to pass the recorder prior to display on the monitor in the so-called E-to-E (electronics to electronics) mode. With some recorders this E-to-E mode picture can only be obtained when the RECORD button of the video tape recorder is pressed. If the monitor is the last set to receive the video signal, it has to be 'terminated'. This means that the 75 ft switch at the back of the monitor must be set to '75 ft' position to avoid disturbances. With monitors operating in the E-toE mode the switch is set to 'HI-Z' position. Likewise, this must be done for all the other pieces of equipment in line. If there is no switch 75 ft/HI-Z the VIDEO OUT connector of the last piece has to be terminated with a plug containing a built-in 75 ft resistance. More recent equipment is automatically terminated. During image processing brightness and contrast of the image are altered electronically. It is strongly recommended to set the brightness and contrast controls of the monitor always to their standard positions. This measure will provide scenes of the same brightness level which will fit together properly when they are edited or processed further. The timer and the scale bar are added to the video image by most image processors. To estimate the correct magnification, especially when photographing off the monitor record the scale of an object (stage) micrometer both in horizontal and vertical position. This is used to calibrate any scaling function or scale bar (see Figure 6) of the processor and to adjust the horizontal and vertical axes of the monitor. Too high a level of the video signal gives rise to snowy and noisy images when scenes are replayed from the video tape recorder. It is, therefore prudent to check this signal (at the VIDEO OUT connector of the image processor or video tape recorder) with the aid of an oscilloscope. A video level higher than 1 V will cause distortion in the bright regions of the image. The recording should be checked by playing back the tape. If the video signal is too high, the service personnel should be asked to reduce the output video signal to a maximum possible value of 1 V. Although the video recording level is controlled automatically we have to adjust the audio recording level manually on some recorders, unless they are 138

3: Video microscopy equipped with an audio level limiter which minimizes audio distortion at the peaks. It is very helpful to record vocal comments during working at the microscope, for example such events as changes of optics, the position of the specimen, or the focal plane (note presence of microphone in Figure 1). If the video tape recorder offers two separate audio channels and the audio dubbing feature one should use channel 2 for these comments, as one can add a second commentary afterwards (dubbing) to the previously recorded scenes only onto channel 1. Recorded tapes can be protected against accidental erasure. For that purpose the red cap on the bottom of U-matic cassettes is removed, or respectively, the plastic tab at the back of 1/2" cassettes is broken off. If you later decide to record again onto these tapes, replace the button or affix adhesive tape over the safety tab slot. The unintentional use of erasure protected tapes is the likely cause if the video tape recorder stops working when the RECORD mode is activated. Some frequent mistakes, which should be avoided, are the following. It is very annoying to the person who evaluates the images if the sequences are of short duration. Some scientists tend to try to improve the focus of the microscope continually while recording, or to shift the specimen around, incessantly expecting better positions. These faults become conspicuous especially when using a time lapse recorder. During recording one must remember that such mistakes cannot be improved afterwards and one ought to keep in mind the designed application of the scenes. For example, scenes which are to be used for motion analysis or other evaluation should last at least five, better ten, minutes without changing focus or stage position. 5.1.7 Recording pseudocolour sequences The pictures obtained by the camera and fed into image processors are usually monochrome (B/W). For special purposes, such as, for example, to enhance the contrast by adding colours or simply to get more aesthetical pictures, many image processors permit assignment of different colours to the different grey values (pseudocolour or false colour). Thus all areas of the same grey value are coloured equally. All generated colours are coded as a mixture of the three basic colours red, green, and blue (RGB) and split up into the corresponding colour channels. This RGB signal can be displayed at high image quality but only on monitors with RGB input capability. To record coloured images a composite video signal of one of the international standards PAL, SECAM, or NTSC is needed. Several image processors provide only analogue RGB output. These four or five signals (R, G, B, composite SYNC, or horizontal and vertical SYNC) have to be encoded into the composite standard colour video signal. For this purpose a colourencoder for either PAL, SECAM, or NTSC is required. This encoder is connected between the RGB outputs of the image processor and the video input of the video tape recorder. The encoder, however, requires a very regular and 139

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen exact SYNC signal, as can only be supplied by a TV camera. It is not difficult to add colour to original microscopic scenes coming directly from the TV camera, through the image processor, and to record these scenes after encoding the RGB signal. To colour previously recorded sequences from a tape, the video tape recorder as input source for the image processor may not provide as regular a SYNC signal as would be required. In this case, a time base corrector (TBC) may help and is connected either to the TBC socket (if available) of the first video tape recorder or to the encoder. Using a TBC is often advisable as it always provides the best possible playback picture, whether for copying, editing, mixing two signals, or additional processing. 5.1.8 Digital storage media: an outlook A whole range of digital storage devices that might well replace the video tape recorders in the future are presently being available or about to be introduced. Generally it should be noted that a two hour movie with sound would require in the order of 4 GByte storage, depending on the algorithm of compression applied. From the usefulness of the different storage media in our field we can say the following: i. Hard disks and RAM of PCs hold nowadays typically 5-10 GBytes or 100 MBytes respectively. This means that they are very useful as temporary storage during accumulation of special scenes, editing and preparing time lapse sequences. Hard disks are the typical intermediate storage for slowly writing to other mass storage devices such as CD-ROM or DVD-ROM which are presently less suitable for direct recording. The writable CD-ROM with a capacity of 650 MByte is suitable for short video clips only, however for archiving of up to a few thousand single frames this would be the medium of choice. Typical B/W images are 0.4 MByte, colour images more than 1 MByte, sometimes less, depending on the resolution and degree of compression. ii. The digital-VHS (D-VHS) tape has similar handling properties and quality as S-VHS, with the advantage of avoiding losses in quality during copying processes. Since this technology is relatively recent only few companies produce D-VHS recorders. iii. The digital video device (DVD) system was introduced for the home entertainment market. One DVD-ROM disk may hold a full two hour movie. With the introduction of DVD writers this system can be employed to store video demo and raw data. This may become a very useful technology when the writing process is possible at real time rather than sequences have to be exported slowly from hard disk. The earlier technology of optical disk memory recorders (OMDR) is now extinct because the disks were too bulk, not rewritable, and too expensive (over 200.00 EURO). 140

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5.2 Obtaining printouts for presentation and publication To obtain printouts for presentation one can choose from a whole range of methods. While the 35 mm camera is still occasionally used to obtain still frames from a video sequence, digital image capturing and printing, mainly performed on PCs, has become the technique of choice for obtaining publication quality digital images and prints. Since full documentation and handbooks come with the equipment and software for the latter techniques, it may suffice to outline the types of hardware in use. 5.2.1 Digital prints i. Frame grabber boards A scientific quality frame grabber is not only necessary for use with high-end CCD cameras and other imaging equipment at a resolution of 640 X 480 (768 X 512 for CCIR) pixels but it is also needed to capture images from input sources including S-VHS and RGB tapes or live video. Once the image is captured it can be processed with a large number of appropriate imaging programs such as PhotoShop (Adobe Systems, Inc.) or Paint Shop Pro (Jasc Software, Inc.). To obtain maximum image information, brightness and contrast must first be correctly adjusted at the level of the A/DC (digitizer) by applying the appropriate LUT and then optimized for presentation at the level of the digitized image (see Section 1.3.4). Special spatial imaging filters can be employed to remove background noise and to improve the signal-to-noise ratio. Manufacturers of frame grabbers include Data Translation, Inc., Fast Electronic GmbH, HaSoTec GmbH, Imaging Technology, Inc., Matrox Electronic Systems Ltd., Scion Corp., or Silicon Graphics, Inc. ii. Video printers Some effort has to be made to obtain publication quality prints. This is achievable with some high-end video printers which have a built-in frame grabber. Some of these printers offer lower quality hard copies on thermographic paper, which are sufficient for many uses such as documentation. Video printers (e.g. Sony Corp. or Mitsubushi Corp.) are connected to the VIDEO OUT socket of the video tape recorder. The video signal should pass the printer in the E-to-E mode to a control monitor. When the PRINT button is pressed the actual frame or an average of the last few frames is stored and printed within a few seconds. The print quality depends on the resolution of the printer (its number of grey levels, regularly 256), and on the quality of the paper used. The advantage of such video printers is the instant availability of a hard copy. The latest video printers attain almost photographic quality and have become available also in colour mode. iii. High quality printing from digital sources Several computer-controlled, high quality printing devices are available, e.g. 141

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen laser printers, dye sublimation printers, and ink jet printers, all of which are now capable of producing photo-quality printouts. Choosing among the different printing techniques will largely depend on budget and amount of printing needed. Dye sublimation printers are rather expensive and have high running costs (up to a few EURO for one A4 page). Laser printers are also expensive but running costs are low, while ink jet printers are cheapest but create higher costs for the ink cartridges and high-glossy, photo quality paper. iv. Digital slide makers Once video images are digitized they can also be printed directly to slide film. Several companies provide such slide printing devices, e.g. Eastman-Kodak Company, Nikon Europe B.V., Polaroid Export Europe. Printing a slide is carried out similarly to printing to paper. While the slide makers are optimized for colour slides, they can also be used to generate B/W slides either by using a special B/W slide film or by using a B/W negative film in combination with a reverse B/W developer kit. In contrast to colour film full saturation in black can only be achieved using B/W film. 5.2.2 Photographic prints If no high quality printer is available photographs can be taken from the monitor with a 35 mm single lens reflex camera. The camera should be equipped with a 90-100 mm lens (and possibly a close-up lens) or an appropriate macro lens. Automatic exposure is not always an advantage so a camera capable of manual exposure is appropriate. Most of the latest cameras are supplied with an ultra-fast photocell, which is unfortunately even affected by the running scan ray, which is invisible to the human eye. Therefore the information from the exposure meter fluctuates widely and pictures taken with automatic exposure become often either too dark or too bright. Exceptions are the Olympus OM-2 and OM-4 cameras which do not fix the exposure at the moment the shutter is released but measure continuously during the entire exposure (off-the-film). Likewise worth mentioning are cameras with very fast vertical shutters as found in more recent cameras, which avoid the scan bars. To shoot B/W pictures we need moderately fast negative film such as Agfapan 100, Kodak T-Max 100, or Ilford FP4. For colour slides and colour prints load the camera only with reversal (slide) film. This is because commercial laboratories will be overtaxed to print correctly the mostly very unnatural colours of a false colour display from negative film. Using reversal film one can point out that the colours of the print should correspond to those on the slide. Colour reversal films for good results are for example the Fuji-Velvia, Ektachrome 64, or Agfa RSXII. For shooting photographs it is advisable to use a high resolution monitor with a small screen. Mount the camera on a firm tripod and adjust it at right 142

3: Video microscopy angles to the screen. The level of the camera lens should correspond to the centre of the screen. The picture in the viewfinder may show either the full monitor with some dark background around it or only a section of the screen. Try photographing with a shutter speed of 1/8 sec or longer. Shorter times (even 1/15 sec) can result in heterogeneous pictures because of the video scan rate. When shooting the running image at 1/8 sec exposure time the camera effects a kind of averaging by integrating about four frames. This reduces the noise but may be totally inappropriate for scenes with rapidly moving objects. In these cases it is better to shoot still-frame pictures. Pressing the PAUSE button of the recorder provides usually only one field, that is half the number of video scan lines as the running picture. The consequence is an image with clearly visible scan lines. It is better to store (freeze) a whole frame (two fields) in the image processor or use tape recorders with multiple heads that display whole frames also in still mode.

5.3 Preparing and presenting video sequences To present video sequences to small audiences monitors or LCD projecting devices for overhead projectors are used, while for larger audiences video and data projectors are essential (the complicated and expensive copying to 16 mm movie film is now obsolete). With the improved display and projector technology a whole range of analogue and digital projectors is available (e.g. from Barco International, Sharp, or Sony Corp.) which allows the presentation of video signals equally well from analogue sources (tape) or digitally from hard disk or CD-ROM of a PC or laptop. Usually tapes recorded in the laboratory are not directly suitable for presentation to an audience so that a demonstration tape or clip with all scenes in suitable sequence and of proper lengths have to be compiled. Today this is done conveniently in a fully digital way with dedicated software packages or with special digital video editing hardware on the PC, which is supplied with sequences from various analogue (tape) or digital sources (hard disks, CDROMs, Internet, etc.). However, for those who still need to do it the classical (analogue) way with two video tape recorders the following hints may be helpful. 5.3.1 Copying and editing video tapes i. Preparatory considerations for editing To copy video sequences two recorders of the same standard (PAL, SECAM, or NTSC) are required. It is possible to copy from time lapse video tape recorders with variable recording speeds onto standard speed recorders. Before beginning to edit it is profitable to compose a script where all titles and scenes are put together consecutively. For titles schedule about five seconds, for long titles the time allowance should be sufficient to enable them to be read 143

Dieter G. Weiss, Willi Maile, Robert A, Wick, and Walter Steffen slowly. Scenes should run for periods of time not shorter than 20 seconds, only singular scenes may be longer than one minute, important ones can be repeated. For the copying process try to improve the video signals as much as possible. If available, the same video tape recorder as used for the original recording should be used since even recorders of the same standard and format are not always completely compatible. This is caused above all by different mechanical adjustment (e.g. the stretch of tape) of the different sets. Checking the quality of the video signal is described in Section 5.1.2. In the following description the source video tape recorder with the original tape is called video tape recorder I, the receiving recorder with the editing tape is video tape recorder II. ii. Procedure for editing video tapes Before copying advance all source tapes to the beginnings of the required scenes, so that one scene after another can be added quickly onto the second tape. The original cassettes should be protected from accidental erasure. To obtain the more pleasing black frames instead of noise and noisy bars on parts of tape II which will not be recorded on, one should first prepare a 'black' tape by running the tape (new or used) completely through in recording mode. Many video tape recorders will assure a smooth transition between recorded scenes when the procedure outlined in Table 3 is applied. In STOP mode (Table 3) the last few seconds of the preceding scene will be erased, but the scenes are smoothly added without frame breakdown at the splicing point. It is only with high grade video editing equipment possible to insert or exchange one scene for another between sequences already recorded (i.e. insert editing) in this same way without noisy frames resulting at the end of the scene. It may in some cases become necessary to further correct the signal from video tape recorder I by using a time-base corrector (TBC) which is also useful when adding a time bar (from a separate time date generator or an image processor), a scale bar, or other overlays to an existing video sequence during copying. 5.3.2 Digital video editing Due to the large volume of data video capture and playback require a high level of computer processing power. Even for a short video sequence the system has to deal with a large volume of data (15^000 images for 10 min at 25 f.p.s.). Processing of full screen video sequences (640 X 480) would require a P200 Pentium processor or better, a minimum of 64 MB RAM, and 4 GB of free hard disk space. There are several systems using different approaches available and we mention only one typical example. Many details and useful hints on the production and presentation of digital movies can be found in ref. 73. 144

3: Video microscopy Table 3. Assembly editing of video tapes for smooth transitions between scenes Video tape recorder II (editing tape)

Video tape recorder I (original tape)

Rewind the tape to the start. Run the tape for about 5 sec in PLAY mode as a leader. Press the PAUSE button, then the PLAY and RECORD button simultaneously without releasing PAUSE. Locate the prospected scene or title and start the tape (PLAY mode) about 5 sec before beginning of this scene. At the beginning of the scene to be recorded release the PAUSE button, press it again to stop the recording. Change the cassette or wind to the next sequence; start the tape about 5 sec before the scene starts. At the end: press the STOP button and rewind the tape to check it. When tape recorder II has been switched to STOP: Prepare the next sequence. Rewind the tape briefly to inspect the last scene. A few sec before its end press the PAUSE button, then the RECORD and PLAY button simultaneously without releasing the PAUSE button. Start the tape (PLAY) about 5 sec prior to the beginning of the desired scene. At the beginning of the scene to record release the PAUSE button.

One way is to control a set of tape recorders from a PC with a special hardware/software combination allowing to run video sequences one after another through the image processor, improve contrast etc., select from various types of smooth or other formats of transitions between the scenes, and add them to an editing tape (e.g. Fast Video Machine from Fast Electronic GmbH). A second approach requires a software package such as Premiere (Adobe Systems, Inc.) with a high performance scientific quality frame grabber capable of acquiring not only single frames but live video sequences (video acquisition board). The software package performs digital image processing functions instead of single frames (as PhotoShop or related packages) on whole sequences of stored, digital images. Dedicated programs such as Premiere (Adobe Systems, Inc.) will provide all the essential tools for compiling video clips, adding special effects, composing a complete movie film, and output it in one of a variety of different file formats. 145

Dieter G. Weiss, Willi Maile, Robert A. Wick, and Walter Steffen Several, fully digital stand-alone systems, especially semi-professional and professional ones, are available for sometimes forbiddingly high prices. A reasonably priced exception is for instance the Casablanca system (MS MacroSystem Computer GmbH). It is capable to acquire many scenes from analogue and digital video sources, store them digitally and put them to order according to a prepared storyboard with the selected transition effects, add titles and audio, and copy the result to a variety of analogue or digital tape formats. Other manufacturers offer partially different, interesting approaches such as MetaMorph (Universal Imaging Corp.), EDITBOX (Quantel Ltd.), or the various versions of video software for Silicon Graphics computers.

Acknowledgements The authors are greatly indebted to the late Bob Allen (1927-1986), Shinya Inoue, and the other colleagues of the Woods Hole video microscopy community for sharing their knowledge and experience, for numerous discussions and for valuable advice.

References 1. Inoue, S. (1981). J. Cell Biol., 89,346. 2. Inoue, S. and Spring, K. R. (1997). Video microscopy, 2nd edn, p. 741. Plenum Press, New York. 3. Allen, R. D., Allen, N. S., and Travis, J. L. (1981). Cell Motil., 1,291. 4. Allen, R. D., Travis, J. L., Allen, N. S., and Yilmaz, H. (1981). Cell Motil., 1, 275. 5. Allen, R. D. and Allen, N. S. (1983). J. Microsc., 129,3. 6. Lange, B. M. H., Sherwin, T, Hagan, I. M., and Gull, K. (1996). Trends Cell Biol., 5, 328. 7. Suzaki, E., Kobayashi, H., Kodama, Y., Masujima, T., and Yerakawa, S. (1997). Cell Motil Cytoskel., 39, 215. 8. Allen, R. D., Weiss, D. G., Hayden, J. H., Brown, D. T., Fujiwake, H., and Simpson, M. (1985). J. Cell Biol, 100,1736. 9. Weiss, D. G. (1986). J. Cell Sci. Suppl, 5,1. 10. Chalfie, M. and Kain, S. (1996). GFP, green fluorescent protein: strategies and applications. John Wiley & Sons, New York 11. Oshiro, M. (1998). Methods Cell Biol, 56, 45. 12. Hecht, E. and Zajac, A. (1984). Optics. Addison-Wesley Publishing Co., Reading, MA. 13. Weiss, D. G. (1987). Mol. Toxicol., 1, 465. 14. Russ, J. C. (1994). The image processing handbook, 2nd edn, p. 674. CRC Press, London. 15. Erasmus, S. J. (1982). /. Microsc., 127, 29. 16. Anonymous (1995). Am. Lab., April, 25. 17. Wick, R. A. (1987). Appl Opt., 26, 3210. 18. van Vliet, L. J., Sudar, D., and Young, I. T. (1998). In Cell biology: a laboratory handbook (ed. J. E. Celis), 2nd edn, Vol. 3, p. 109. Academic Press, New York. 146

3: Video microscopy 19. Inoue, T. (1993). In Electronic light microscopy (ed. D. Shotton), p. 95. Wiley-Liss, New York. 20. Foskett, J. K. (1993). In Optical microscopy: emerging methods and applications (ed. B. Herman and J. J. Lemasters), p. 237. Academic Press, New York. 21. Kuznetsov, S. A. and Weiss, D. G. (1998). In Cell biology: a laboratory handbook (ed. J. E. Celis), 2nd edn, Vol. 3, p. 344. Academic Press, New York. 22. Bennett, H. S. (1950). In Handbook of microscopical techniques (ed. C. E. McClung), p. 501. Harper & Row (Hoeber), New York. 23. Piston, D. W. (1998). Biol. Bull., 195,1. 24. Inoue, S. (1961). In The encyclopedia of microscopy (ed. G. L. Clark), p. 480. Reinhold, New York. 25. Hansen, E. W., Conchello, J. A., and Allen, R. D. (1988). J. Opt. Soc. Am., A5, 1836. 26. Schnapp, B. J. (1986). In Methods in enzymology (ed. R. B. Vallee), Vol. 134, p. 561. Academic Press, Orlando. 27. Brenner, M. (1994). Am. Lab., April, 38. 28. Kachar, B. (1985). Science, 227, 766. 29. Dodt, H.-U. and Zieglgansberger, W. (1994). Trends Neurosci., 537,453. 30. Weiss, D. G. and Maile, W. (1993). In Electronic light microscopy (ed. D. M. Shotton), p. 105. Wiley-Liss, New York. 31. Ellis, G. W. (1985). J. Cell Biol., 101, 83a. 32. Lichtscheidl, I. and Url, G. W. (1987). Eur. J. Cell Biol., 43, 93. 33. De May, J. (1983). In Immunocytochemistry (ed. J. M. Polak and S. W. Van Noorden), p. 92. Wright-PGS, London. 34. Hoffman, R. (1977). J. Microsc., 110,205. 35. Ellis, G. W. (1978). In Cell reproduction (ed. E. Dirksen, D. Prescott, and C. F. Fox), p. 465. Academic Press, New York. 36. Allen, R. D. (1985). Annu. Rev. Biophys. Biophys. Chem., 14,265. 37. Allen, R. D., David, G. B., and Nomarski, G. (1969). Z. Wiss. Mikr. Mikrotech., 69,193. 38. Bereiter-Hahn, J. and Vesely, P. (1989). In Cell biology: a laboratory handbook (ed. J. E. Celis), 2nd edn, Vol. 3, p. 54. Academic Press, New York. 39. De Brabander, M., Nuydens, R., Geuens, G., Moeremans, M., and De Mey, J. (1986). Cell Motil. Cytoskel., 6,105. 40. Geerts, H., De Brabander, M., Nuydens, R., Geuens, S., Moeremans, M., and De Mey, J. (1987). Biophys. J., 52,775. 41. Mathog, D., Hochstrasser, M., and Sedat, J. W. (1985). J. Microsc., 137, 241 and 253. 42. Wang, Y. L. (1998). Methods Cell Biol., 56,305. 43. Kachar, B., Evans, D. F., and Ninham, B. W. (1984). J. Coll. Interface Sci, 100,287. 44. Spring, H. and Trendelenburg, M. (1990). J. Microsc., 158, 323. 45. Sase, L, Miyata, H., Corrie, J. E., Craik, J. S., and Kinosita, K. Jr. (1995). Biophys. J., 69,323. 46. Vale, R. D., Reese, T. S., and Sheetz, M. P. (1985). Cell, 42, 39. 47. Weiss, D. G., Langford, G. M., Seitz-Tutter, D., and Keller, F. (1988). Cell Motil. Cytoskel., 10,285. 48. Haugland, R. P. (1996). Handbook of fluorescent probes and research chemicals, 6th edn, p. 679. Molecular Probes, Eugene, OR. 147

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49. Mitchison, T. J., Sawin, K. E., and Theriot, J. A. (1998). In Cell biology: a laboratory handbook (ed. J. E. Celis), 2nd edn, Vol. 3, p. 127. Academic Press, New York. 50. Mittal, B., Sanger, J. M., and Sanger, J. W. (1987). J. Cell Biol, 105,1753. 51. Peters, R. (1985). Trends Biochem. Sci., 10,223. 52. Kapitza, H. and Jacobson, K. (1986). In Techniques for the analysis of membrane proteins (ed. C. I. Ragon and R. J. Cherry), p. 345. Chapman and Hall, London. 53. Yanagida, M., Morikawa, K., Hiraoka, Y., Matsumoto, S., Uemura, T., and Okada, S. (1986). In Application of fluorescence in the biomedical sciences (ed. D. L. Taylor, et al.), p. 321. Alan R. Liss, New York. 54. Kron, S. J. and Spudich, J. A. (1986). Proc. Natl. Acad. Sci. USA, 83, 6272. 55. Wampler, J. E. (1986). In Application of fluorescence in the biomedical sciences (ed. D. L. Taylor, et al.), p. 301. Alan R. Liss, New York. 56. Yoshimoto, Y., Iwamatsu, T., Hirano, K., and Hiramoto, Y. (1986). Dev. Growth Differ., 28, 583. 57. O'Kane, D. J., Lingle, W. L., Wampler, J. E., Legocki, M., Legocki, R. P., and Szalay, A. A. (1988). Plant Mol Biol, 10,387. 58. Purves, D. and Voyvodic, T. (1987). Trends Neurosci., 10, 398. 59. Blasdel, G. G. and Salama, G. (1986). Nature, 321, 579. 60. DeBiasio, R., Bright, G. R., Ernst, L. A., Waggoner, A. S., and Taylor, D. L. (1987). J. Cell Biol, 105, 1613. 61. Zecevic, D. (1996). Nature, 381,322. 62. Shotton, D. (1993). In Electronic light microscopy (ed. D. Shotton), p. 39. WileyLiss, New York. 63. Cohn, S. A, Ingold, A. L, and Scholey, J. M. (1987). Nature, 328,160. 64. Weiss, D. G., Keller, F., Gulden, J., and Maile, W. (1986). Cell Motil CytoskeL, 6, 128. 65. Weiss, D. G., Galfe, G., Gulden, J., Seitz-Tutter, D., Langford, G. M., Struppler, A., et al. (1990). In Biological motion (ed. W. Alt and G. Hoffmann). Lecture notes in biomathematics, Vol. 89, p. 95. Springer-Verlag, Berlin. 66. Soll, D. R. (1988). Cell Motil. CytoskeL, 10, 91. 67. Wampler, J. E. and Kutz, K. (1989). Methods Cell Biol., 29,239. 68. Shaw, P. J. (1993). In Electronic light microscopy (ed. D. M. Shotton), p. 211. Wiley-Liss, New York. 69. Tsien, R. Y. and Poenie, M. (1986). Trends Biochem. Sci., 11, 450. 70. Bright, G. R., Rogowska, J., Fisher, G. W., and Taylor, D. L. (1987). BioTechniques, S, 556. 71. Bright, G. R., Fisher, G. W., Rogowska, J., and Taylor, D. L. (1989). Methods Cell Biol., 30,157. 72. Allen, T. D. (1987). J. Microsc., 147,129. 73. Waterman-Storer, C. M., Shaw, S. L., and Salmon, E. D. (1997). Trends Cell Biol., 7, 503.

Further reading Video microscopy Ref. 2. Sluder, G. and Wolf, D. E. (ed.) (1998). Video microscopy. Methods in cell biology, Vol. 56. Academic Press, New York. 148

3: Video microscopy Shotton, D. M. (1998). In Cell biology: a laboratory handbook (ed. J. E. Celis), 2nd edn, Vol. 3, p. 73 and p. 85. Academic Press, New York. Shotton, D. M. (ed.) (1993). Electronic light microscopy: the principles and practice of video-enhanced contrast, digital intensified fluorescence and confocal laser scanning microscopy, p. 355. Wiley-Liss, New York. YEC microscopy Ref. 36. Weiss, D. G. (1998). In Cell biology: a laboratory handbook (ed. J. E. Celis), 2nd edn, Vol. 3, p. 99. Academic Press, New York. VIM Refs 6,18,48, and 71. Taylor, D. L., Waggoner, A. S., Murphy, R. F., Lanni, F., and Birge, R. R. (ed.) (1986). Applications of fluorescence in the biomedical sciences. Alan R. Liss, New York. Digital image processing Refs 14 and 62. Jahne, B. (1991). Digital image processing, p. 337. Springer-Verlag, Heidelberg. Anonymous (1985). Image analysis. Principles and practice. Published by Joyce-Loebl, distributed by IRL Press. Baxes, G. A. (1984). Digital image processing. Prentice-Hall, Englewood Cliffs, New York. Instrumentation Refs 2 and 23. Guide to biotechnology products and instruments. (1988). Science, 239, G73 and G164-G180.

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4

Microscopy of chromosomes A. T. SUMNER and A. R. LEITCH

1. Introduction Chromosomes have been studied for something like 120 years, and for most of this period they were visualized using uniform staining with a variety of dyes. This was adequate for studies of the gross structure and behaviour of chromosomes, but identification of individual chromosomes was difficult, being based only on chromosome morphology and size, and specific chromosome substructures could not be studied. Just over 25 years ago, two new techniques emerged that revolutionized the study of chromosomes: (a) Chromosome banding, a series of staining techniques that produced patterns of longitudinal differentiation on chromosomes, and allowed identification of specific chromosomes and chromosomal substructures. (b) In situ hybridization, which labels specific DNA sequences in chromosomes. Methods for chromosome banding are described in Section 4, and methods for in situ hybridization in Section 5. In recent years, immunocytochemistry has been applied increasingly to studies of chromosomes; these methods are described in more detail in Chapter 5. However, methods of preparing chromosomes for immunocytochemistry do differ from routine chromosome preparation methods, and a range of preparation procedures is described in Section 2. Methods of recording images of chromosomes are of vital importance: fluorescent images are increasingly popular for a variety of reasons (greater sensitivity and contrast, and multiple labelling techniques), yet can suffer from low brightness and impermanence. It is therefore not surprising that some of the technically most advanced equipment for image capture has been applied to the study of chromosomes, although photographic film will no doubt also continue to be important for the foreseeable future. Methods for recording images of chromosomes are discussed in Section 6.

2. Methods of preparing chromosomes In this section it is only possible to give a few examples of how to obtain mitotic cells, and of methods of preparing them for the different procedures

A. T. Sumner and A. R. Leitch described later in, this chapter. For a wider variety of procedures, including different types of mammalian tissues and a variety of plants, lower vertebrates, and invertebrates, see refs 1-6. Methods for mammalian meiotic chromosomes are described in detail by Chandley et al. (7). There are two aspects to preparing chromosomes: obtaining growing cells with a sufficient proportion of mitotic cells to make it practicable to study their chromosomes, and spreading the chromosomes on to a microscope slide so that they can be seen easily. It is very hard work if only one cell in a thousand is dividing, but it may sometimes be necessary to use such material if it is difficult to obtain or grow. If the dividing cells are present at a few per cent, that is perfectly adequate for most purposes, although by using synchronized cells, preparations can be obtained in which over 90% of cells are in mitosis.

2.1 Routine preparation of mammalian chromosomes In this section, two methods of culturing mammalian cells will be described, and a single method of fixing them and spreading them on slides. Such preparations are suitable for plain staining of chromosomes, to count the number of chromosomes in the cell, to study chromosome aberrations, and to measure the amounts of DNA in chromosomes, for chromosome banding, and for in situ hybridization. They are not generally usable for immunocytochemical studies. 2.1.1 Culture of human blood lymphocytes This is a routine type of procedure used in clinical cytogenetics laboratories all over the world. A small blood sample is added to a suitable culture medium. The medium contains phytohaemagglutinin, which induces the lymphocytes to grow and divide. After an appropriate period of culture (usually two to three days), colchicine or some other inhibitor of micro tubule formation is added to arrest the cells in metaphase, and the cells can then be harvested, fixed, and spread. Protocol 1.

Human lymphocyte culture

Reagents • Culture medium: 8 ml F10 medium (Gibco BRL), 2 ml fetal calf serum, 0.1 ml phytohaemagglutinin (PHA), 0.01 ml antibiotic mixture (10000 U/ml penicillin, 10 mg/ml streptomycin)—keep in the refrigerator or cold room

• 10 ug/ml colcemid (Sigma)—keep in the refrigerator and protect from light

Method 1. Collect adult blood by venipuncture into a sterile container coated with lithium heparin (anticoagulant). Mix gently to prevent clotting. Blood

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4: Microscopy of chromosomes should only be taken by a qualified person, and all procedures must be carried out under sterile conditions. 2. Incubate 0.5 ml blood for 72 h at 37 °C in 5 ml culture medium. 3. 3 h before harvesting (i.e. 69 h after setting up the culture), add 0.1 ml colcemid solution. 4. Harvest cells (see Protocol 3).

2.1.2 Chinese hamster ovary (CHO) cells CHO cells are a continuously growing cell line. During interphase the cells remain attached to the substratum, but become detached during mitosis. A very high proportion of mitotic cells can therefore be obtained simply by shaking the culture vessel gently, and pouring off the culture medium, which will generally contain more than 90% mitotic cells.

Protocol 2. Culture of CHO (Chinese hamster ovary) cells Equipment and reagents • C02 incubator set at 37 °C • Inverted microscope with phase-contrast optics • 10 (jig/ml colcernid (see Protocol 1)

• 50 ml Falcon plastic culture vessels (Becton Dickinson) • RPMI 1640 medium containing 10% fetal calf serum (Sigma)

Method 1. Add approx. 106 CHO cells to the culture vessel, and make up to 8 ml with the culture medium. 2. Culture for approx. 48 h in an incubator at 37°C, ventilated with an air mixture containing 5% carbon dioxide. 3. Examine the culture flask using the inverted microscope. If the cells are nearly confluent, proceed to the next step; otherwise, return them to the incubator, and continue to examine the flasks at intervals until the required density of cells is attained. 4. Add 40 |xl colcemid, and culture for a further 3 h. 5. Harvest the mitotic cells by shaking the flask gently, and pouring off the supernatant culture medium, which contains mainly metaphase cells. 6. Treat the cells as described in Protocol 3.

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A. T. Sumner and A. R. Leitch 2.1.3 Fixation with methanol:acetic acid Protocol 3. Fixation and spreading of routine chromosome preparations The same procedure is used for both cultured human lymphocytes and for CHO cells. Equipment and reagents • Microscope slides, preferably with frosted ends. These can be bought already cleaned, but it is nevertheless advisable to clean them further by soaking them in acid:alcohol (1% concentrated HCI in absolute alcohol). Keep a large jar with a supply of slides in it, and take these out and clean them just before you are ready to harvest the cells. Slides should be wiped dry and polished with muslin.

» Bench-top centrifuge, with buckets for 10 ml tubes • Vortex mixer with speed regulator • 0.075 M (hypotonic) potassium chloride solution (5.6 g/litre) • Methanol:acetic acid fixative: 100% methanol, 100% glacial acetic acid (3 parts:1 part), prepared immediately before use—discard any unused fixative

Method 1. Place the culture medium containing the cells in a 10 ml centrifuge tube, and centrifuge at about 200 g for 5 min. 2. Pour off the supernatant, and add 10 ml hypotonic potassium chloride solution to the pellet of cells, with gentle shaking on the vortex mixer. Ensure that the cells have all dispersed, and stand for 10 min at room temperature. 3. Centrifuge again at 200 g, and pipette off the supernatant without disturbing the pellet of cells. 4. Disperse the cells with the vortex mixer, and, keeping the cells moving by gentle application of the vortex mixer, add the methanol:acetic acid fixative drop by drop until there is a great excess of fixative, then make up to approx. 10 ml. 5. Centrifuge the cells at 200 g again, and decant off the supernatant. 6. Add 10 ml fixative again, with vortexing, and centrifuge the cells as before. 7. Decant off the supernatant, add fixative, and centrifuge, as in step 6. 8. Decant off the supernatant and add sufficient fixative, with vortexing, to produce a slightly milky fluid. The precise appearance required at this stage to obtain a good number of cells on the slide, with good spreading of chromosomes, will be found by experience. 9. Take a clean microscope slide, and write on the frosted end the identification of the culture, plus any other relevant details (e.g. date).

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4: Microscopy of chromosomes 10. Using a polythene pipette, make sure the fixed cell suspension is thoroughly dispersed, and take an aliquot up into the pipette (0.10.2 ml should be ample at this stage). Drop two or three drops of the cell suspension on to different parts of the slide from a height of 5-10 cm. Breathe gently on the slide to aid spreading of the chromosomes. 11. Allow the cells to dry, and examine the slide with a phase-contrast microscope for density of cells and proportion of metaphases (this can easily be assessed with a x 10 objective); a x 40 objective may be used to assess the quality of spreading of the chromosomes if necessary. 12. If everything is satisfactory, prepare as many slides as required in the same way. 13. The fixed cell suspension can be stored overnight in the refrigerator; for longer storage, place in a freezer at -20°C. Allow the cell suspension to warm to room temperature, centrifuge, and resuspend in fresh fixative, before using again.

2.2 Preparation of cells by cytocentrifugation for immunocytochemical studies of chromosomes Preparations fixed in alcohol:acetic acid are generally unsuitable for immunocytochemical studies, as the fixative extracts or denatures many proteins. A radically different approach is therefore required to produce spreads without acid treatment. In general, such methods spread the chromosomes by cytocentrifugation, in which the cells are centrifuged on to a microscope slide. Such preparations may still contain a considerable amount of cytoplasm, which may have to be removed to allow access of immunocytochemical reagents. Fixation is not necessary, and indeed may be undesirable, before cytocentrifugation. Once the cells have been centrifuged on to the slide, they may be fixed in the most appropriate way for the antigen being studied, as different antigens differ in their susceptibility to destruction by fixation. The method given here has produced satisfactory results in the first author's experience, but other published methods are equally suitable (8). The quality of the chromosome morphology is unlikely to be as good as that obtained routinely with alcohol:acetic acid fixed preparations: some chromosomes may be very poorly spread, especially if the cells are too concentrated on the slide, while, particularly at the edges of the preparations, others may become excessively stretched. In between, there should be a proportion of cells in which the quality of the chromosomes is adequate.

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A. T. Sumner and A. R. Leitch Protocol 4. Cytocentrifuge preparation of chromosomes The method described here is suitable for cells cultured as described in Protocols 1 and 2. Equipment and reagents • Cytocentrifuge: any model appears to be equally suitable, but the instructions given below refer to a Shandon Cytospin 2 (quantities, centrifugation speeds, times, etc., may have to be adjusted for other models) • Phosphate-buffered saline (PBS): Dulbecco 'A' tablets, dissolved according to the manufacturer's instructions (Oxoid Ltd.) . 0.1% Triton X-100 (v/v) in PBS

• Stenman's hypotonic solution (9): 10 mM Hepes Na ((/V-2-hydroxyethyl) piperazineW-2-ethanesulfonic acid monosodium salt, 2.603 g/litre) pH 7, 30 mM glycerol (2.19 ml/ litre), 1 mM calcium chloride (CaCI2.6H2O, 0.2191 g/litre), 0.8 mM magnesium chloride (MgCI2.6H20, 0.1626 g/litre)—store in the refrigerator

Method 1. Place the culture medium containing the cells in a 10 ml centrifuge tube, and centrifuge at about 200 g for 5 min. 2. Pour off the supernatant, and add 10 ml Stenman's hypotonic solution to the pellet of cells, with gentle shaking on the vortex mixer. Ensure that the cells have all dispersed, and stand for 10 min in the refrigerator at approx. 4°C. 3. Disperse the cells in the Stenman's hypotonic, and add aliquots of 0.3-0.4 ml of the cell suspension to the centrifugation chambers of the Shandon Cytocentrifuge. 4. Centrifuge for 10-15 min at 1500 r.p.m. 5. Remove the slides from the Cytocentrifuge, lay them flat, and allow them to dry out. 6. Later (the same day or the next day), treat the slides with Triton X-100 solution at room temperature for 5-30 min to remove cytoplasm. 7. Wash for 5 min each in three lots of PBS. 8. Fix the preparation as required, and proceed with immunocytochemical labelling according to standard procedures (see Chapter 5, and also Protocol 9).

2.3 Preparation of chromosomes from plant cells Plant mitotic cells can be found in the apical meristems of roots and shoots, and in dividing tissues in ovaries, anthers, and endosperm. In addition they can be generated from tissue cultured materials. Whatever the source, it is important that the material is vigorous and healthy at the time of harvest. The quality of material is an important factor in obtaining high quality chromosome preparations. 156

4: Microscopy of chromosomes Root tips are the commonest source of mitotic divisions in plants. Typically, seeds are germinated on moist filter paper or tissues, or more rarely on 2% (w/v) agar overlaid with cellophane discs. In either way young, fresh, root tips can be obtained in many species. If it is impossible to obtain roots from germinating seeds, then try to obtain them from cuttings or root cultures. The final and least favourable option is root material from soil grown plants. In all cases make sure the roots are young, vigorous, and in almost all cases with a white apex. Other good sources of metaphases are young anthers and ovaries; meiotic tissues can also be obtained from anthers at later stages. However accumulating metaphases in buds is more problematic than it is with roots. The final source is tissue cultured material. Long-term cultures, especially suspension cultures, will almost certainly have rearranged karyotypes. However experimental manipulation of growing conditions and the use of cell cycle inhibitors can lead to high metaphase indices. 2.3.1 Accumulation of metaphases In general metaphase indices are lower for plant material than for mammalian material, although accumulation protocols have been developed which can substantially increase mitotic indices (10). Often it is desirable to spend time increasing the mitotic index of plant material to help chromosome banding and in situ hybridization experiments. Examine vigorously growing material first without any treatments. Then experiment with the protocols described below, which are simple and will provide a basis for accumulating metaphases. Excised root tips, buds, or other meristems are treated with one of the following solutions. Times and conditions need to be determined empirically. When accumulating metaphases in small seeded plants, treat the whole seedling. For big seeded plants, cut off the root tip. 0.01-0.05% (w/v) colchicine is ideal for most plants. Material should be placed in the solution for 3-6 h at room temperature or 16-24 h at 4°C. For cereals, treatment with aerated distilled water at 0°C (ice-water), typically for 24 h, is ideal. For some dicotyledonous plants, material can be placed in 2 mM 8-hydroxyquinoline for 1-2 h at room temperature followed by 1-2 h at 4°C. 2.3.2 Fixation and chromosome spreading There are two main ways to prepare plant material for chromosome studies: 'squashing' and 'dropping'. Dropping involves fixing material in ethanol: acetic acid (3:1), making a cell suspension with enzymes, and dropping the suspension on to a glass slide, much as is done for mammalian cells (Protocol 3) (1). This method gives the most sensitive in situ hybridization data and may be essential for detecting single and low copy DNA sequences (11). However, because there is no hypotonic step, the chromosomes tend to be poorly separated. Squashing, on the other hand, gives good separation of chromosomes, ideal for high quality chromosome banding, but at the cost of signal strength when performing in situ hybridization. The protocol below is a 157

A. T. Sumner and A. R. Leitch gentle squashing method which combines advantages of both methods and in our hands is successful. It is important through practice to develop a method that works for your material. Get good spreads before chromosome staining, as the quality will never improve after the staining! Under the phase-contrast microscope and especially the Nomarski differential interference microscope, cytoplasm can be seen easily. Avoid slides with too much cytoplasm overlying the cells. Chromosomes and interphase nuclei with nucleoli should be clearly seen. Cells and nuclei from meristematic cells after squashing should be round. They should not have rectangular outlines and bright walls. Such cells are usually poor and slides containing a lot of material like this should be discarded. The optimal density of cells is when individual cells do not overlap but are not so few that they are hard to find. Material with tiny chromosomes may be screened best using DAPI fluorescence, and slides that are selected washed in 2 X SSC (saline sodium citrate buffer, see Protocol 6) and immediately used for in situ hybridization. Poor slides give poor in situ hybridization results and it is worth screening carefully. Protocol 5. Fixing and spreading plant chromosomes Equipment and reagents • • • • • •

Acid cleaned slides (see Protocol3) Watch glass Hypodermic needles and forceps Razor blade Diamond pen Binocular microscope, and phase-contrast or Nomarski interference microscope • Fixative: ethanol, glacial acetic acid (3:1), freshly prepared . 10 x enzyme buffer pH 4.8: 40 mM citric acid, 60 mM sodium citrate—store as a stock solution at 4°C

• 1 x enzyme buffer: dilute 10 x enzyme buffer with water (1:9) • 2 x enzyme solution: 2% (w/v) cellulase (1.8% (w/v) dry powder from Aspergillus niger [Calbiochem] plus 0.2% (w/v) 'Onozuka' RS [Yakutt Pharmaceutical Ind]) and 10% (v/v) pectinase (Sigma) in 1 x enzyme buffer—store in aliquots at -20°C • 45% acetic acid • 2 ug/ml DAPI (4',6-diamidino-2-phenylindole) in 2 x SSC • Dry ice

Method Vigorously growing healthy material is fixed in freshly prepared fixative for at least 10 h at room temperature. Material can be stored for at least a month at -20°C. 1. Place 0.5-1 cm of root tip or individual anthers/ovaries into 1 x enzyme buffer in a watch glass. Replace solution continuously over 25 min period by pipetting fresh 1 x enzyme buffer into the watch glass. 2. Remove 1 x enzyme buffer with a pipette and flood material with 2 x enzyme solution. Incubate at 37°C. 3. Regularly check the state of digestion. Material should be close to

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4: Microscopy of chromosomes falling apart but still integral, e.g. root tips picked up by their cut end should not be able to support their own weight. Typical digestion time is 30 min. 4. Remove 2 x enzyme solution and replace with enzyme buffer. Wash for 25 min with continuous replacement of 1 x enzyme buffer. 5. Replace 2 x enzyme solution with fresh fixative by removing half the volume of 1 x enzyme buffer and replacing with fixative. Repeat three times and then three times in fresh fixative. 6. Remove fixative and replace with 45% acetic acid. 7. Under a binocular microscope excise meristematic tissues with the needles and place in a small drop of 45% acetic acid on a glass slide. In the case of root tips the area of interest is the white zone immediately behind the root cap. Do not take more tissue than is needed, as more tissue usually means a lower quality of spreading. Break up the material on the slide with the hypodermic needles. 8. Carefully add a 24 x 24 mm coverslip, making sure that no air bubbles are trapped. 9. Gently tap the material and monitor its dispersal with the phasecontrast or Nomarski microscope. 10. Place a filter paper over the coverslip, blot gently, and press lightly without shearing the coverslip. Examine the slide again. Regulate the pressure of spreading and the amount of tapping to optimize spreading. In general, the harder that it is pressed, the lower the quality of in situ hybridization. 11. Place good slides on dry ice, leave for 5 min, and flick off the coverslip with the razor blade. Leave slides to dry. 12. Select high quality slides under phase or Nomarski microscopes.

2.4 Assessment of the quality of chromosome preparations Chromosome preparation from animal cell cultures and plant meristems remains as much an art as a science, although attempts are being made to understand the process better (12). It is therefore necessary to be able to judge the quality of chromosome preparations as soon as the first slides are made, so that appropriate adjustments can be made at once to obtain satisfactory spreads from the material. Occasionally, cells fail to grow vigorously, or produce such a low mitotic index that they are virtually useless; if this occurs, seek advice from an expert in the material of interest. For routine, alcohol:acetic acid fixed preparations, the aim is to achieve metaphase spreads in which there are few if any overlapping chromosomes, yet none have been lost, while at the same time eliminating all the surrounding cytoplasm, which is likely to interfere with subsequent staining and labelling reactions. Possible remedies for a variety of faults are described in refs 3, 5, 159

A. T, Sumner and A. R. Leitch and 13. Although it is possible to stain slides quickly to assess their quality, it is quicker to look at the unstained preparations using a phase-contrast microscope, or even better, a Nomarski interference microscope if one is available. The advantage of the latter is that chromosomes, cytoplasm, and cellular components are easily visualized in unstained dry spreads. For cytocentrifuge preparations, it is important to have the right number of cells present: if they are too concentrated, the chromosomes will not spread, and if there are too few cells, the chromosomes may tend to be overstretched. To some extent these problems can be overcome by adding less or more cell suspension to the cytocentrifuge buckets, but it is probably better to dilute or concentrate the cell suspension.

3. Uniform (solid) staining of chromosomes Although it would be impossible to carry out a comprehensive chromosomal study of a species without using banding methods (see Section 4), there are still good reasons for using uniformly, non-banded chromosomes from time to time. The chromosomes do not suffer the distortion that often occurs with banding, the morphology of the chromosomes is clearer and the ends, in particular, are clearly visible, and chromosome abnormalities and fragile sites are more easily seen. Solid staining with methods such as Feulgen (2,4) is valuable for making DNA measurements. In addition, solid staining is valuable as a counterstain for various other banding and labelling techniques, as described below. Solid staining methods can be divided into two classes: transmitted light stains, such as aceto-Orcein, Giemsa, and Feulgen, which provide permanent preparations; and fluorescent stains such as DAPI, propidium iodide, and Acridine Orange, which are impermanent, but often very quick to carry out, provide good counterstains for fluorescent labelling (for in situ hybridization and irnmunocytochemistry), and can be used to measure DNA (both on slides and by flow cytometry). Full details of these methods will not be given here. For many purposes, indeed, it is not necessary to specify a precise procedure: an appropriate volume of water or any buffer that is to hand (e.g. phosphate-buffered saline) can be put into the staining vessel, and a small amount of a stock dye solution added. The chromosome preparations are stained until sufficient staining is obtained, and may be put back for longer if necessary. For fluorochromes, the dye concentration and staining time required may be surprisingly low. For the use of fluorochromes as counterstains to other procedures, see Protocols 9 and 12. Giemsa staining can be done very quickly and easily, but will stain any surrounding cytoplasm as well, a particular problem with chromosomes prepared by squash techniques (i.e. plant and invertebrate chromosomes). In such cases, it will be better to use aceto-Orcein (5) or Feulgen (2, 4) staining, although both of these methods are rather long and complicated. If a fluorescence procedure is acceptable (i.e. permanent preparations are not required) 160

4: Microscopy of chromosomes DAPI is appropriate, as its blue fluorescence cannot be confused with the red autofluorescence of chloroplasts, for example.

4. Chromosome banding Chromosome banding refers to the production of longitudinal differentiation along chromosomes by the use of special staining methods, in the absence of any structural differentiation (14). Specific chromosomes can be identified by their characteristic patterns, not only within a species, but" also in related species. Some types of banding draw attention to specific parts of chromosomes, and others are related to the functional properties of specific regions, such as nucleolar organizing regions (NORs) and kinetochores. Thus chromosome banding can also be used to study various aspects of chromosomal function. Banding patterns also draw our attention to certain aspects of chromosome organization which are of interest in their own right (14, 15).

4.1 The classification of chromosome bands Four different classes of chromosome bands have been recognized (14). Heterochromatic bands (C-bands, Section 4.2, Protocol 6) form discrete blocks on chromosomes, and are nearly always found at the centromeres, and sometimes also at terminal and interstitial sites. They commonly (but apparently not invariably) contain highly repeated DNA sequences, and lack conventional genes. Heterochromatic bands appear to contain distinctive proteins which may contribute to their compactness. Note that all the methods for heterochromatic bands stain only constitutive heterochromatin, but not facultative heterochromatin (such as the inactive X chromosome in female mammals). Euchromatic bands consist of a series of positively and negatively stained bands throughout the non-heterochromatic parts of the chromosomes. The euchromatic bands, which can be stained by such methods as G-banding (Section 4.3, Protocol 7), R-banding, or various fluorochromes, are largely confined to higher vertebrates (reptiles, birds, and mammals), although replication bands, which generally show patterns corresponding to those produced by the aforementioned methods, may well be universal (14). It is still not clear whether the lack of euchromatic bands in the chromosomes of almost all plants, invertebrates, and lower vertebrates genuinely represents a difference in the organization of the chromosomes in these organisms, or whether their absence is due to technical problems which have not yet been solved. An extensive listing of the properties of euchromatic bands is given by Holmquist (16). During chromosome condensation, dark G-bands fuse together, obliterating the pale bands between them, so that the chromosomes show fewer bands at metaphase than at prophase. The important practical feature of euchromatic bands is that they form distinctive patterns characteristic of each chromosome pair of a species, and can be used to identify that chromosome, even when translocated or rearranged. 161

A. T. Sumner and A. R. Leitch The NORs are those segments of the chromosomes that contain the genes for ribosomal RNA, and on which the nucleoli are formed. The NORs, which contain hundreds or even thousands of copies of the ribosomal genes, commonly appear as constrictions of the chromosomes. They can be stained specifically with silver (Section 4.4, Protocol 8). The final class of band is formed by staining the kinetochores, which are the sites of attachment of the spindle microtubules to the chromosomes. The preferred method for demonstrating kinetochores uses an immunocytochemical procedure (Section 4.5, Protocol 9). In the following sections, one principal banding method has been described in detail for each of the four classes of chromosome banding. There are, however, a considerable number of methods that may prove useful in various circumstances, and although there is no space here to describe the protocols, it is as well that the reader should be aware that other techniques are available. Technical details will be found in standard works (3, 5,14,17).

4.2 C-banding C-banding is the universal technique for demonstrating constitutive heterochromatin. In higher vertebrates this contributes to the complete characterization of the karyotype, and allows the study of certain types of chromosome evolution. In those organisms (plants, invertebrates, and lower vertebrates) that lack euchromatic bands, C-banding is of fundamental importance for the identification of chromosomes. Euchromatic bands cannot be demonstrated satisfactorily in meiotic chromosomes, even in higher vertebrates, and here again C-banding is useful for chromosome identification. Protocol 6. The BSG method for C-banding (20) Equipment and reagents • 0.2 M hydrochloric acid (17.2 ml concerntrated acid/litre) . 5% barium hydroxide in distilled water at 50°C. Warm 40 ml distilled water to 50°C in a Coplin jar in a water-bath; a few minutes before use add 2 g Ba(OH)2.8H20 and stir well to dissolve. The layer of barium carbonate that forms on the surface may be skimmed off if necessary, but does not usually cause any problems. Discard the solution after an hour or so.

• 2 x SSC at 60°C: 0.3 M sodium chloride, 0.03 M trisodium citrate (17.53 g NaCI and 8.82 g trisodium citrate per litre of distilled water)—discard the heated working solution after an hour or two • Giemsa: add 1 ml Gurr's Giemsa Improved R66 (6DH) to 50 ml buffer pH 6.8, made with Gurr's buffer tablets—discard after an hour or two, as the dye precipitates after dilution

Method 1. Allow the chromosome preparations to age for about a week before C-banding. 2. Put the chromosome preparations in 0.2 M hydrochloric acid for 1 h at room temperature.

162

4: Microscopy of chromosomes 3. 4. 5. 6. 7. 8. 9.

Rinse briefly with distilled water. Place in the barium hydroxide solution for 1-5 min at 50°C. Rinse thoroughly with distilled water. Immerse the slides in 2 x SSC at 60°C for 1 h. Rinse with distilled water. Stain with Giemsa, 45 min. Rinse the Giemsa solution off with distilled water and carefully blot the slides dry. 10. Allow the slide to dry thoroughly for several minutes at room temperature, then mount in a synthetic neutral mountant (e.g. DPX).

It is characteristic of C-bands that they are heteromorphic, varying in size between homologues in the same individual, and between different individuals of the same species (14). Such variations may characterize inbred strains, as in mice (18), or distinct races of a species, as in certain grasshoppers (19). C-banded material appears to be selectively neutral in most species (and is sometimes regarded as junk), and in humans, no convincing evidence has been produced to associate heteromorphisms with clinical conditions such as infertility, congenital abnormality, and mental retardation. A C-banded mammalian metaphase spread is illustrated in Figure 1.

Figure 1. A C-banded human lymphocyte metaphase. Note the large blocks of heterochromatin on chromosomes 1, 9,16, and Y.

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4.3 G-banding G-banding methods are the standard procedures for investigating the karyotypes of higher vertebrates, and are in routine use in clinical cytogenetics laboratories throughout the world. The ASG method given here (21) is one of the standard methods for G-banding, and tends to give better chromosome morphology than the somewhat more widely used trypsin methods.

Protocol 7. ASG method for G-banding (21) Reagents « 2 x SSC at 60°C (see Protocol

6)

• Giemsa (see Protocol SI

Method 1. Allow the chromosome preparations to age for about a week (acceptable results can probably be obtained between 3-14 days, or sometimes much longer). 2. Incubate slides in 2 x SSC at 60°C for 1 h. 3. Rinse with distilled water. 4. Stain for 45 min. 5. Rinse thoroughly with distilled water. 6. Blot carefully, allow to dry thoroughly, and mount in a neutral mountant (e.g. DPX).

A human metaphase and karyotype banded by this procedure are illustrated in Figure 2.

4.4 Ag-NOR staining for nucleolus organizing regions The sites of nucleolar organizers on chromosomes can be stained specifically with silver (Ag-NOR staining). Ag-NOR staining is used to locate NORs on chromosomes of both animals and plants. Ag-NOR staining actually reflects NOR activity at the preceding interphase, so that not all sites of genes for ribosomal RNA are necessarily stained (14). Ag-NOR staining can be used to study changes in ribosomal gene activity during embryonic development, during gametogenesis, and in cells with different genotypes and metabolic activities.

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Figure 2. Human lymphocyte metaphase chromosomes (top) and karyotype (bottom), G-banded using the ASG technique. Reprinted with permission from Nature New Biology, 232, pp. 31-32, Copyright© 1971, Macmillan Magazines Ltd.

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A. T. Sumner and A. R. Leitch Protocol 8. Ag-NOR staining (22) Equipment and reagents • Hotplate at approx. 70°C • Colloidal developer: dissolve 2 g gelatin in . Silver nitrate: dissolve 4 g AgN03 in 8 ml 100 ml distilled water with gentle warming distilled water—keep the solution in the and continuous stirring; when the gelatin dark, and discard it if any blackening occurs has dissolved, add 1 ml pure formic acid . Giemsa: 5% Gurr's Improved R66 (BDH) in (discard this soulution after about two buffer pH 6.8, made with Gurr's buffer weeks) tablets (BDH)

Method 1. Slides are best used two to three days after the chromosomes have been spread. 2. Mix two drops of colloidal developer and four drops of silver nitrate solution in an Eppendorf tube. Pipette the mixture on to the chromosome preparation, and cover with a coverslip. 3. Place the slide on a hotplate pre-heated to approx. 70°C. The slide should be removed when the solution turns golden yellow, after 1-2 min. 4. Remove the slide from the hotplate, and wash the coverslip off with a stream of distilled water. Wash thoroughly with distilled water. 5. Counterstain with Giemsa for about 5 min. 6. Rinse with distilled water, blot, allow to dry thoroughly, and mount with a neutral mountant (e.g. DPX).

Chromosomes showing Ag-NOR staining are illustrated in Figure 3.

4.5 CREST labelling of kinetochores This immunocytochemical method for labelling kinetochores differs from all the others described above in that it cannot be used on standard alcohol: acetic acid fixed chromosome preparations, but requires cytocentrifuge preparations that have been made with minimal fixation (Protocol 4). CREST serum is obtained from patients with a particular form of the autoimmune disease scleroderma; these vary considerably in the amount and type of anti-kinetochore antibodies that they contain, and a new sample needs to be tested on material that is known to give a good reaction. Formerly it was necessary to obtain CREST sera from a friendly hospital rheumatology department, but nowadays they are also available commercially (often referred to as anti-centromere antibody, ACA). Because this is an immunocytochemical reaction involving different preparation procedures, it is difficult to combine CREST labelling with routine banding to identify individual chromosomes, which can be a serious disadvantage of this procedure. If it is necessary to do kinetochore 166

4: Microscopy of chromosomes

Figure 3. Ag-NOR staining of CHO metaphase chromosomes. The NORs are arrowed. Note the variability in size of the NORs. Counterstained with Giemsa.

labelling on alcohol:acetic acid fixed chromosomes, the C d -banding procedure of Eiberg (23) could be tried, or alternatively, in situ hybridization with centromeric DNA sequences, such as alphoid DNA in humans (24). Protocol 9. CREST labelling of kinetochores Equipment and reagents • Moist chamber: place paper towels on the bottom of a plastic sandwich box, and soak them with PBS, Add glass or plastic rods to keep the slides above the wet towels; note that the slides must be able to lie horizontally to avoid the reagents running away. « CREST serum, obtainable from hospital rheumatology departments, or commercially (The Binding Site) . Pure methanol at -20"C

. P B S (see Protocol 4} . Bovine serum albumin (BSA) • Anti-human IgG, conjugated with FITC (fIuorescein isothiocyanate) • Propidium iodide solution (approx. 1 mg/ml} or DAPI (2 mg/ml) in PBS • Antifadant mountant (e.g. Citifluor API, Citifluor Ltd., or Vectashield, Vector Laboratories I

Method 1. Fix cytocentrifuge preparations of chromosomes for 10 min in methanol at-20°C. 2. Allow the slides to warm to room temperature and dry. 3. Incubate the chromosome preparations for 30 min in a moist chamber 167

A, T. Sumner and A. R. Leitch Protocol 9.

4. 5. 6. 7. 8. 9.

Continued

at room temperature in CREST serum diluted with PBS containing 1% 8SA. The optimal dilution for each serum must be found by experience, but try 1:500 to start with. Put a drop of the diluted serum over the area of the slides with the chromosomes and cover with a coverslip (about 40 ul of the diluted serum for a 22 mm square coverslip). Rinse the slides in three lots of PBS containing 1% BSA, Incubate for 30 min in the moist chamber in anti-human IgG, conjugated with FITC, diluted 1:5 with PBS containing 1% BSA. Rinse the slides in three lots of PBS containing 1% BSA. Counterstain with propidiurn iodide or DAPI. Rinse the slides with PBS. Mount with antifadant mountant, and seal the edges of the coverslip with rubber solution.

A metaphase spread with the kinelochores labelled with CREST serum is shown in Figure 4.

5. In situ hybridization DNA:DNA in situ hybridization, which involves hybridizing DNA probes directly to chromosomes (Figure 5), is an important method that has been put to many uses in genetic studies. These include: (a) Mapping sequences to chromosomes, which is now a major tool in genome mapping projects.

Figure 4. Imrrmnofluorescent labelling of kinetochores of CHO chromosomes with CREST serum, followed by FITC labelled anti-human IgG. The kinetochores appear as pairs of bright dots at the centromeres. Counterstained with ethidium bromide. 168

Figure 5. In situ hybridization of a rye dispersed sequence AF1/4 (25) labelled with digoxigenin-11-dUTP to a root tip metaphase of a Triticum aestivum (wheat) x Secale cereale (rye) hybrid, Triticale cv TC400. (A) DAPI staining for DNA (blue fluorescence). The rye chromosomes have bright DAPI positive subtelomeric heterochromatin, which become visible because of the denaturation step. Subtelomeric heterochromatic bands are major C-bands in rye. (B) Sites of probe hybridization were detected using anti-digoxygeninFITC (yellow/green fluorescence). Probe hybridization is restricted to the euchromatic regions of the rye chromosomes, (C) Double exposure for DAPI and FITC fluorescence. Note that the rye chromosomes show label (cyan fluorescence) only on the euchromatic regions of the chromosome; the subtelomeric heteroctiromatin is unlabelled but fluoresces brightly with DAPI.

A. T. Sumner and A. R. Leitch (b) (c) (d) (e) (f)

Determining genome structure. Understanding genome evolution and ancestry. Analysing interphase nuclei. Assessing sequence copy number. Examining DNA condensation (11, 26-28).

The method enables sequences on more than one chromosome to be mapped at metaphase. The resolution of mapping is about 1 Mb, while on pachytene bivalents it is thought to be a few hundred kilobases. Less than 1 kb has been detected in situ, and recently fibre spreading methods have enabled sequences separated by a few kilobases to be resolved (29). Furthermore several sequences can be detected simultaneously by using more than one probe label. Using ratio or spectral imaging all human chromosomes can now be simultaneously painted in a single preparation (30).

5.1 Probe preparation There are three main sources of DNA for use as probes for in situ hybridization. (a) Total genomic DNA. These probes have been used to perform genomic in situ hybridization (GISH) experiments to identify and localize alien chromosomes and chromosome sections introduced into crop plants (28) and to determine the ancestry of allopolyploid plants (27). (b) PCR (polymerase chain reaction) generated probes. PCR probes have been used to make chromosome paints (31), to amplify specific DNA sequences (32), and to elongate sequences for in situ localization (33). (c) Cloned sequences. These are still most commonly used and the vast majority of sequences that are localized by in situ hybridization are first cloned into plasmids, although cosmids, bacterial artificial chromosomes (Bacs), and yeast artificial chromosomes (Yacs) are increasingly used. The larger the cloned DNA fragment the more important it is to include blocking DNA in the probe mixture. The blocking DNA allows in situ suppression of repeats in the probe and thus prevents in situ hybridization signal from labelled repeats occurring across the whole genome (34). The method is known as chromosomal in situ suppression. The most commonly used DNA labelling methods are designed to incorporate nucleotides conjugated to biotin, digoxigenin, or fluorochromes directly into the DNA. Sometimes, with elegant ratio labelling experiments, two different labels are incorporated into the same probe at known ratios. By varying the ratio between probes, and with electronic imaging methods (including spectral karyotyping), just two or three labels can uniquely identify more than 20 probes simultaneously (30). 170

4: Microscopy of chromosomes Biotin is bound to nucleotides with a linker arm to minimize steric hindrance of probe hybridization. The most commonly used biotin derivative is biotin-11-dUTP, but other modified nucleotides are available. Biotin has a strong affinity to avidin which, when conjugated to a fluorochrome, is used to detect the probe hybridization sites. Digoxigenin is a natural steroid occurring in the plants Digitalis purpura and D. lanata. The derivative, digoxigenin-11dUTP, is incorporated into DNA to make the probe. Antibodies against digoxigenin, typically conjugated to fluorochromes, are used to detect probe hybridization sites. Three fluorochromes conjugated directly to nucleotides are commercially available (Amersham), which are listed in order of decreasing sensitivity: fluorescein (fluoresces yellow/green), rhodamine (fluoresces red), and coumarin (fluoresces blue). They are useful because after in situ hybridization and stringent washing the probe hybridization sites can be viewed directly and no detection steps are necessary. Thus there are advantages of speed, they can be used easily in multiple labelling experiments, and there is little background. However, the sensitivity of the label is less than either biotin or digoxigenin (particularly the rhodamine and coumarin fluorochrome conjugates) and this has restricted the use of these labels to the localization of highly reiterated sequences. For this reason, and to explain the extra steps for biotin and digoxigenin the protocols are restricted to these labels. However fluorochrome conjugates are worth using where possible. Fluorochrome-11-dUTP conjugates are incorporated into the probe in the same way as for digoxigenin-11-dUTP. 5.1.1 Probe labelling There are many methods to incorporate labels into the DNA probe. The one described here is nick translation (Protocol 10) which employs two enzymes, DNase I and E. coli DNA polymerase, that incorporate labelled nucleotides along both strands of the DNA probe by nicking the DNA and substituting nucleotides. Commercially available enzyme mixtures have been optimized to produce > 50% incorporation of label into DNA in about 90 minutes. In addition they generate probe lengths of about 200-400 bp which is ideal for in situ hybridization. Increasingly, polymerase chain reaction labelling of the probes is used, and during the amplification cycles, label is incorporated directly into the probe by adding labelled nucleotides to the reaction mixture. If PCR generated probes are more than 1 kb long there may be probe penetration problems during the in situ reaction and signal strength reduced. Protocol 10. Labelling of DNA by nick translation Equipment and reagents • Water-bath at 15°C • 100 mM dithiothreitol . 0.3 M EDTA pH 8

• Biotin labelled nucleotide: 0.4 mM biotin11-dUTP (Sigma) made up from powder in 100 mM Tris-HCI pH 7.5

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Continued

. • DNA for labelling: optimal concentration of DNA polymerase l/DNase I: 0.4 U/u1 (Gibco DNA for labelling is 0.5-1 ug/ml; ideal DNA BRL)—do not leave at room temperature; it is active at 15°C and is damaged at higher length is greater than 500 bp . 10 x nick translation buffer: 0.5 M Tris-HCI temperatures . Digoxigenin labelled nucleotide mixture: pH 7.8, 0.05 M MgCI2, 0.5 mg/ml (w/v) mix digoxigenin-11-dUTP (1 mM stock bovine serum albumin, nuclease-free solution; Boehringer Mannheim) and dTTP • Unlabelled nucleotide mixture: make 0.5 (1 mM stock) to a final concentration of 0.35 mM solutions of each nucleotide (dCTP, mM digoxigenin-11-dUTP and 0.65 mM dGTP, and dATP; Sigma) in 100 mM dTTP Tris-HCI pH 7.5 and prepare a 1:1:1 mixture . 1 x TE: dilute 100 x TE (1 M Tris-HCI pH 8,> 3 M sodium acetate . 70% and 100% ethanol (ice-cold) 0.1 M EDTA) with water

Method 1. In a 1.5 ml Eppendorf tube prepare the nick translation solution as follows: 5 ul 10 x nick translation buffer, 5 ul unlabelled nucleotide mixture, either 1 ul digoxigenin labelled nucleotide mixture, or 2.5 ul biotin labelled nucleotide, 1 ul 100 mM dithiothreitol, a quantity of solution containing 1 ug DNA, water to make up the total volume to 45 ul. 2. Add 5 (ul DNA polymerase l/DNase I solution, mix gently, and centrifuge briefly. 3. Incubate at 15°C for 90 min. 4. Stop the reaction by adding 5 ul 0.3 M EDTA pH 8. 5. Precipitate the DNA by adding 5 ul 3 M sodium acetate (or 5 ul 4 M LiCI) and 150 ul ice-cold ethanol. Place the DNA in the freezer (-20°C) overnight or on dry ice for 1-2 h. 6. Centrifuge the tubes at-10°C for 30 min at 12000 g. 7. Discard the supernatant by inverting the Eppendorf tube on to fresh tissue paper in a single action. 8. Wash the pellet by adding 0.5 ml ice-cold 70% ethanol and then spin for 5 min. Tip off solution as in step 7. 9. Leave the pellet to dry (several hours at room temperature, less under vacuum). 10. Resuspend the DNA in 1 X TE (typically 10 ul to give an estimated concentration of 0.1

5.1.2 Checking label incorporation After DNA labelling it is advisable to check label incorporation either by using a dot blot (see Protocol 11) or by running a gel and observing retardation of mobility in labelled samples. The latter is particularly useful if direct label fluorochromes are used in PCR labelling reactions. 172

4: Microscopy of chromosomes Protocol 11. Checking incorporation of the probe Equipment and reagents • Hybond N* (Amersham) • • Autoclave bags . Buffer 1: 0.1 M Tris-HCI pH 7.5, 0.15 M NaCI .Buffer 2: 0.5% (w/v) blocking reagent • (Boehringer Mannheim) in buffer 1, dissolved by heating the solution to 50-70°C for at least 1 h—the solution can be stored at 4°C for up to one month • . Buffer 3: 0.1 M Tris-HCI pH 9.5, 0.1 M NaCI, 0.05 M MgCI2

Detection of biotin incorporation: 1:500 dilution of alkaline phosphatase-avidin D (Vector Laboratories) in buffer 1 Detection of digoxigenin: 1:5000 dilution of anti-digoxigemn conjugated to alkaline phosphatase (Boehringer Mannheim) in buffer 1 NBT (4-nitroblue tetrazolium chloride)/BCIP <5-bromo-4-chloro-2-indolyl-phosphate) stable mixture (Gibco BRL)

Method 1. Cut the Hybond N+ membrane to the size required. 2. Soak the membrane in buffer 1 for 5 min and then blot dry between filter paper. 3. Load the DNA on to the membrane (1 ul) and leave to dry for 5 min. 4. Place the membrane in buffer 1 for 1 min and then into buffer 2 for 30 min. Shake gently during this period. 5. Drain the membrane and place in a Petri dish. 6. Add appropriate alkaline phosphatase conjugate in buffer 1 to the membrane, cover with plastic cut from an autoclave bag, and incubate at 37 °C for 30 min. 7. Wash the membrane in buffer 1 three times for 5 min. 8. Transfer the membrane to buffer 3 for 2 min. 9. Add NBT/BCIP mixture, apply fresh plastic cover, and incubate in the dark for 10 min for the colour to develop fully. Wash the membrane to stop the reaction in water and air dry. 10. Select only those probes that produce a good clear dot. 11. Code the probes with a strong signal so that it is easy to reference the probe to the dot blot. Discard weakly labelled probes.

5.2 In situ hybridization reaction The in situ hybridization reaction is carried out over a two day period. The reaction has a large number of steps. (a) RNase treatment. This pre-treatment removes cellular RNA so that the probe can only bind to cellular DNA, most of which is chromosomal. (b) Pepsin treatment. This pre-treatment is optional and should be omitted if possible. It increases the sensitivity of in situ hybridization, and is useful, 173

A. T. Sumner and A. R. Leitch sometimes essential, in detecting low copy sequences. However, chromosomes become more vulnerable to damage during denaturation steps and the treatment may add unnecessary steps to the protocol. (c) Denaturation. Denaturation of the chromosomal DNA is a critical step which should be monitored carefully. The denaturation solution contains formamide which destabilizes DNA and reduces the melting temperature of the DNA by about 0.6 X the percentage of formamide in the solution. In contrast, the concentration of cations (Na+) increases the melting temperature by about 17 X log molarity of cations. Thus a balance of salts, formamide, and temperature regulate the denaturation conditions (for more information see ref. 35). Too much denaturation and the chromosomes are stripped of DNA; too little and the DNA is not denatured. The protocol describes general conditions. For new material try denaturing chromosomes at a range of temperatures, wash slides, dehydrate them through an ethanol series, and examine the condition of the chromosomes with phase or preferably Nomarski phase-contrast microscopy or following staining with DAPI. Choose the hottest solution that maintains chromosome structure. (d) Hybridization and hybridization mixture. The hybridization mixture contains probe DNA, blocking DNA, salts, formamide, dextran sulfate, and sodium lauryl sulfate (SDS). The dextran sulfate increases the hybridization reaction rate by a factor of three and works by forming a matrix in the hybridization mixture which concentrates the probe without affecting the denaturation conditions. The probe(s) are typically labelled with fluorochromes, digoxigenin, biotin, or ratios of each. The blocking DNA has different roles in different experiments and is typically used at concentrations of 2-250 times the probe concentration. In all experiments it reduces non-specific probe hybridization. With CISS and chromosome painting experiments, the blocking DNA comes from the organism under study and the blocking DNA increases the specificity of probe hybridization. With GISH experiments, the blocking DNA is obtained from one of the parents in the hybrid organism being studied and once again increases the specificity of probe hybridization (31, 34). Probe and blocking DNA need to be denatured in the hybridization mix by heating, and applied to the material for in situ hybridization overnight at 37 °C. (e) Stringent washing. This wash is designed to strip away weakly bound and non-specifically bound probe which may occur during the overnight in situ hybridization reaction. The solution contains salts and formamide at concentrations that destabilize weakly bound probes leaving only strongly bound probe hybridized. The protocol leaves a stable sequence identity of greater than 80-85%. Increasing the temperature of this wash can further reduce background and non-specific labelling but the strength of desirable signal may also decline. 174

4: Microscopy of chromosomes (f) Detection. In double label experiments using biotin and digoxigenin the biotin is detected with avidin conjugated to one fluorochrome (here Cy3) while digoxigenin is detected with an antibody conjugated to another fluorochrome that fluoresces at a different wavelength (here FITC). The choice of fluorochromes arises because Cy3 is only obtained conjugated to avidin and it is a particularly good fluorochrome. (g) Counterstain. Use only DAPI fluorescence for a chromosomal counterstain unless there are compelling reasons to use propidium iodide. Propidium iodide results in a great loss of information due to fluorescence across a wide spectrum. (h) Antifadant. This increases the stability of the fluorochromes, e.g. FITC, Cy3, and DAPI which is essential for photography. Protocol 12. In situ hybridization Equipment and reagents • Humid chamber. Place 2 x SSC soaked tissues in a floating box with a lid. Put into the box two glass rods on to which the slides will be placed; the rods will keep the slides above the wet tissues. Float the box in a water-bath at 37°C (or place in a 37°C room) and allow the box to equilibrate at the temperature. • Plastic coverslips: made from plastic autoclave bags cut to an appropriate size to cover the material, but not to overlap the edge of the slide . Water-baths at 37°C and 70°C • 20 x SSC pH 7: 3 M NaCI, 0.3 M sodium citrate . 2 x SSC: dilute 20 x SSC with water (1:9) • RNase A: prepare a stock solution by dissolving 10 mg/ml of DNase-free RNase in 10 mM Tris-HCI pH 7.5, 15 mM NaCI. Boil for 15 min and allow to cool. Store frozen in aliquots. For use dilute to 100 ug/ml RNase A in 2 x SSC. • 0.01 M HCI, optional step • 1 ug/ml pepsin in 0.01 M HCI: 3200-4500 U/mg protein (porcine stomach mucosa) (Sigma) • Pre-hybridization fix: ethanol, glacial acetic acid (3:1), freshly prepared . Ethanol series: 70%, 90%, 100% ethanol • Ice-cold ethanol series: 70%, 90%, 100% ethanol stored on ice • Formamide denaturation solution: heat 70% (v/v) formamide in 2 x SSC (70 ml formamide, 10 ml 20 x SSC, 20 ml water) to exactly 65°C (solution temperature) in a water-bath—prepare immediately before

• Formamide: deionized, high grade formamide is used, stored frozen at -20°C (if all the formamide does not freeze, buy fresh formamide) • 50% (w/v) dextran sulfate (M, 500000): dissolve in water at 37°C overnight and store in aliquots at -20°C—warm to 37°C to reduce viscosity before use • Blocking DNA: store at -20°C, defrost and vortex well before use. (a) For mapping short cloned sequences: salmon total genomic DNA (autoclaved, 10 min). (b) For GISH experiments: total genomic DNA (autoclaved, 10 min) from the organism under study, (c) For chromosome painting and CISS experiments: total genomic DNA (autoclaved, 10 min) from the organism under study. • Probe DNA-dig: digoxigenin labelled DNA (0.1 ug/ml) stored in freezer, defrost and vortex well before use • Probe DNA-bio: biotin labelled DNA (0.1 ug/ml) stored in freezer, defrost and vortex well before use • 10% (w/v) SDS in water • Hybridization mix (see Table 1) • Stringent wash: 20% formamide in 0.1 x SSC (20 ml formamide, 0.5 ml 20 x SSC, 79.5 ml water) • 4 x SSC/Tween: dilute 4 parts 20 x SSC with 16 parts water and add 0.2% (v/v) Tween 20 • Bovine serum albumin (BSA) . BSA block: 5% (w/v) BSA in 4 x SSC/Tween 20 • Detection reagent: prepare a mixture of 10 (ug/ml anti-digoxigenin-FITC and 5 avidin-Cy3 in BSA block

175

A. T. Sumner and A. R. Leitch Protocol 12. Continued • DAPI: stock solution (100 (ig/ml DAPI in water), stored in aliquots at -20°C—dilute stock solution to 2 ug/ml DAPI in 2 x SSC • Anti-digoxigenin-FITC (Boehringer Mannheim) (digoxigenin detection reagent): make stock solutions of 500 ug/ml in sterile distilled water according to manufacturer's instructions, and store aliquoted in freezer (freeze solution once only)

.

.

Cy3-avidin (Amersham) (biotin detection reagent): supplied at stock concentration (1 mg/ml)—store aliquoted in freezer (freeze solution once only) Antifadant: Vectashield antifadant reduces bleaching of fluorochromes; also available with DAPI added (Vector Laboratories)

A. Day 7: slide pre-treatments and hybridization steps 1. Place slides in an oven at 37°C overnight. 2. Add 200 ul RNase A, cover with a plastic coverslip, remove air bubbles, and incubate for 1 h at 37°C in a humid chamber. 3. Wash slides in 2 x SSC, three times for 5 min each. 4. If pepsin is used, place material in 0.01 M HCI for 2 min. 5. Optional step. Add 200 ul pepsin solution, cover with a plastic coverslip, and incubate for 10 min at 37°C. 6. If pepsin is used, stop reaction by placing in water for 2 min, and wash in 2 x SSC, twice for 5 min each. 7. Place material in pre-hybridization fix for 10 min. 8. Dehydrate for 3 min each in 70%, 90%, and 100% ethanol, and then air dry. 9. Prepare the hybridization mix as shown in Table 1 prior to denaturing slides. 10. Denature the hybridization mix at 70°C for 10 min, vortex briefly, and place on ice. 11. Denature slides. Place slides in formamide denaturation solution at 65°C for 3 min. The temperature needs to be determined empirically for your material (see text). This is one of the most crucial steps. When using a standard Coplin jar, each slide causes a reduction in the temperature of the formamide solution of about 2°C. Therefore denature slides one at a time and note the solution temperature change. 12. Place slides in ice-cold ethanol series, 3 min in each, and air dry. 13. Add 40 ul of the freshly denatured hybridization solution and cover with a plastic coverslip. Ensure there are no air bubbles. 14. Incubate overnight at 37°C in the humid chamber. B. Day 2: post-hybridization washing and label detection 1. Float coverslips off in 2 x SSC at 35-42°C. 2. Place slides in stringent wash, twice for 5 min each, at 42°C.

176

4: Microscopy of chromosomes 3. Wash slides in 2 x SSC at 42°C, three times for 3 min each. 4. Take Coplin jar out of water-bath and leave to cool for 5 min. 5. Wash slides in 2 x SSC three times for 3 min each. 6. Place slides in 4 x SSC/Tween for 5 min. 7. Drain but do not allow slide to dry. Add 200 ul BSA block to each slide and apply a plastic coverslip, incubate for 5 min. 8. Remove coverslip and drain but do not allow slide to dry. Add 30 ul detection reagent per slide. Replace coverslip and incubate for 1 h at 37°C in a humid chamber. 9. Wash slides in 4 x SSC/Tween at 37°C, three times for 8 min each. 10. Drain solution, but do not allow slide to dry. Add 100 ul DAPI per slide to counterstain the chromosomes, cover with plastic coverslip, and incubate for 10 min. 11. Rinse briefly in 4 x SSC/Tween. 12. Drain but do not allow slide to dry. Apply antifadant solution. 13. Place a thin coverslip (preferably UV transparent and of high quality) over material. Gently squeeze excess antifadant from the slide with filter paper.

Table 1. Composition of hybridization mixture Solution

To prepare 40 ul

Final concentration

Formamide 50% (w/v) dextran sulfate 20 x SSC Probe DNA-dig (0.1 p-g/pl) Probe DNA-bio (0.1 ug/Ml) Blocking DNA (1 ug/ul) 10%(w/v)SDS Water

20 Ml

50% 10% 2X 2.5ng/ul 2.5 ng/ul 250 ng/ul 1.25%

8 ul 4Ml 1 ul(100 ng/slide) 1 ul(100 ng/slide 2 Ml (2 ug/slide) 0.5 Ml 3.5 ul

Use 40 (ul per slide. Adjust ratios appropriately for different volumes.

6. Observation and recording of images of chromosomes Many of the features that can be demonstrated on chromosomes are at or near the limit of resolution of light microscopy. A good quality microscope, equipped with high quality lenses and properly adjusted, is therefore necessary for observing chromosomes. A high quality camera is also required, and increasingly, confocal microscopes and CCD cameras are being used; the 177

A. T. Sumner and A. R. Leitch latter are particularly useful for fluorescence studies, particularly in in situ hybridization, as they can be used to obtain multicoloured images at high resolution and high sensitivity (as well as high price!). Practical details of how to set up a microscope to obtain optimal results have been given elsewhere (36) (also Chapter 1). Requirements for observation of chromosomes stained with absorbing dyes, and those stained with fluorochromes are, in many ways, rather different. However, one important factor for the study of chromosomes is the use of flat field objectives. These are not only necessary for photography and other image recording systems, but are also valuable for direct observations. It is excessively tiresome if the focus has to adjusted continually to observe chromosomes in different parts of the field and comparison between different chromosomes becomes much more difficult if only one is in-focus at a time.

6.1 Observation of banding with absorbing dyes A low power (x 10) objective is necessary to locate metaphase spreads on the slide, where they may be quite rare compared with the interphase nuclei, but this magnification is totally inadequate to see banding. In fact, a X 90 or X 100 oil immersion objective is necessary to assess the quality of the preparation, and to make definitive observations. The contrast of banded chromosomes can be improved by optical means, either by the use of phasecontrast or differential interference (Nomarski) microscopy, or by the use of filters. Phase-contrast has been used for improving the visibility of Giemsa stained R-bands, which are sometimes rather pale, and can also be used with Ag-NOR staining if the chromosomes have not been counterstained. In the latter case, the Ag-NORs appear as bright objects against a darker background. Green filters can be used to increase the visibility and contrast of Giemsa stained chromosomes.

6.2 Observation of fluorescent chromosomes Fluorescently stained chromosomes generally have a low level of fluorescence, and therefore a fluorescence microscope of the highest quality is needed. The characteristics of such a system are described in detail elsewhere (37) (Chapter 6). The important features are the greatest possible efficiency of illumination (ideally a 100 W mercury lamp), and of collection of the emitted fluorescence. These are achieved by using epi-illumination (incident illumination through the objectives), and objectives of the highest possible light transmission, i.e. high numerical aperture, flat field, fluorescence objectives. These are available from the major microscope manufacturers especially for fluorescence work. It is worth purchasing oil immersion objectives of both approximately X 50 and X 100 magnification, as fluorescence microscopy requires continual changing between these two magnifications, the lower being used for detailed scanning of the slide, and the higher for observation. 178

4: Microscopy of chromosomes Fluorescence microscopy requires the use of exciter filters, to ensure that only light of the required wavelengths reaches the specimen to excite fluorescence, and barrier filters, to cut out the exciting light and to transmit only the fluorescence. With epi-illumination, these filters are combined in a single module with a dichroic reflector which enables the epifluorescence system to work by reflecting short wavelengths and transmitting long wavelengths. It is vital to use the correct filter combinations for the fluorescence to be studied (Table 2). Unfortunately, there is no universal system for designating filter modules for fluorescence, and each manufacturer has its own system. It is always advisable to try out any objectives and filter modules on the types of specimen you are interested in, to see which gives the best results, before spending any money. A serious problem in all fluorescence microscopy is fading of fluorescence during illumination. Special mountants that retard the fading of fluorescence are now available from various sources (Citifluor; Vector), and the use of Table 2. Useful fluorochromes for use in cytogenetics" Excitation

(max, nm)

Emission (max, nm)

Fluorescence colour

615

Red

Microscope fittersb

Fluorochromes that stains DNA Propidium iodide

340, 530

BP 525-560

CBS 580 DAPI

355

450

Blue

LP590 BP 340-380

CBS 420 Chromomycin A3

430

570

Yellow

LP420 BP436

CBS 455 LP470 Fluorochromes conjugated to probe detection systems Fluorescein (FITC)

495

Cy3

550

515

Green

BP 450-490

CBS 510 570

Orange

LP520 BP515-560C

CBS 580 Coumarin (AMCA)

350

450

Blue

LP590 BP 340-380

CBS 420 LP420

aThe microscope filters are for visualizing single fluorochromes. Double and triple bandpass filters enable combinations of fluorochromes to be visualized simultaneously and can be obtained from leading microscope suppliers. b BP = bandpass filter, light above or between wavelength given is transmitted. CBS = chromatic beam splitter, light above wavelength given is transmitted, below it is reflected. LP = long pass filter, light above wavelength given is transmitted. c Use rhodamine filter block.

179

A. T. Sumner and A. R. Leitch some of these has been described in some of the protocols given above. Two points should be made about such mountants. First, although they do retard the rate of fading (but do not usually stop it entirely), they may also reduce the brightness of the fluorescence, which may limit their value. Secondly, there is no universal antifadant, so that what works well with one fluorochrome may be useless with another. Some fluorochromes, such as quinacrine, cannot be stabilized by any antifadant; on the other hand, it has been found that storage overnight or for a few days before observation greatly stabilizes the fluorescence, as with DAPI and chromomycin.

6.3 Photography of chromosomes Photography of chromosomes provides a permanent record, not only for publication and to provide a reference in clinical studies, but also for detailed analysis of banding and in situ hybridization patterns. General principles for photographing banded chromosomes have been described by Davidson (38). The first requirement, of course, is that the microscope should be properly set up (36) (Chapter 1). It is necessary to take a test strip both when using new equipment and when trying out a new type of film or developer. Exposures should be varied on either side of those recommended by the manufacturer of the film, or prescribed by the automatic exposure meter. The effect of various filters should also be tested. All relevant data must be recorded, so that the optimal exposure can be used in the future. 6.3.1 Black and white photography For photographing chromosomes stained with absorbing dyes, a slow black and white film such as Kodak Technical Pan or Ilford Pan F is quite adequate, as there is no problem with the staining being bleached, and these slow films produce the finest grain. For photographing fluorescence, the fastest possible film is desirable, because of the problem of fading, and the low light levels involved. Formerly it was impossible to obtain a reasonable film speed (400 ASA or more) without excessively coarse grain, but nowadays such films not only have a reasonably fine grain, but can also be rated at a higher speed using suitable development—consult the manufacturers' instructions on this point. For fluorescence work it is essential to have a light meter that can select a specific area of the specimen, i.e. part of a fluorescent chromosome, as it is impossible to calculate the correct exposure from an area consisting largely of black background. A particular problem with photographing fluorescence is reciprocity failure: in effect, the longer the exposure, the slower the effective film speed. For photographing very dim fluorescence, therefore, it may be necessary to rate the film at half, or even a quarter, of its usual speed. 6.3.2 Colour photography Colour photography can be very valuable for fluorescence work, particularly where two or more different colours have to be distinguished, as in double 180

4: Microscopy of chromosomes labelling in situ hybridization. Here again, films have been greatly improved in recent years, and it is possible to get high speed colour films without excessively coarse grain. The recording of images by colour photography offers many advantages over black and white photography: (a) Multiple fluorochrome labels can be distinguished. (b) Dark-room time is eliminated. (c) Film processing can be done quickly. For epifluorescence microscopy it is necessary to have a high speed, fine grain film with high colour contrast. The films we use are Fujicolor 400 and Fujichrome 400. Slide films (Fujichrome 400) give the most accurate representation of colour, but this may not be as important as clear colour distinction between fluorochromes. They also have smaller grain sizes than print films and can be processed anywhere, even in the laboratory. The main disadvantage is that slides are difficult to analyse. Print films (Fujicolor 400) are convenient, as several copies of prints can be made as a matter of routine and prints are easily analysed. In addition, when using print films there is more flexibility with exposure times because simple adjustments during printing can generate acceptable images from negatives with widely different densities. However, printing can be a problem as a photographer needs to be found who has the facility to print the exposure times and filter settings on the back of each print. Most towns have such photographers. In consultation with the photographer the optimal settings and ranges are established for printing. In a short time it is then only necessary to drop off the films and pick up the prints. But setting up the necessary dialogue with the photographer takes time.

6.4 Other methods of image capture Increasingly, electronic images are being captured to record images, in particular those from fluorescent in situ hybridization experiments. This is due to an increasing use of confocal microscopes, low light cameras, and image enhancement. In addition, images recorded on photographic film are being scanned into an electronic image for printing with high contrast on colour printers. 6.4.1 Confocal microscopes The confocal microscope offers advantages in that it can be used to obtain high resolution images, and if a sufficiently small aperture is used, much of the out-of-focus information can be removed. The microscope scans the specimen with a spot of laser light at an appropriate wavelength to excite fluorochromes. Fluorescent light passing through the confocal aperture is recorded with a photomultiplier and displayed point by point on to an appropriate high 181

A. T. Sumner and A. R. Leitch resolution monitor. Confocal microscopy is used routinely by many laboratories and the technique offers certain advantages. (a) The brief period of fluorochrome excitation required for producing a confocal image (usually about 2 sec) reduces the bleaching of fluorochromes. (b) The digital images recorded by the confocal microscope can be stored and recombined enabling easy alignment of different images. (c) Instant images are obtained, whereas conventional microscopy requires film processing, which can take days. (d) The resolution of images may, if the instrument is used at its highest possible performance, be higher. (e) Optical sections can be taken and three-dimensional objects reconstructed. The potential increased resolution and the possibility of reconstruction from optical sections does mean that in some circumstances the confocal is extremely powerful. However there are several major disadvantages. (a) In practice a confocal microscope slows down analysis of slides and fewer cells will be examined. (b) The images recorded can be subjected to too much manipulation which can distort or lose information. (c) The microscopes are very expensive to buy and run. For further information see Chapter 6, Section 7.3. 6.4.2 Low light (CCD) cameras These cameras record fluorescent images electronically (see Chapter 6, Section 7.4). They have one major advantage over conventional cameras: the acquisition of instant images. However they have all the disadvantages of a confocal microscope above (except running costs). In theory, fluorescence not visible to the eye can be recorded and imaged, although in our experience, this ability is of little use. Background and non-specific signals tend to be imaged as well as the real signal, so that the latter is hard to distinguish. Thus imaging is restricted to material where the signal is visible. An instant image can be obtained and processed, but there are the dangers of image distortion or misinterpretation. If obtaining an instant image is the desired feature, then be careful to purchase a suitable camera. Do not compare an electronic image with what can be seen down the eyepiece of the microscope, because signal brightness and image size make this difficult. Compare a print of an image taken from a low light camera with a print from a conventional camera. Check resolution, pixel size, chromosome, and signal clarity. Do not take too much notice of the brightness of colours, which can, if required, be changed on a print either photographically or electronically. Be careful to ensure that instant images are really needed, and then be careful which camera is purchased. 182

4: Microscopy of chromosomes 6.4.3 Image processing All electronic images can be processed and printed at high quality. In addition, images obtained by conventional microscopy can be stored electronically, processed, and imaged. Equipment for doing this is now widely available. In all cases published electronic images are brilliant in intensity and colour, precise, and usually impressive. However, image processing is one of the most dangerous tools available to cytogeneticists. It requires scrupulous adherence to ethical manipulation approaches as almost anything can be 'created' or 'destroyed' using modern software. We would recommend that the only acceptable manipulation is that which affects the whole image uniformly.

References 1. Schwarzacher, T. S. and Leitch, A. R. (1994). In Methods in molecular biology (ed. P. Isaac), Vol. 28, p. 153. Humana Press, Totowa, NJ. 2. Macgregor, H. C. and Varley, J. M. (1988). Working with animal chromosomes (2nd edn). John Wiley & Sons, Chichester. 3. Barch, M. J. (ed.) (1991). The ACT cytogenetics laboratory manual (2nd edn). Raven Press, New York. 4. Bennett, M. D. and Smith, J. B. (1976). Phil. Trans. R. Soc. Land. B, 274, 227. 5. Rooney, D. E. and Czepulkowski, B. H. (ed.) (1992). Human cytogenetics: a practical approach (2nd edn). Oxford University Press, Oxford. 6. Gosden, J. R. (ed.) (1994). Chromosome analysis protocols. Humana Press, Totowa, NJ. 7. Chandley, A. C., Speed, R. M., and Ma Kun. (1994). In Chromosome analysis protocols (ed. J. R. Gosden), p. 27. Humana Press, Totowa, NJ. 8. Jeppesen, P. (1994). In Chromosome analysis protocols (ed. J. R. Gosden), p. 253. Humana Press, Totowa, NJ. 9. Stenman, S., Rosenqvist, M., and Ringertz, N. R. (1975). Exp. Cell Res., 90, 87. 10. Pan, W. H., Houben, A., and Schlegel, R. (1993). Genome, 36, 387. 11. Leitch, I. J. and Heslop-Harrison, J. S. (1993). Genome, 36, 517. 12. Spurbeck, J. L., Zinsmeister, A. R., Meyer, K. J., and Jalal, S. M. (1996). Am. J. Med. Genet., 61, 387. 13. Worton, R. G. and Duff, C. (1979). In Methods in enzymology (ed. W. B. Jakoby and I. H. Pastan), Vol. 58, p. 322. Academic Press, New York. 14. Sumner, A. T. (1990). Chromosome banding. Unwin Hyman, London. 15. Bickmore, W. and Sumner, A. T. (1989). Trends Genet., 5, 144. 16. Holmquist, G. P. (1989). J. Mol. Evol., 28, 469. 17. Schweizer, D. and Ambros, P. F. (1994). In Chromosome analysis protocols (ed. J. R. Gosden), p. 97. Humana Press, Totowa, NJ. 18. Davisson, M. T. (1989). In Genetic variation and strains of the laboratory mouse (ed. M. F. Lyon and A. G. Searle), p. 617. Oxford University Press, Oxford. 19. Webb, G. C., White, M. J. D., Contreras, N., and Cheney, J. (1978). Chromosoma, 67, 309. 20. Sumner, A. T. (1972). Exp. Cell Res., 75, 304. 21. Sumner, A. T., Evans, H. J., and Buckland, R. A. (1971). Nature New Biol, 232, 31. 183

A. T. Sumner and A. R. Leitch 22. Howell, W. M. and Black, D. A. (1980). Experientia, 36, 1014. 23. Eiberg, H. (1974). Nature, 248, 55. 24. Mitchell, A., Jeppesen, P., Hanratty, D., and Gosden, J. (1992). Chromosoma, 101, 333. 25. Francis, H. A., Leitch, A. R., and Koebner, R. M. D. (1995). Theor. Appl. Genet., 90, 636. 26. Ferguson-Smith, M. A. (1991). Am. J. Hum. Genet., 48, 179. 27. Leitch, I. J., Parokonny, A. S., and Bennett, M. D. (1996). In Chromosomes today (ed. N. Henriques-Gil, J. S. Parker, and M. J. Puertas), Vol. 12, p. 333. Chapman and Hall, London. 28. Schwarzacher, T., Anamthawat-Jonsson, K., Harrison, G. E., Islam, A. K. M. R., Jia, J. Z., King, I. P., et al. (1992). Theor. Appl. Genet., 84, 778. 29. Zhong, X. B., Fransz, P. F., Wennekes-Vaneden, J., Zabel, P., Vankammen, A., and De Jong, J. H. (1996). Plant Mol Biol. Rep., 14, 232. 30. Schrock, E., Dumanoir, S., Veldman, T., Schoell, B., Wienberg, J., FergusonSmith, M. A., et al. (1996). Science, 273, 494. 31. Telenius, H., Pelmear, A. H., Tunnacliffe, A., Carter, N. P., Behmel, A., Ferguson-Smith, M. A., et al. (1992). Genes Chromosomes Cancer, 4, 257. 32. Pearce, S. R., Harrison, G., Li Dongtao, Heslop-Harrison, J. S., Kumar, A., and Flavell, A. J. (1996). Mol. Gen, Genet, 250, 305. 33. Cox, A. V., Bennett, S. T., Parokonny, A. S., Kenton, A., Callimassia, M. A., and Bennett, M. D. (1993). Ann. Bot., 72, 239. 34. Lichter, P., Tang, C. C, Call, K., Hermanson, G., Evans, G. A., Housman, D., et al. (1990). Science, 247, 64. 35. Meinkoth, J. and Wahl, G. (1984). Anal. Biochem., 138, 267. 36. Bradbury, S. (1989). Introduction to the optical microscope. Oxford University Press, Oxford. 37. Ploem, J. S. and Tanke, H. J. (1987). Introduction to fluorescence microscopy. Oxford University Press, Oxford. 38. Davidson, N. R. (1973). /. Med. Genet, 10,122.

184

5 Immunohistochemistry MICHAEL G. ORMEROD and SUSANNE F. IMRIE

1. Introduction Immunohistochemistry utilizes antibodies to localize specific products in tissue sections. Briefly, a tissue section is incubated with a labelled antibody, the section is washed, and the site of reaction of the antibody is identified by visualizing the label. A variety of histochemical techniques have been used for many years to identify certain constituents of cells. These methods lack the specificity obtained from an antibody and it is this property which enables a worker to map precisely the distribution of a particular product in a tissue. Aside from its power as a tool in research, immunohistochemistry has found an increasing role in diagnostic histopathology with particular application to the diagnosis of tumours. The range of applications of the method has been advanced immeasurably by the development of technology for the production of monoclonal antibodies. This has produced a wide range of antibodies, many of which have specificities which would have been difficult, if not impossible, to obtain from conventional antisera. The first part of the chapter (Sections 2-5) give the background to the techniques. The later sections describe the procedures in detail. Section 2 briefly defines an antibody and its structure, and describes the difference between polyclonal sera and monoclonal antibodies. The possible effects of tissue processing on the antigen are considered in Section 3. A wide variety of different labels may be employed; these are described and compared in Section 4. Section 5 describes and compares in general terms the different methods used for applying antibodies to tissue sections. Section 6 contains the details of the experimental methods used with enzyme labelled antibodies. The use of controls and problem solving are discussed in Section 7 and ways of visualizing two antigens simultaneously in Section 8. Although the chapter concentrates on the use of antibodies, a similar technique can be applied using any reagent that shows a similar specificity. For example, lectins can be used to identify certain groups of carbohydrates. Most of the chapter describes techniques for staining sections cut from

Michael G. Ormerod and Susanne F. Imrie tissue. The same methods can also be applied to cytological preparations and a short section (Section 9) on staining smears is included. Section 10 contains a brief discussion on quantification and Section 11 describes a staining tray—the only special piece of equipment needed. For further reading, several books on immunohistochemistry have been published. Most of these give details of the application of the technique to particular problems (1-5). All the reagents listed in this chapter can be purchased from Sigma Ltd. unless otherwise stated.

2. Antibodies Antibodies, collectively called immunoglobulins (Igs), comprise approximately 20% of the proteins in human plasma. They are produced by plasma cells and can exist in millions of different forms. During an immune response, the foreign body (antigen) stimulates division of those plasma cells responsible for producing an Ig reactive with that particular antigen. This enables the animal to produce large numbers of specific antibodies. Any animal in a normal environment continually undergoes immune responses and its plasma will contain antibodies directed against thousands of different antigens.

2.1 Immunoglobulin structure The basic structure of an Ig is a dimer, each half containing two polypeptide chains, one called heavy (containing ~ 440 amino acids), the other light (containing ~ 220 amino acids). The chains are linked by disulfide bonds (Figure 1). The heavy chain has an invariant (constant) region, which determines the class of the Ig, and a variable region. The latter, in conjunction with the variable region on the light chain, forms the site that binds the antigen. Light chains can be of two classes, kappa or lambda. The major classes of Igs are listed in Table 1. The IgG class is further sub-

Figure 1. A simplified representation of an Ig molecule showing the antigen binding site and the sites of cleavage of papain and pepsin. — represents a disulfide bond.

186

5: Immunohistochemistry Table 1. The major classes of immunoglobulins Name

Total Ig in blood (%)

Comment

IgG IgA IgM igD

80 15 5

lgE

<1

Major antibody in immune sera. Found in sero-mucous secretions. Produced early in immune response, pentameric. Present on lymphocyte surfaces. Responsible for allergic reactions.

<1

divided (IgG1, IgG2, etc.). During an immune response, the bulk of the Igs in the plasma are IgGs. Consequently an antiserum will contain predominantly Igs of this class. Immunoglobulins are sometimes digested enzymatically to produce fragments that are still reactive with the antigen. Pepsin digests the constant part of the heavy chain below the 'hinge' region leaving a dimeric fragment called F(ab')2 (see Figure 1). Papain creates two monomeric fragments, Fab, plus the constant region of the heavy chains, Fc. Fab fragments are sometimes used in place of the whole Ig.

2.2 Polyclonal antisera To raise an antiserum, a group of animals are injected with the purified antigen together with a non-specific stimulant of the immune response (an adjuvant). A protocol can be found in ref. 6. If the immunization is successful, the antigen will have stimulated a variety of lymphocytes, each of which will have undergone several divisions to produce a clone of antibody-producing plasma cells. For this reason, the term 'polyclonal' antiserum is sometimes used. The site on an antigen with which an Ig reacts is called an epitope. An antigen may contain several epitopes and an antiserum will contain Igs directed against each of them. Furthermore different Igs directed against the same epitope may have different binding affinities and also may see a slightly different part of the epitope. An antiserum will therefore contain a variety of Igs reactive with the antigen but with different specificities and affinities. Consequently, two antisera are never identical. Apart from a high concentration of antibodies directed against the appropriate antigen, an antiserum will also contain other Igs that were present before immunization and may contain antibodies reactive with impurities in the original preparation of antigen. These Igs may give undesired reactions requiring their removal.

2.3 Monoclonal antibodies Many of these problems are avoided by the use of monoclonal antibodies. These are made by a cloned line of cells that produce a single Ig. The cells are 187

Michael G. Ormerod and Susanne F. Imrie derived by fusing spleen cells from an immunized animal with a line of drugsensitive myeloma cells. After the fusion, the cells are incubated in the presence of the drug. The only cells to grow will be hybrids resulting from a fusion of a normal spleen cell (which is drug-resistant) and a myeloma cell (which brings immortality to the hybrid). The hybrids are cloned and clones producing specific antibodies are selected and recloned. The resulting cell lines (called hybridomas) each produce indefinitely a single Ig. A full description of these methods can be found in ref. 7 or in another volume in this series, Antibodies: a practical approach (ed. D. Catty), Vol. 1.

2.4 Purific ation of antibodies Sometimes it is necessary to purify an antibody before attaching a label to it. This is comparatively easy if a monoclonal antibody produced in the supernatant of a hybridoma culture is used. An ammonium sulfate precipitation followed by exclusion chromatography would suffice. If a monoclonal antibody from an ascitic fluid or a polyclonal antiserum is used, the specific antibody should be separated from the other Igs. If the original antigen is available, the purification is best performed by affinity chromatography.

2.5 Specificity of antibody reactions An important property of an Ig is the affinity with which it binds to its antigen. The concentration needed to produce a desired end-point clearly depends on this affinity. The 'strength' of a polyclonal antiserum or a preparation of monoclonal antibodies is directly related to the concentration of antibody multiplied by its affinity constant; the higher the affinity, the greater the working dilution. For the reasons given below, antibodies of high affinity generally show greater specificity. Antibodies are used as reagents because of their high specificity. However it is important to realize that they can also give rise to non-specific reactions. These can have three causes. (a) From impurity antibodies. This only arises with a polyclonal antiserum and should be detected by the use of appropriate controls (see Section 7). It is less likely to be a problem if the antiserum can be used at a high dilution. (b) From cross-reactions. These arise if two molecules have similar, but different, structure. Some antibodies may recognize both molecules but often the undesired reaction will be of lower affinity and will only be a problem if the antibody is used at too high a concentration. This emphasizes the desirability of using antibodies of high affinity. (c) From two molecules sharing the same epitope. This possibility will not be revealed by the usual controls and would normally only be shown by a detailed immunochemical study. 188

5: Immunohistochemistry Preparations of monoclonal antibodies should not contain any impurity antibodies. However they are specific for an epitope and could give a misleading result if this epitope was found on more than one protein.

2.6 Storage of antibodies For antibodies and antisera obtained commercially, the supplier's instructions should be followed. Other reagents should be stored without first diluting them. It is advisable to aliquot them into suitable quantities and to store the aliquots frozen. Once an aliquot is thawed it should not be refrozen, as thawing and freezing will denature Igs. If an antibody is used rarely, it may be stored in amounts sufficient for one experimental run and an aliquot thawed when necessary. If this involves microlitre quantities of reagent, the aliquot should be covered with a small quantity of glycerol before freezing to prevent it freeze-drying. If a reagent is in regular use, it may be stored in larger aliquots. We have found that most antisera have a shelf-life at 4°C of at least a month, some considerably longer. For such reagents we regularly make them 0.01 % in sodium azide to prevent bacterial contamination. If reagents are ever contaminated, they should be immediately discarded.

3. Effect of tissue processing on antigens Before applying an antibody, a section of tissue must be prepared. Before deciding how to handle a tissue the effect of any processing on the antigen of interest must be considered. Conventionally, when sections are to be stained with haematoxylin and eosin, tissue is fixed, often in formalin, and embedded in paraffin wax. After sections have been cut, the wax must be removed by immersion in xylene, or a similar solvent, and brought to water through ethanol. This treatment yields sections of high quality. It also alters the proteins in a tissue so that many antigens no longer react with the appropriate antibody. In this case, alternative methods of fixation and processing may be tried but frequently sections must be cut from frozen tissue.

3.1 Choosing conditions for processing When using a new antibody, it is important that the optimum conditions for fixation and processing are determined. While these can only be established empirically, it is possible to lay down some guide-lines. A set of fixatives is chosen. We would recommend formalin, methacarn, 90% ethanol, Bouin's, chloroform:acetone, and formolxalcium followed by chloroform:acetone. These fixatives are then tested on a set of frozen sections. Several frozen sections are cut from the tissue that contains the antigen of interest and one of the sections immersed in each of the selected fixatives for 189

Michael G. Ormerod and Susanne F. Imrie 5 min. The sections are stained with the antibody using the selected method and the result read. If all the fixatives chosen appear to destroy the antigen, the experiment is repeated using more gentle fixatives (e.g. paraformaldehyde: lysine:periodate). Occasionally, the staining must be carried out without prior fixation; this will give sections with poor morphology. Recipes for various fixatives, including those mentioned above, with some indications of their use are given in Protocol 1. Protocol 1. Recipes for some common fixatives A. Fixatives generally used on tissue subsequently processed into blocks of paraffin wax 1. 10% formol:saline. 100 ml 40% formaldehyde, 9 g NaCI in 900 ml water. 2. Methacarn. 60% methanol, 30% chloroform, 10% glacial acetic acid. Tissues are usually fixed at room temperature overnight and then transferred to 70% alcohol. 3. Modified methacarn. Use inhibisol in place of chloroform. 4. Carnoy's fluid. 60 ml ethanol, 30 ml chloroform, 10 ml acetic acid. 5. Bouin's fluid. 75 ml saturated picric acid, 25 ml 40% formaldehyde, 5 ml acetic acid. 6. B5. 60 g mercuric chloride, 21 g sodium acetate trihydrate in 900 ml water. Add 100 ml 40% formaldehyde before use. When using this fixative, after sections have been cut, mercury must be removed before performing an immunohistochemical stain. Take sections to water. Immerse for 5 min in Lugol's iodine.a Wash in water. Immerse for a few seconds in 5% (w/v) sodium thiosulfate. Wash in water. B. The following are usually used on frozen sections 1. Formohcalcium. 100 ml 40% formaldehyde, 100 ml 1 M CaCI2, 800 ml water, plus a few chips of marble (or CaCO3). Store at 4°C. Fixation is followed by a further fixation in chloroform:acetone. 2. Chloroform:acetone (50:50, v/v). 5 min fixation at 4°C. Often used to fix frozen sections prior to using monoclonal antibodies to distinguish lymphocyte phenotypes. 3. Ethanohacetic acid. 95% ethanol, 5% glacial acetic acid. 1 min at 4°C for frozen sections. 4. Ethanol. Various percentages of ethanol at 4°C or room temperature for a predetermined time.

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Periodate:lysine:paraformaldehyde. (a) Prepare a 3.6% (w/v) solution of paraformaldehyde by dissolving 2 g in 0.14 M sodium dihydrogen phosphate, 0.11 M NaOH at 70°C. (b) Filter, cool, and add 2.5 ml 1 M HCI. (c) Prepare a solution of lysine by adjusting the pH of a 0.2 M solution of lysine-HCI to 7.4 by addition of 0.1 M dibasic sodium phosphate. (d) Dilute to 0.1 M lysine by addition of 0.1 M phosphate buffer pH 7.4. (e) Just before use mix one part of paraformaldehyde solution to three parts lysine solution and add solid sodium periodate to a concentration of 10 mM. This is a gentle fixative that is suitable for labile antigens such as the H2 antigen in mouse tissues.

"Lugol's iodine. 2 g Kl, 1 g iodine in 100 ml water.

If the antigen survives one or more of the fixatives, tissue is fixed in the optimum fixative and embedded in paraffin wax. Sections are cut and stained. If the processing has destroyed the antigen, this is repeated using a paraffin wax with a low melting point (45 °C as opposed to 58°C). At this stage it should be possible to select the optimum conditions for a particular antibody. The antigens found on the surfaces of lymphoid cells, such as the histocompatibility antigens and the subset-specific markers, are often unstable. Some care must be taken to preserve their integrity. To demonstrate the necessary steps, a procedure used to prepare sections for staining labile markers on the surface of lymphocytes in human tissue is given in Protocol 2. Protocol 2. Preparation of sections for staining for lymphocytic markers Equipment and reagents • • • • •

Microtome mounted in a cryostat Glass microscope slides Slices of cork OCT embedding compound Isopentane (Aldrich Chemical Co.)

• Formokcalcium (see Protocol 1) • Acetone • Chloroform:acetone (50:50, v/v) • Phosphate-buffered saline (PBS)

A. Freezing the tissue 1. Cover small pieces of tissue (maximum size 3 x 3 x 15 mm) with OCT embedding compound on a slice of cork. 2. Freeze in isopentane pre-cooled in liquid nitrogen. 3. Store the tissue and cork in plastic ampoules in liquid nitrogen.

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Michael G. Ormerod and Susanne F. Imrie Protocol 2.

Continued

B. Cutting sections 1. Cut sections, 8 um thick, on a microtome mounted in a cryostat and mount as usual on glass slides. 2. Dry sections at 37°C for 1 h. They may now be used immediately or stored. 3. To store sections, wrap them in plastic film (the type sold for 'cling' wrapping food) and store at -20°C. Before use, bring to room temperature and remove film. C. Fixation 1. 5 min in formol:calcium. 2. Dip in cold acetone. 3. 5 min in chlorofornracetone (50:50, v/v) at -20°C. 4. Dip in cold acetone. 5. Wash twice in PBS.

It should be noted that the various epitopes on an antigen might be affected differently by a fixative. If several monoclonal antibodies to the same antigen are available, each should be optimized separately. Sections of calcified tissue, such as a tumour metastasized to bone, are normally decalcified with 5% formic acid or 0.5 M EDTA before histological staining. If the antigen has survived fixation in formol:saline, it will usually also survive the process of decalcification.

3.2 Revealing hidden antigens Sometimes treatment of a section of fixed tissue with a either heat in a microwave oven or with a proteolytic enzyme will 'reveal' an apparently destroyed antigen. The mechanism by which these procedures permit a previously inhibited reaction of one of the proteins with an antibody is not properly understood. Possibly, after fixation, the epitope in question is unaffected but surrounded by a matrix of cross-linked proteins whose removal allows the antibody access. The use of a proteolytic enzyme adds to the procedure an extra variable that is difficult to control precisely. As a general practice we would not recommend it. However, in diagnostic pathology, sometimes the only available tissue has already been fixed in formalin and embedded in paraffin wax and these may not be the optimal conditions for a particular antigen. In these circumstances there is little choice but to try this approach. A suitable recipe is given in Protocol 3. A procedure for revealing antigens using treatment in a microwave oven (8) is given in Protocol 4. 192

5: Immunohistochemistry Protocol 3. A method for treatment of sections with pronase Reagents • Phosphate-buffered saline

• Pronase

Method This is used on sections from fixed tissue prior to applying the first antibody. 1. Dewax the section and take it through alcohol to water. 2. Incubate the section in PBS at 37°C for 5 min. 3. Incubate the section in PBS, 50 ug/ml pronase at 37°C for 20 min. 4. Wash in running tap-water for 5 min. 5. Wash twice in PBS. 6. Apply antisera according to the desired protocol. After this treatment, the sections are very fragile and must be handled with care.

Protocol 4. A method for the treatment of sections in a microwave oven Equipment and reagents • Domestic microwave oven equipped with a temperature probe • 3-aminopropyl triethoxysilane

• Citrate buffer: dissolve 2.1 g citric acid in 1 litre distilled water, add 26.5 ml 1 M NaOH, adjust to pH 6

Method 1. Cut 5 p.m sections and mount them on slides coated with aminoalkyl silane (see Section 6.9.1). 2. Dewax sections, take them down to water, and place the sections in a slide holder. 3. Pour 500 ml citrate buffer into a plastic container with a hole cut in the lid to take the temperature probe of the microwave oven. Heat the buffer to 90°C. 4. Place slides in the pre-warmed citrate buffer and heat in the microwave oven at 90°C for 10 min. 5. Remove slides from the oven and leave to cool for 15 min. 6. Wash sections in water and continue as for the chosen method. 193

Michael G. Ormerod and Susanne F. Imrie The effect is demonstrated in Figure 2 which shows a stain for glial fibrillary acidic protein on a section of formalin fixed human cerebellum embedded in paraffin wax using an indirect method. In Figure 2b the section was pre-treated with pronase according to the protocol in Protocol 3; Figure 2a was untreated. The primary antibody was a rabbit polyclonal antiserum raised by Mr N. Bradley (Institute of Cancer Research) and used at a dilution of 1/100. The secondary antibody was a peroxidase-conjugated swine anti-rabbit (Dako) used at a dilution of 1/100. Colour was developed using diaminobenzidine (DAB), the section counterstained with Mayer's haemalum (see Section 6.9.2), and the coverslip mounted in DPX (see Section 6.9.3). The antibody picks out the astrocytes, the fine fibrils are the cellular processes delineated by the stain. The stronger staining on the section pre-treated with pronase can be seen.

4. Choice of label The labels used to visualize an antibody fall into three main classes: fluorescent, enzymatic, and gold with silver enhancement. It is also possible to use a radioactive label followed by autoradiography. This is not a normal immunohistochemical procedure and the technique will not be discussed here.

4.1 Fluorescent labels An ideal fluorescent label has a high quantum yield, good separation between the wavelengths of excitation and emission, a wavelength of maximal absorption close to a strong line from a mercury arc lamp (used for fluorescence microscopy), and an emission wavelength suitable for photographic film and the human eye. In practice, the two substances that are in common use for fluorescent microscopy are fluorescein and rhodamine which can both be attached to protein by reaction of lysine residues with the isothiocyanate. Phycoerythrin, a fluorescent protein found in red algae, is a popular label for flow cytometry and consequently a wide range of antibodies labelled with this protein is available. Filters designed for use with rhodamine are also suitable for phycoerythrin. The advantage of using a fluorescent label is its speed. Once the slide has been incubated with labelled antibody, the coverslip can be mounted and the result read. The disadvantages are the need for specialist equipment and that one has literally to work in the dark. The architecture of the tissue and the cellular morphology are not revealed. Although the earliest immunohistochemistry employed antibodies labelled with fluorescein, this method is being superseded by methods employing enzymes.

4.2 Enzymatic labels When an enzyme is used as a label, it is visualized by means of a reaction that gives an insoluble coloured product. An ideal enzyme has a low molecular weight (for ease of attachment to Ig) and a high turnover number (to give a 194

5: Immunohistochfiirtintry

Figure 2. Human cerebellum stained for glial fibrillary acidic protein showing the effect of pre-treatment with a protease, (a) Untreated, (b) Pre-treated with pronase. The bar represents 30 um.

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Michael G. Ormerod and Susanne F. Imrie high yield of product), is absent from normal tissue, and can be used to give a product which is insoluble in water, ethanol, and xylene (so that the coverslip can be mounted conventionally). 4.2.1 Horseradish peroxidase Horseradish peroxidase (HRP) was the first such enzyme to be used and remains a popular choice. It fulfils the above criteria except that it is found in normal tissue, particularly granulocytes, erythrocytes, and cells of the myeloid series. Usually it is necessary to block the activity of enzyme endogenous to the tissue. The substrate is hydrogen peroxide and the product oxidizes a chromogen. It is commonly used with DAB that gives a brown precipitate at the site of reaction. An advantage of this stain is that, for all practical purposes, it is permanent. Care must be taken not to confuse the reaction product with endogenous brown pigment. 4.2.2 Alkaline phosphatase The substrate for this enzyme is usually a naphthol phosphate with a diazonium salt. The phosphatase releases the naphthol that couples with the diazonium salt to form a precipitate. Used in conjunction with Fast Red it gives a red precipitate that dissolves in ethanol and xylene so that the coverslip must be mounted in an aqueous medium. Alkaline phosphatase is found in several tissues including bone marrow, breast, endothelium, kidney, placenta, and intestine. That found in the intestine is a different isoenzyme and is the more robust, surviving many procedures for processing which destroy the enzyme at other sites. The red colour catches the eye and we have found this label particularly useful when trying to identify rare cells (e.g. micrometastases in bone marrow smears) (9). It has the disadvantage that the reaction product fades over a period of months. 4.2.3 Glucose oxidase Glucose oxidase is found in bacteria and is absent from mammalian tissue. The substrate is oxidized in the presence of a tetrazolium salt and the hydrogen acceptor, phenazine methosulfate. Upon reduction, the tetrazolium forms a coloured precipitate that, if the nitroblue derivative is used, is blue. If used when labelling two antigens on the same slide, it makes a pleasing contrast with the product from alkaline phosphatase. It is necessary to use Methyl Green if a counterstain of the nuclei is required. 4.2.4 Galactosidase The enzyme B-galactosidase is extracted from Escherichia coll and is readily conjugated to other proteins. The optimal pH for the bacterial enzyme (7-7.5) differs from that of human 3-galactosidase (5-5.6) so that, if the correct buffer is used, there is no need to block the endogenous enzyme. Furthermore, the latter will be inactivated by heating above 55 °C so that tissue embedded in paraffin wax will contain no active enzyme. 196

5: Immunohistochemistry 4.2.5 Conjugating enzymes to antibodies Several manufacturers now produce a range of conjugated second antibodies of high quality. Producing conjugates in the laboratory is not recommended if commercial reagents are available.

4.3 Colloidal gold This technique employs colloidal gold particles onto which an antibody has been absorbed. It was originally developed for electron microscopy where it has the advantage that gold is electron dense and that different antibodies may be labelled with gold particles of different sizes, enabling two or three antigens to be localized simultaneously. Used in light microscopy, the gold particles have to be visualized by a silver precipitation. The method has proved to be more sensitive than those employing enzymatic labels. Since immunogold is usually used in an indirect method and several manufacturers produce suitable reagents, these are usually best purchased. Streptavidin absorbed onto colloidal gold is also obtainable and can be used in conjunction with biotinylated antibodies. The pH, particle size, ionic concentration, and the concentration of protein affect the amount of protein absorbed onto the surface of colloidal gold particles. The first of these, pH, is critically important and needs to be close to the isoelectric point of the protein being absorbed. Methods for absorbing proteins onto gold are described in the article by J. Roth (ref. 2, Vol. 2), and the manufacturer, Janssen Pharmaceutica, has produced an excellent booklet giving detailed recipes for different types of protein.

4.4 Selecting a label There is no right or wrong choice of label. Selection should be guided by the tissue to be studied. Alkaline phosphatase would probably be a poor choice if a study is to be made of the gut since there is a high concentration of the enzyme in this tissue. Peroxidase is usually best avoided if the tissue contains large amounts of endogenous peroxidase or brown pigment (which can be confused by the colour produced by a commonly used substrate), particularly if the antigen is likely to be affected by the procedures necessary to eliminate these. Another important factor is the availability of labelled reagent, since for most applications it is usually easier and cheaper to buy rather than make labelled antibody. The ultimate choice may come down to the personal preference of the individual worker.

5. Methods of application The simplest method of detecting an antigen in a tissue section is to apply a labelled antibody (the so-called direct method). More commonly the primary 197

Michael G. Ormerod and Susanne F. Imrie antibody is left unlabelled and the label is attached to a different reagent that is then used to detect the primary antibody. For example, if the primary antibody is a mouse IgG, it may be detected by a labelled goat anti-mouse IgG antibody (the indirect method). Of the methods described below, the direct method gives the fastest result. The indirect method combines ease of use with acceptable sensitivity, while the enzyme-anti-enzyme and immunogold methods are the most sensitive. As in many things, there is no one 'correct' method and the choice is often governed by the personal preferences of the investigator.

5.1 The direct method The label is attached directly to the antibody (Figure 3d). To do this, the Ig must first be purified. The advantage of the method is its speed—it has only one step. The disadvantages are that it is less sensitive than other methods and that each antibody has to be labelled separately.

5.2 The indirect method A second antibody is raised to the Igs of the species from which the antibody of interest was obtained. Sections are incubated with the first antibody, washed, and then incubated with the labelled second antibody (Figure 3b). The major advantage is that, for a series of first antibodies, only one preparation of labelled second antibody is needed. This creates less work and also is more economical with the primary antibody since some reagent is always lost during a chemical procedure. The indirect method is also more sensitive than the direct method because more than one second antibody molecule can react with each first antibody.

Figure 3. The (a) direct and (b) indirect methods of visualizing the reaction site of an antibody on a tissue section. 1, primary antibody; 2, labelled second antibody.

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5: Immunohistochemistry In a particularly sensitive variant of the indirect method, the second antibody is absorbed onto particles of colloidal gold (see Section 4.3).

5.3 Enzyme-anti-enzyme methods This is a variation of the indirect method which is used to increase sensitivity. The label used in immunohistochemistry is frequently an enzyme. For ease of description, it is assumed that the first antibody is a mouse monoclonal. A mouse antibody to the enzyme is also required. A further requirement is an unlabelled antibody (e.g. raised in a rabbit) to mouse IgG. As it outlines in Figure 4, the section is incubated in the first (mouse) antibody, washed, and then incubated in an excess of rabbit anti-mouse IgG serum and washed again. A solution of complexes of enzyme-anti-enzyme is prepared by adding mouse antibodies to a solution of enzyme. This is added to the slide and, after incubation and washing, a colour reaction for the enzyme performed. Because Igs are dimers, they can react with two separate molecules of antigen. If an enzyme carries more than one epitope and the enzyme and antibody are mixed in the correct concentrations, a cross-linked network of enzyme and antibody will be formed. The purpose of the rabbit anti-mouse IgG serum is to link the enzyme-anti-enzyme complexes to the first antibody. The advantage of this method is its greater sensitivity since even more molecules of label can be added to each molecule of first antibody. It also avoids having to link covalently an enzyme to the second antibody. The disadvantages are the need for an additional reagent (the anti-enzyme antibodies) and the additional time required because of the extra step in the sequence of reactions. If the enzyme used is HRP the method is called PAP (peroxidase-anti-

Figure 4. The enzyme-anti-enzyme method. 1, primary antibody; 2, second (linking) antibody; 3, enzyme-anti-enzyme complex. Note that the primary and the anti-enzyme antibodies must be raised in the same species.

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Michael G. Ormerod and Susanne F. Imrie peroxidase); alkaline phosphatase-anti-alkaline phosphatasc (APAAP) is also frequently used, Figure 5 shows a human iymph node stained for the leucocyte common antigen (CD45) using the indirect method with alkaline phosphatase (Figure 5a) and the APAAP method (Figure 5b). Tissue was fixed in modified methacarn and embedded in paraffin wax. The primary antibody was a murine monoclonal at a dilution of 1/200. The secondary antibody in Figure 5a was a rabbit anti-mouse alkaline phosphatase used at a dilution of 1/200. In Figure 5b rabbit anti-mouse Ig was applied at 1/20 followed by mouse APAAP at 1/100. The reagents were obtained from Dako. The colour was developed using Fast Red TR, the counterstain was Mayer's haemalum (Section 6,9.2), and the coverslip mounted in glycerine jelly (Section 6.9.3). The stain demonstrates that the antibody reacted with the surface of the lymphocytes and a comparison of Figure 5a and 5b demonstrates the greater sensitivity of the APAAP as compared to the simple indirect method.

5.4 Systems using biotin-avidin Avidin, a protein extracted from egg white, has four binding sites of high affinity for biotin which is found is liver. Biotin can be covaicntly bound to either the first or second antibody which is then visualized using labelled

Figure 5. Human lymph node stained for leucocyte common antigen using (a) the indirect and (b) the APAAP method. The bar represents 200 (j.m,

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Figure 6. The biotin-avidin method. 1, primary antibody; 2, biotinylated second antibody; 3, labelled avidin.

avidin (Figure 6). Avidin has an isoelectric point close to 10 and is positively charged in neutral buffers. It is therefore likely to bind negatively charged molecules in a tissue. Streptavidin, isolated from Streptomyces avidinii, also has four binding sites for biotin but has an isoelectric point close to 7. For enzyme labelling, rather than attach an enzyme directly to avidin or Streptavidin, the enzyme may be biotinylated and unlabelled avidin used as a bridge. For increased sensitivity, this system may be used in a manner analogous to the enzyme-anti-enzyme technique. Complexes of biotinylated enzymeavidin are preformed and applied in place of the labelled avidin—the socalled ABC method (avidin-biotinylated peroxidase complex). It is claimed that larger complexes can be created than in the enzyme-anti-enzyme method thereby giving greater sensitivity. Because of the presence of biotin in liver, particular care should be exercised if avidin or Streptavidin are used on sections of this tissue.

5.5 Other methods The avidin-biotin system can be mimicked by attaching a small molecule (hapten) to the first antibody and using a labelled antibody to the hapten. Common haptens are dinitrophenol (DNP) and arsinilate. This method can be useful when labelling a section with two antibodies from the same species (see Section 8). The methods above may be used in a variety of combinations. In attempts to increase sensitivity, antibodies may be piled on antibodies. Except in 201

Michael G. Ormerod and Susanne F. Imrie special circumstances, this is generally neither necessary nor desirable. If an antigen has apparently been destroyed in fixed tissue, its presence may be revealed by using a highly sensitive method. However, the increased sensitivity will also show weak cross-reactions and amplify any slight non-specific staining. Whenever possible, it is preferable to select the correct conditions for processing a tissue.

6. Experimental methods Work sheets for the different methods are presented below. They may vary in detail from those of other authors. There is no one correct method and variation can be introduced as long as certain guide-lines are followed. In the following methods, all incubations and treatments are at room temperature unless otherwise stated. In many laboratories, this can vary from 18-24 °C and this will lead to considerable variation in the rate of reaction. If standardization is desired, and particularly if quantitative measurements are to be made, the temperature should be controlled. If a cold room is used, the times of incubation should be lengthened. It has been reported that times of incubation with antibody solutions can be reduced from 1 h to 1 min by use of a microwave oven (10). This relies on the speed of the antibody-antigen reactions at the high temperatures induced by localized heating by the microwaves.

6.1 A general method A generalized method for applying antibodies to a section is given in Protocol 5. This is for an indirect method but can be adapted for all the other methods above. It is to be used together with the detailed protocols for each type of label in the methods listed in the rest of this section. Protocol 5. The indirect method Equipment and reagents • Moist chamber (see Section 11) • PBS, 5% serum: the serum being obtained from the same species as the second antibody

• Phosphate-buffered saline (PBS) • PBS, 0.5% bovine serum albumin (BSA) . PBS, 0.01% detergent (Brij or Tween 80)

Method 1. Bring the section to water or buffer. 2. When using an enzyme conjugate, if necessary block endogenous enzyme. 3. Rinse in PBS and wipe any excess from the slide with a paper tissue. This ensures that the antiserum is not diluted on the slide.

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5: Immunohistochemistry 4. Place 100 ul of the first antibody, diluted appropriately,a over the section. Incubate for 1 h at room temperature in a moist chamber. 5. Wash the section with PBS, 0.5% BSA. 6. Wash several times in PBS, 0.01% detergent. 7. Wash with PBS and wipe excess from the slide. 8. Place 100 ul of the second (conjugated) antibody, diluted appropriately, over the section. Incubate for 1 h in a moist chamber at room temperature. 9. Repeat steps 5 and 6. 10. If using an enzymatic label, wash the slide in an appropriate buffer and develop the colour. Mount the coverslip. ' Dilute antibodies in either PBS, 0.5% BSA or preferably, in PBS, 5% serum.

The purpose of most of the additives to the buffers is to prevent Igs sticking non-specifically to the section and to remove the last traces of unreacted Ig after incubation. Protein is added to prevent the antibody absorbing onto the surface of the section and the addition of a small amount of detergent is intended to reduce hydrophobic interactions between Igs and proteins in the section. If large numbers of sections are being stained, for washing, they can be placed in racks and gently agitated in a staining trough. If there are only a few sections, solutions for washing may be kept in wash bottles. When a section is washed, the slide is held at a slant and a stream of solution directed just above the section. The sections will be washed several times and care must be taken not to wash the section off the slide. This also emphasizes the need to prepare sections of high quality.

6.2 Choosing the correct dilution of antibody When using a new antibody, a number of sections should be cut from a tissue known to contain the antigen. They should be stained using a set of dilutions which should be in geometric, not arithmetic, progression (e.g. 1:10, 1:20, 1:40, etc.). It is best to start with large steps (e.g. in fives) to find the approximate range and then to repeat the experiment using doubling dilutions. The optimal dilution is that which just stains a section close to the maximum strength. With some antisera, at high concentrations, non-specific staining of the section may be observed. In this case, a dilution has to be sought at which the non-specific staining has been diluted out while the specific stain is still acceptably strong. This should not be a problem when using a monoclonal antibody. 203

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6.3 Fluorescent labels Some fixatives (e.g. glutaraldehyde) make a tissue autofluorescent. This should be checked before starting. After following the procedure in Protocol 5, coverslips should be mounted in an aqueous, non-fluorescent mountant (such as the Fluorescence Mounting Medium from Dako Ltd.) or glycerol:40% formaldehyde (9:1, v/v). Another mountant can be made up as follows. (a) To 3 g analytical grade glycerol add 1.2 g polyvinyl alcohol (PVA) and stir. (b) When the PVA and glycerol have mixed completely, add 3 ml water, stir, and leave for 4 h at room temperature. (c) Add 6 ml 0.1 M Tris-HCl buffer pH 8.5, and keep at 50°C with occasional agitation until the PVA has dissolved.

6.4 Peroxidase The chromogens used with peroxidase often produce a brown colour that might be confused with brown pigment in the section, for example, that found in red blood cells in tissue fixed in formol:saline. It is therefore often necessary to bleach the section before starting any other procedure. This is done by immersing the slide in H2O2 (e.g. 7.5%) for 5 min and then washing well in water. A check should be made to see if this has a deleterious effect on the antigen being studied. 6.4.1 Blocking endogenous enzyme Before placing antibody on the section, incubate the slides in 2.3% periodic acid for 5 min, wash in water, rinse in 0.03% freshly prepared potassium borohydride, and wash in water. Another method often used is to incubate the slides for 30 min in 0.3% H2O2 in methanol. If these treatments destroy the antigen, an alternative is to incubate the slides in PBS, 0.1% phenylhydrazine for 5 min. 6.4.2 Chromogens for peroxidase From the large number of possible chromogens for use with peroxidase, only about five have found general use in immunohistochemistry. Their recipes are given below. i. Diaminobenzidine (DAB) with or without enhancement This is the most commonly used chromogen. It gives a brown precipitate that is insoluble in water, ethanol, and xylene. DAB has been suspected as a carcinogen and should be handled with care. (a) Just before use dissolve 100 mg DAB in 100 ml 0.1 M Tris-HCl buffer pH 7.2. (b) Add 100 ml water containing 70 ul 30% H2O2. 204

5: Immunohistochemistry (c) Immerse sections for 5 min. Wash thoroughly in water. (d) Counterstain in Mayer's haemalum and mount coverslips in DPX (see Section 6.9). It is possible to intensify the stain using either imidazole or silver. For the former, make the DAB solution above 0.01 M in imidazole before use. For silver enhancement, a method is given in Protocol 6. The result of a silver enhancement is shown in Figure 7. Sections are of human colon fixed in formalin and embedded in paraffin wax. They were stained for carcinoembryonic antigen (CEA) by the indirect method. The primary antibody was rabbit anti-CEA raised by one of the authors (M. G. O.) and used at a dilution of 1/4000. Secondary antibody and colour development as Figure 2. In Figure 7a there was no further treatment; in Figure 7b the colour was enhanced with silver. CEA is located on and in the epithelial cells. The staining is barely visible at this dilution of primary antibody without enhancement but is clearly demonstrated after the silver reaction. Protocol 6. Silver enhancement of a peroxidase/diaminobenzidine stain Reagents 2.5 mM gold chloride pH 2.3 100 mM sodium sulfide pH 7 Acetic acid Sodium carbonate Ammonium nitrate Silver nitrate Dodeca-tungstosililic acid Mayer's haemalum

' Silver reagent. Make up the following solutions by dissolving each given quantity in 100 ml twice distilled water. Solution A: 5.08 g sodium carbonate; B1: 0.83 g ammonium nitrate; B2: 0.82 g silver nitrate; B3: 3.97 g dodeca-tungstosililic acid. Add 1 ml solutions B1, B2, and B3 to 1 ml water. Add 5 ul 40% (v/v) formaldehyde solution. Add this solution to 4 ml solution A with vigorous mixing.

Method The section is first stained using a suitable method with a peroxidase labelled antibody and the colour developed with DAB. The colour may then be enhanced using the method below. 1. Wash the sections in distilled water. 2. Immerse for 5 min in 2.5 mM gold chloride pH 2.3. Wash in distilled water. 3. Immerse for 5 min in 0.1 M sodium sulfide pH 7. Wash in distilled water. 4. Immerse for 2-6 min in the silver reagent. Wash thoroughly in distilled water leaving the slide in water for 10 min between washes. 5. Immerse in 1% (v/v) acetic acid for 15 min changing the acid once during this time. Wash in water. 6. Counterstain in Mayer's haemalum and mount. 205

Michael G. Ormerod and Susanne F. Imrie

Figure 7. Human colon stained for CEA showing the effect of silver enhancement on the product of the peroxidase reaction. The bar represents 50 um,

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5: Immunohistochemistry ii. Hanker-Yates reagent This consists of one part p-phenylenediamine-HCl to one part (w/w) pyrocatechol. It gives a blackish/brown precipitate insoluble in water, ethanol, and xylene. (a) Just before use dissolve 150 mg Hanker-Yates reagent in 100 ml 0.1 M Tris-HCl buffer pH 7.6, and add 120 ul 30% H2O2. (b) Incubate sections for 15 min. Wash in water. (c) Counterstain in Mayer's haemalum and mount in DPX. Hi. 3-amino-9-ethyl carbazole This gives a red precipitate soluble in ethanol. (a) Dissolve 2 mg 3-amino-9-ethyl carbazole in 0.5 ml dimethyl formamide (DMF) in a glass tube. (b) Add 9.5 ml 0.2 M acetate buffer pH 5. (c) Just before use add 5 ul 30% H2O2. Incubate sections for 15 min. (d) Counterstain in Mayer's haemalum and mount in glycerine jelly. iv. 4-chloro-l-naphthol This can be used if a blue colour is required. The precipitate is soluble in xylene. We have not found a suitable nuclear Counterstain since Methyl Green is soluble in water. (a) Dissolve 20 mg 4-chloro-l-naphthol in 40 ml 20% methanol in Tris-saline pH 7.6, by heating to 50°C. (b) Before use add 15 ul 30% H2O2. Incubate sections for 10 min. (c) Mount in glycerine jelly. v. Tetramethyl benzidine This is an alternative blue stain, insoluble in ethanol and xylene. (a) Dissolve 5 mg tetramethyl benzidine in 2 ml dimethyl sulfoxide. (b) Add to 50 ml 0.02 M acetate buffer pH 3.3, containing 20 ul 30% H2O2 immediately before use. Incubate sections for 15 min. (c) Counterstain in Methyl Green and mount in DPX.

6.5 Alkaline phosphatase 6.5.1 Blocking endogenous enzyme Before applying the first antibody, immerse the slides in 20% acetic acid for 5 min. Wash well in water. This will destroy all alkaline phosphatase activity including that in the intestine. If it also destroys the antigen, alkaline 207

Michael G. Ormerod and Susanne F. Imrie phosphatases other than intestinal can be inhibited by making the substrate solution 1 mM in levamisole (increased to 2 mM for frozen sections of tissues rich in alkaline phosphatase such as placenta and kidney). This takes advantage of the fact that the alkaline phosphatase used for preparing conjugated antibodies is extracted from calf intestine. 6.5.2 A cautionary note Phosphate buffer may inhibit alkaline phosphatase. If the last antibody has been washed off in phosphate buffer (as recommended in the protocol in Protocol 5), the sections should be washed thoroughly in distilled water before developing the colour. 6.5.3 Chromogens for alkaline phosphatase i. Fast Red This is the usual chromogen for use with alkaline phosphatase. It gives a red precipitate soluble in ethanol. (a) Dissolve 5 mg sodium naphthol AS BI phosphate in a few drops of DMF in a glass tube. (b) Add to 5 mg Fast Red TR salt in 10 ml veronal acetate buffer pH 9.2. (c) Add levamisole if required. Filter. (d) Incubate slides for 1 h and wash in water. (e) Counterstain with Mayer's haemalum and mount in glycerine jelly. (f) The intensity of the reaction may be increased if the substrate is renewed after 30 min. ii. Fast Blue salt This gives a blue product soluble in ethanol. (a) Dissolve 5 mg Fast Blue BB salt in 10 ml 0.1 M Tris buffer pH 9, and add 5 mg sodium naphthol AS BI phosphate in DMF as above. (b) Add levamisole if required. Filter. (c) Incubate for 15 min replacing the substrate after each 5 min. Wash in water. There are problems finding a counterstain for this dye since haemalum stains the nuclei blue, Methyl Green is soluble in water, while the blue precipitate is soluble in xylene. Hi. NewFuchsin This produces a red precipitate insoluble in ethanol and xylene. (a) Mix 250 ul 4% New Fuchsin in 2 M HC1 with 250 ul 4% NaNO2 and leave to stand in the cold for 5 min. 208

5: Immunohistochemistry (b) Add to 40 ml 0.2 M Tris-HCl buffer pH 9, and add 10 mg sodium naphthol AS TR phosphate dissolved in 0.2 ml DMF. (c) Add levamisole if required. Filter. Incubate sections for 10 min. (d) Counterstain with Mayer's haemalum and mount in DPX.

6.6 Glucose oxidase The chromogen usually used with glucose oxidase is nitroblue tetrazolium that gives a dark blue precipitate insoluble in ethanol and xylene. It should not be necessary to block endogenous enzyme. (a) Dissolve 335 mg B-D-glucose and 33.5 mg nitroblue tetrazolium in 50 ml 0.05 M Tris buffer pH 8.3. (b) Heat at 37°C for 1 h in the dark and add 8.3 mg of phenazine methosulfate (this compound may be carcinogenic and should be handled with care). (c) Incubate sections for 1 h at 37 °C in the dark. Wash in water. (d) Counterstain in 0.1% Methyl Green and mount in DPX.

6.7 Galactosidase If the reagents below are used, the product is blue, stable in ethanol and xylene. It should not be necessary to block the endogenous enzyme. (a) To 7 ml PBS, 1 mM MgCl2 pH 7, add 0.5 ml 50 mM potassium ferricyanide and 0.5 ml 50 mM potassium ferrocyanide. (b) Add 10 mg 5-bromo-4-chloro-3-indolyl-B-D-galactosidase previously dissolved in a drop of DMF (this solution may be stored frozen for two months). (c) Incubate sections in the above for 1 h at 37°C. Wash in water. (d) Counterstain in 0.1% Methyl Green and mount in DPX.

6.8 Immunogold The gold is visualized by silver precipitation by a chemical process similar to that used to develop photographic film. If the tissue has been fixed in formalin and embedded in paraffin wax, it is necessary to pre-treat the sections with Lugol's iodine or some other oxidizing agent (11). The reason for this is not clear since it is not necessary when other conditions of fixation and processing are used. A suitable protocol is given in Protocol 7. 209

Michael G. Ormerod and Susanne F. Imrie Protocol 7. Silver enhancement of colloidal gold Reagents Lugol's iodine (see Protocol 1, step 6) Sodium thiosulfate Tris-buffered saline Citric acid Trisodium citrate Gum accacia Silver lactate

Hydroquinone Silver enhancement solution: 20 ml 1 M citrate buffer (1 M citric acid, 0.5 M trisodium citrate) pH 3.5; 33 ml 30% gum acacia; 15 ml silver lactate (0.11 g in 15 ml); 15 ml hydroquinone (0.85 g in15 ml); 17 ml water

A. Before adding the first antibody 1. Wash the section in water. 2. Immerse the section for 5 min in Lugol's iodine. Wash in water. 3. Rinse in 2.5% (w/v) sodium thiosulfate in water. 4. Wash in TBS, 1% Triton X-100. B. Adding the antibody 1. Apply antibodies as described in Protocol 5 substituting Tris-buffered saline (TBS) for PBS and using antibody absorbed on immunogold in the final step. 2. The immunogold method is very sensitive and can bring up any slight background staining. 3. Before applying an antibody, it may be necessary to incubate the section for 10 min in a normal serum (from the same species as the second antibody). C. To develop the stain 1. Wash in water. 2. Incubate in the silver enhancement solution in subdued light (e.g. a dark-room safe light) for 40-60 min. 3. Wash in water. 4. Counterstain. Take through ethanol to xylene (or Histoclear) and mount.

6.9 Some general procedures 6.9.1 Coating the slides During an immunohistochemical stain the slides are washed frequently. It is important that the section adheres well to the glass slide. Several different procedures are used to coat slides to improve adhesion. Those commonly 210

5; Immunohistochemistry used (gelatin:formaldehyde, gelatin, albumin, poly-L-lysine, and 3-aminopropyl triethoxysilane) are given in Protocol 8 that also indicates the circumstances under which they are used. Pre-coated slides can be purchased from PolySciences (aminoalkyl silane coated called SectionLock) or Sigma (aminoalkyl silane coated called Silane-Prep and poly-lysine coated called Poly-Prep slides). Protocol 8. Solutions used for coating slides Reagents • Gelatin • Formaldehyde • Chrome alum • Glacial acetic acid • Ethanol • Egg albumin

Sodium chloride Glycerine Thymol Poly-L-lysine (high molecular weight, Sigma) Acetone 3- aminopropyl triethoxysilane

A. Gelatin:formaldehyde (often used for frozen sections) 1. Mix 100 ml 1% gelatin (warm gently to dissolve) and 100 ml 2% formaldehyde. 2. Immerse slides for 3 sec and dry at room temperature. 3. Store at room temperature and use as required. B. Another gelatin-based adhesive (sometimes used for frozen and soft wax sections) 1. Melt 15 g gelatin in 500 ml warm water. 2. Dissolve 1 g chrome alum in 220 ml distilled water. 3. Mix the two and add 70 ml glacial acetic acid and 300 ml 95% ethanol. 4. Store at room temperature. 5. Coat slides as in step 1 above. C. Albumin (routinely used for ordinary paraffin wax sections) 1. Dissolve 2.5 g egg albumin, 0.25 g NaCI in 50 ml distilled water (warm to 37°C). 2. Add 50 ml glycerine, 0.05 g thymol. 3. Coat slides just before use. D. Poly-L-lysine 1. Immerse slides in 100 ug/ml poly-L-lysine in water. 2. Dry and store.

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Michael G. Ormerod and Susanne F. Imrie Protocol 8.

Continued

E. 3-aminopropyl triethoxysilane 1. Immerse slides in a 3% solution in either acetone or 95% ethanol. 2. Wash in either acetone or 95% ethanol. 3. Wash thoroughly in water. 4. Dry and store.

6.9.2 Counterstaining It is general practice to counterstain the nuclei in a section in order to reveal the architecture of the tissue. If the immunohistochemical stain is red, black, or brown, either haematoxylin or Mayer's haemalum is generally used. We use the latter. If the immunostain is blue, then Methyl Green is preferred. i. Mayer's haemalum (a) Dissolve 1 g haematoxylin in 1 litre of distilled water using gentle heat. (b) Add 50 g aluminium potassium sulfate, heat if necessary. (c) Add 0.2 g sodium iodate, mix well, and leave overnight. (d) Add 1 g citric acid. Mix well. (e) Add 50 g chloral hydrate. (f) Immerse the slide in the haemalum for 5-20 min (depending on the desired strength of nuclear stain) and wash thoroughly in running tapwater. (g) Dip in a saturated solution of lithium carbonate that renders the stain blue and wash in tap-water. ii. Methyl Green Use a 0.1% solution of Methyl Green in distilled water. The stain will dissolve out in water and the coverslip should be mounted in a non-aqueous mountant. 6.9.3 Mounting the coverslip If the reaction product is insoluble in ethanol and xylene, the slides are brought from water through ethanol to xylene. The coverslips are mounted in a natural or synthetic resin such as DPX. If not, they are left in water and the coverslips mounted in glycerine jelly or a similar water-based mountant. DPX consists of 10 g distrene 80, 5 ml dibutylphthalate, and 35 ml xylol. It is usually purchased ready made-up. To make glycerine jelly: (a) Dissolve 10 g gelatin in 60 ml distilled water using gentle heat. (b) Add 70 ml glycerine and 0.25 g phenol and mix well. 212

5: Immunohistochemistry (c) Aliquot into 10 ml batches and store in the cold. (d) Before use melt in a water-bath; avoid shaking as this creates air bubbles.

7. Controls and problem solving There are two types of problem encountered: the unwanted presence of stain and the unexpected absence of stain. That there is a problem will be revealed by the appropriate controls.

7.1 Controls Two types of control are needed—positive and negative. For each antibody, a block of tissue known to contain the antigen should be selected and a large number of sections cut. One of these should be included in every run in order to monitor the strength of the staining reaction. Each run should also include an experimental section on which the first antibody has been omitted. This checks for non-specific staining by the reagents used to detect the primary antibody. If the direct method is being used, this control would be omitted. If an enzyme stain is being used, a section that is developed for colour only is included to monitor endogenous enzyme activity. These controls, although necessary, do not check for non-specific staining by the primary antibody. When using a polyclonal antiserum, an aliquot of serum can be absorbed with the original antigen. This should reduce the specific and reveal non-specific stain. It will not demonstrate the presence of cross-reacting antigens. It is unnecessary to do this control with every run but it should, if the antigen is available, be performed when first using a new antiserum and on one or two key sections. This type of control is meaningless with a monoclonal antibody. The test section itself may act as a control. The distribution of an antigen is probably known; for example, an antibody against T lymphocytes should not stain epithelial cells. The correct structures should be well stained and other structures quite clean. If the 'wrong' cells are stained then non-specific staining can be suspected. In particular, stromal cells and muscle cells tend to give a 'dirty' background stain if the washing procedures are inadequate.

7.2 Problem solving Problem solving is a matter of applying simple logic. The controls discussed above should pin-point which part of the procedure is in error. Each step should be carefully considered in turn. Frequently the source of the problem is something quite trivial. For example, it is advisable to use slides with ends of frosted glass and to mark the slide clearly in pencil. Without this, it is sometimes difficult to recognize one side of the slide from the other and hence stain the wrong side of the slide. 213

Michael G. Ormerod and Susanne F. Imrie There are three types of incorrect staining—under-, over-, and non-specific staining. No staining at all on the positive control usually suggests that a reagent has been inadvertently omitted or the wrong reagent used. If a large number of sections have been stained with different antibodies, perhaps the incorrect second antibody has been used on that section (anti-rabbit on a section stained with a mouse antibody). Weak staining suggests that one of the more labile reagents has deteriorated. For example, if a peroxidase stain is in use, it is important that the solution of H2O2 is fresh. Sodium azide is frequently added to buffers to prevent bacterial growth. This compound inhibits many enzymes and could cause difficulties if a buffer containing azide has been used during the development of the chromogen. Over-staining is often caused by accidentally diluting one of the reagents incorrectly. Temperature can occasionally cause a problem. Most routine work is carried out at room temperature, which can vary as much as 10°C in a laboratory without air-conditioning. Enzyme reactions proceed much faster at higher temperatures and a procedure worked out during a chilly day might over-produce chromogen during a heat wave. If the volume of antibody solution is insufficient to cover the section properly, any evaporation during incubation will concentrate antibody at the edge of the section. This will cause over-staining. In the extreme, if solution actually dries onto part of the section, a heavy background stain will result. Non-specific staining will increase if the slide is over-stained. Apart from this, if this is a general problem, attention should be paid to the procedures used for washing the slides and, in particular, to the protein in the buffer. If a problem arises with a particular primary antibody, it may help if the section is pre-incubated for 15 min with PBS containing 5% serum from another species (see footnote to Protocol 5). If a particular secondary reagent causes a problem, it should be discarded and another purchased from a different source.

8. Detecting two antigens on the same section The application of two antibodies to the same section requires some care. If both antibodies have been raised in different species, an indirect method may be used with two differently labelled second antibodies selected to ensure that there is no cross-reaction between them. If the two antibodies are from the same species, as will often be the case when working with murine monoclonal antibodies, it may be possible to carry out a complete immunohistochemical stain for the first antigen followed by a stain for the second using a different enzyme. (The formation of the coloured precipitate will often prevent the first antibody reacting with the second set of reagents.) If this does not work, then either a direct method should be used with the antibodies each conjugated to a different label or a distinguishing compound must be attached to each anti214

5: Immunohistochemistry body. For example, one antibody could be biotinylated and then detected using labelled avidin and the other reacted with dinitrophenol (DNP) and detected with a labelled anti-DNP. The correct order of application of the reagents must be established empirically. If antibodies from two different species are used, good results are obtained usually by applying both first antibodies together followed by both second antibodies, and theoretically this should always work. However, sometimes better results are obtained if the reagents are applied sequentially. Enzyme labels are satisfactory if the two antigens are located on separate cells (e.g. when distinguishing different transplantation antigens in chimeric mice) or are found in separate cellular compartments (e.g. nucleus and plasma membrane). If not, then it is difficult to distinguish unequivocally a singly from a doubly labelled cell. In this case, fluorescent labels must be used together with a microscope that allows the operator easily to switch filter combinations (i.e. from those appropriate for fluorescein to those for rhodamine and back again) while observing a particular cell.

9. Cytological preparations Procedures for staining cytological preparations are the same as those for sections. The key to achieving good results lies in the method used to prepare the cells. On conventionally prepared smears, there is often protein or mucus overlying the cells and this can hinder good interaction between cellular antigens and antibodies. An excessive number of red blood cells is also undesirable. It is important to wash the cells, and if necessary remove erythrocytes, before making a smear or centrifuging cells onto a glass slide. Some methods for the preparation of smears of cells from serous effusions, bone marrow, and cervical scrapes are given in Protocols 9-11. In our hands, the smears were subsequently stained with antibodies to epithelial antigens such as epithelial membrane antigen or cytokeratin. The methods might have to be modified for more labile antigens. Protocol 9.

Preparation of smears from serous effusions

Equipment and reagents • Bench centrifuge . 95% ethanol • Carbowax fixative

• Lymphoprep, density 1.077 (Nyegaard) or Histopaque 1077

A. With little contamination from red blood cells 1. Centrifuge the specimen at 300 gfor 5 min. 2. Remove the supernatant and examine the deposit. 3. If the deposit is blood-stained, remove red cells (see below).

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Michael G. Ormerod and Susanne F. Imrie Protocol 9. Continued 4. If the deposit is essentially free of blood, resuspend in 20 ml PBS and recentrifuge. Repeat. 5. Remove the supernatant and resuspend the cells in as small a volume of PBS as possible. 6. Place a drop on a clean slide and smear with a second slide as for a blood film. Fix immediately in 95% alcohol and leave for a minimum of 1 h. 7. Store in alcohol or spray the smear with Carbowax fixative and store at-20°C. B. From blood-stained effusions 1. Wash the centrifuged deposit with 20 ml PBS, and recentrifuge. Remove the supernatant. 2. Mix the centrifuged deposit with 5 ml PBS. 3. Underlay the cell suspension with 10 ml Lymphoprep or Histopaque. 4. Centrifuge at 300 g for 20 min. 5. Remove the layer of nucleated cells at the top of the interface and transfer to a clean centrifuge tube. 6. Centrifuge at 300 g for 5 min and remove the supernatant. 7. Continue as in part A, step 5.

Protocol 10. Preparation of smears from aspirates of bone marrow Equipment and reagents • Bench centrifuge . 50 ml centrifuge tubes . Heparin

• Lymphoprep, density 1.077 (Nyegaard) or Histopaque 1077 • Sterile PBS

Method 1. Aspirate 1-4 ml marrow using a heparinized syringe and place in a 50 ml centrifuge tube containing 1000 U heparin plus 5 ml tissue culture medium. 2. Mix the sample and then make it up to 35 ml with sterile medium. 3. Underlay with 15 ml Lymphoprep. 4. Centrifuge at 400 g for 20 min. 5. Aspirate the cell layer, transfer to a clean centrifuge tube, and make up to 20 ml with PBS.

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5: Immunohistochemistry 6. Centrifuge at 400 g for 15 min. 7. Aspirate down to 10 ml and make up to 20 ml with sterile PBS. 8. Centrifuge at 400 g for 5 min. 9. Aspirate down to ~ 0.6 ml, mix well, and transfer to a 1 ml siliconized conical centrifuge tube. 10. Centrifuge. 11. Aspirate supernatant leaving volume approximately equal to pellet size. 12. Resuspend very gently and disaggregate the cells by taking up the suspension into a 20 ul pipette. 13. Prepare thin smears and fix immediately in absolute ethanol. Leave for at least 30 min. Store at -20°C. A similar method can be used to prepare nucleated cells from peripheral blood.

Protocol 11. Preparation of smears from cervical scrapes Reagents • Cellfix solution: 1 g dithiothrietol dissolved in 600 ml PBS plus 400 ml ethanol

• Dithlothrietol

Method 1. Take a cervical scrape in a conventional manner using a wooden spatula. 2. Break the end from the spatula and drop into 10 ml Cellfix solution. Agitate violently. Remove the spatula. The cells can be stored in this form at 4°C. 3. Centrifuge the ceils and wash in PBS. 4. Resuspend the cells in the smallest possible volume of PBS. Spread 10 ul on a clean microscope slide and allow to air dry. 5. Store the smears at-20°C.

Figure 8 shows a photograph of a smear made from a cervical scrape and stained by the indirect method. The primary antibody was rabbit anti-epithelial membrane antigen used at a dilution of 1/500; second antibody sheep antirabbit Ig conjugated to alkaline phosphatase and used at 1/100. Both reagents were produced in this laboratory. The colour was developed using Fast Red TR, counterstained in Mayer's haemalum, and the coverslip mounted in glycerine jelly. The normal squamous cell is negative while the neighbouring dyskaryotic cell has both its membrane and cytoplasm stained. 217

Michael G. Ormerod and Susanna F. Imrie

Figure 8. Two cells from a cervical scrape from a lesion diagnosed as cervical intraepithelial neoplasia, grade 2. The bar represents 10 um.

10. Quantification Immunohistochemical stains can he quantified using a microscope equipped with a system for image analysis. For further information, see ref. 12. It is probably easiest to perform on cytologica! preparations since the cells arc separated and the whole cell can be examined. When undertaking quantitative measurements, all the variables, including temperature, should be carefully controlled.

11. Equipment Apart from the normal equipment needed to cut tissue sections and to slain sections and cytological smears, the only special piece of equipment needed is a staining tray. This can he quite simple; it is used to keep slides horizontally in a humid atmosphere. We use trays, 36 cm X 36 cm, .5 cm deep with 2.5 cm high cross pieces and a lid, made of polymethylmethacrylate (Perspex, Lucite) (see Figure 9). The tray is levelled on the bench using a spirit-level and by placing pieces of card appropriately under the edges. The slides are placed on the cross pieces and a little water is put on the bottom of the tray. With the lid in place, this ensures that the solutions of antibody do not evaporate during an incubation. Some protocols require incubations at higher temperatures and it is useful to have an incubator available. To tit in an incubator, a smaller version of the

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5; Immunohistochemistry

Figure 9. Photograph of a tray suitable for immunohistochemical staining.

staining tray may be needed. This can easily be devised from a sandwich box sold in most hardware stores. For incubation at a lower temperature, the tray can be placed in a cold room. The equipment needed and the methods used to prepare blocks of tissue and to cut sections are well described in standard books of reference (13).

References 1. Stcrnberger, S. S, and DC Lellis, R. A. (ed.) (1982). Diagnostic immunohistochemistry. Masson Publishing USA Int., New York. 2. Bullock, G. R. and Petrusz. P. (ed.) (1982 and 1983). Techniques in immnnocytochemistry, Vols 1 and 2. Academic Press, London. 3. Cuello, A. C, (ed.) (1982), Immunohistochemistry, Vol 3. IBRO Handbook Series. John Wiley and Sons, Chichester. 4. Polak, J. M. and van Noorden, S. (ed.) (1983), Immunocytochemistry. John Wright and Sons, Bristol. 5. Polak, J. M. and van Noorden, S, (1987). An introduction to immunocytochemistry: current techniques and problems, 2nd edn. R. M, S. Handbook 11, Oxford University Press, Oxford, 6. Johnslone. A. and Thorpe. R. (1982). ImmiiHocheminlry in practice. Blaekwetl Scientific, Oxford. 7. Galfre, G. and Milstein, C. (1981). In Methods in enzymology (ed. J. J. Langone and H. V. Vunakis), Vol. 73, p. 3. Academic Press. New York. 219

Michael G. Ormerod and Susanne F. Imrie 8. Cuevas, E. C., Bateman, A. C., Wilkins, B. S., Johnson, P. A., Williams, J. H., Lee, A. H. S., et al. (1994). J. Clin. Pathol.,47, 448. 9. Dearnaley, D. P., Ormerod, M. G., Sloane, J. P., Lumley, H., Imrie, S. F., Jones, M., et al. (1981). Br. J. Cancer, 44, 85. 10. Chui, K. Y. (1987). Med. Lab. Sci, 44, 3. 11. Holgate, C. S., Jackson, P., Cowen, P. N., and Bird, C. C. (1983). J. Histochem. Cytochem., 31, 938. 12. Read, N. G. and Rhodes, P. C. (1993). In Immunocytochemistry: a practical approach (ed. J. E. Beesley), pp. 127-49. IRL Press at Oxford University Press, Oxford. 13. Bancroft, J. D. and Stevens, A. (ed.) (1982). Theory and practice of histological techniques, 2nd edn. Churchill Livingstone, Edinburgh.

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6

Calcium and pH imaging in living cells RICHARD M. PARTON and NICK D. READ

1. Introduction Calcium and pH imaging, involving the use of ion-sensitive fluorescent dyes, have had a profound impact on our understanding of signal transduction in animal, plant, and fungal cells. This chapter provides an introduction to the techniques involved in using the light microscope to image and measure intracellular free calcium and pH within living cells and tissues.

2. Study of calcium and pH in living cells 'Second messengers' are intracellular mediators which provide important links in signal-response coupling within cells and act in both signal propagation and amplification (1). The most common and ubiquitous second messenger within eukaryotic cells appears to be Ca2+ (2-4). The role of pH has received far less attention as a second messenger. Nevertheless, a variety of cellular responses to stimuli involve changes in cytoplasmic pH (5-8). In addition, pH homeostasis is important in regulating numerous other aspects of cell metabolism (9-11). In order to demonstrate a causal relationship between a change in Ca2+ (or pH) and a cellular response to a specific stimulus, there are several criteria which one should aim to establish: (a) A positive correlation between the response to a stimulus and the change in ion concentration. (b) The change in ion concentration should precede the response. (c) Prevention of the change in ion concentration using inhibitors or chelators should inhibit the response to a stimulus. (d) Artificial changes to the ion concentration (e.g. by using caged probes or ionophores), mimicking the changes which occur upon stimulation, should evoke a comparable cellular response.

Richard M. Parton and Nick D. Read For all of these criteria to be met the concentration of the ion in question must be directly measured in living cells and ideally over the whole period between stimulation and the initiation of the response. Furthermore, it is desirable that the method used to study Ca2+- or pH-mediated signalling provides accurate information on the dynamic spatial and temporal aspects of changes in ion concentration, and the magnitude of these changes, as all of these features can encode essential signal transduction information (4). Ideally the method of measurement should: (a) Show high selectivity for the ion of interest over other ions which may be present. (b) Allow accurate and precise quantification of ion concentration. Even very small differences in ion concentration may be biologically relevant so methods should be precise enough to measure these differences. The precision of measurement should be of the order of 100 nM Ca2+ and 0.1 pH unit. (c) Provide high spatial and temporal resolution. This may require measurements over submicrometre distances and in the millisecond range. (d) Not interfere with normal physiological activities or cellular responses (i.e. be as non-invasive as possible).

3. Imaging intracellular free calcium and pH Ion imaging is a technique which typically combines microscopy with the use of ion-sensitive fluorescent probes in order to study intracellular ion concentrations. Imaging ion-sensitive fluorescent dyes in living cells can be used to fulfil all of the criteria defined in the previous section to establish a causal relationship between a change in Ca2+ (or pH) and a cellular response to a specific stimulus. (For information on other methods of ion measurement consult refs 12-16.) The key aspects of intracellular ion imaging with fluorescent dyes are: (a) The use of Ca2+- and pH-sensitive dyes which are introduced into cells (Sections 4 and 5). These ion reporters allow intracellular ion concentration to be determined optically by virtue of their emission of photons (fluorescence) in a manner which is dependent upon ion concentration. (b) The use of microscope optics (Section 6) to focus the optical signal from the intracellular ion-sensitive probes onto a suitable detector (Section 7) to form an image. (c) The quantitative relationship between ion concentration and photons emitted by ion-sensitive dyes which allows quantitative determination of ion concentration (Sections 4 and 12). The exact quantitative relationship is dependent upon the properties of the probe (Section 4.3). 222

6: Calcium and pH imaging in living cells Ion imaging can provide quantitative, spatially and temporally resolved information on intracellular Ca2+ or H+ concentrations as they undergo dynamic changes within the living cell. In many cases, imaging may also be considered less invasive than other methods of intracellular ion measurement and so able to give a better picture of normal signal-response coupling. The technique lends itself well to being used in conjunction with the experimental manipulation of living cells and can even be employed simultaneously with other methods of ion measurement (e.g. ion-sensitive microelectrode techniques; see Section 14). Ion imaging, however, is not without its problems and limitations, which are discussed throughout this chapter.

4. Fluorescent dyes for free calcium and pH 4.1 Properties of calcium and pH dyes Ion-sensitive dyes are fluorescent molecules (i.e. they absorb light at one wavelength and emit light at a longer wavelength) which reversibly bind to specific ions. A measure of the affinity of ion binding to the dye is the dissociation constant (Kd) for that dye-ion interaction. Binding of the ion causes conformational changes in the dye molecule altering its fluorescence excitation and emission properties which can be used to report ion concentration (Figure 1). Many different fluorescent dyes are commercially available for measuring Ca2+ concentration and pH (see Tables 1 and 2) (17), and it is important to select the most appropriate dye for one's needs. Important parameters to consider are: (a) Dissociation constant (Kd for Ca2+ dyes or pKa for pH dyes). Determines the range over which ion concentrations can be measured, generally 0.1 X Kd to 10 X Kd (see Tables 1 and 2), but this range can be limited by other factors (e.g. dye precipitation, dye brightness). In the case of dextran conjugated dyes the actual Kd will vary between batches. (b) Ion selectivity. Determines how strongly the dye binds the ion of interest relative to other ions (e.g. Mg2+, H+, and K+) likely to be present in vivo. (c) Spectral properties. Dictate the imaging procedures which may be used (see Figure 1, Tables 1 and 2, Sections 4.3 and 6.3) (17). (d) Kinetics of photon emission and ion-dye interaction. Determines how well rapid, transient changes in ion concentration are reported (18). (e) Photon absorption and quantum yield. Determine efficiency with which dye absorbs and emits photons. These dictate the effective dye brightness and hence the concentration of dye required (17). (f) Photobleaching. Results in irreversible destruction of the excited dye (17). 223

Richard M. Parton and Nick D. Read

Figure 1. Excitation and emission spectra of selected Ca2+ and pH dyes determined in aqueous buffer solutions in vitro by Molecular Probes. (A) Oregon Green 488 BAPTA-1, a single wavelength dye. (B) lndo-1, a dual emission ratiometric dye. (C) Fura-2 10 kDa dextran, a dual excitation ratiometric dye. (D) Fluo-3/Fura Red, a dual excitation ratiometric dye-pair. (E) Calcium Green-1/Texas Red 70 kDa dextran, a dual excitation-dual emission ratiometric dye-pair. (F) BCECF, a dual excitation ratiometric dye. (G) Carboxy SNARF-1, a dual emission ratiometric dye. Reproduced from ref. 17 with permission.

224

6: Calcium and pH imaging in living cells This can significantly reduce the fluorescence signal and distort the relationship between ion concentration and fluorescence during an experiment (19). (g) Cytotoxicity. Determines the dye concentration tolerated by the specimen and varies considerably between different dyes, loading methods, irradiation conditions, and different cell types. (h) Size, charge, and hydrophobicity. Influence the intracellular behaviour and loading properties of dyes (Sections 4.2 and 5).

4.2 Intracellular dye behaviour It is now well established that dye-ion interaction and dye spectral responses are significantly influenced by various factors within the intracellular microenvironment, including the concentration of other ions, hydrophobicity, viscosity, and temperature (20-22). The interpretation of dye imaging results is complicated because the precise nature of the intracellular microenvironment influencing intracellular dye behaviour is unknown (Section 12.5). Indeed, intracellular dye behaviour is arguably the biggest problem in ion imaging. The influence of intracellular pH on Ca2+ dyes is a particularly

Figure 2. cSNARF-1 pH response examined in vitro. Dye solutions (50 uM) were ratio imaged using a laser scanning confocal microscope (Bio-Rad MRC600). Ratio values (580 nm/ 640 nm) at defined pH were compared for MES/Hepes buffer solutions and an artificial cytosol preparation comprising: 25% (v/v) ethanol, 60% (w/v) sucrose, 10 mM MES, 10 mM Hepes, 100 mM KCI, 20 mM NaCI, and 1 mM MgSO, (24). Fluorescence intensities at both wavelengths were roughly twice as high in artificial cytoplasm as in buffer alone. From ref. 30 with permission.

225

Table 1.Ca2+ •sensitive fluorescent dyesa Ca2+ dye

Typical excitation X (nm)b

Typical emission X (nm)6

FreeCa2+ range (uM)c

Bis-fura BTC Calcium Crimson Calcium Green-1 Calcium Green-2 Calcium Green-5N Calcium Green-C18 Calcium Orange Calcium Orange 5N Calcium Green-1 Texas Red Fluo-3

340/380 400/480 570

510 540 615

0.04-4 0.7-7 0.02-2

490

530

0.02-2

3, 10, 70, 500

490

535

0.06-6

NA

490

530

1.4-140

NA

490

530

0.03-3

NA

NA

545

575

0.02-2

+

NA

Lipophilic derivative for detecting Ca2+ near membrane surfaces More photostable than Calcium Green-1 or Fluo-3

545

580

2-200

+

NA

More photostable than Calcium Green-5N

488/568

535/615

0.04-4

NA

70

490

525

0.04-4

Fura-2 Fura-C18

340/380 340/380

510 510

0.02-2 0.02-2

+ NA

3, 10, 70 NA

FuraIndoline-C18

500/610

700

0.04-4

NA

NA

This dye consists of Ca2+-sensitive Calcium Green-1 and Ca2+-insensitive Texas Red linked to 70 kDa dextran Magnitude of Ca2+-dependent fluorescence increase greater than for Calcium Green-1 More resistant to photobleaching than lndo-1 Lipophilic derivative for detecting Ca2+ near membrane surfaces Lipophilic derivative for detecting Ca2+ near membrane surfaces

AM esterd fluorescent

+ +

Dextran conjugate (Mr, kDa)

Notes

NAe NA 10,70

Higher fluorescence output than Fura-2

NA

More photostable than Calcium Green-1 or Fluo-3 More fluorescent than Fluo-3 in Ca2+ free and Ca2+ bound forms Larger fluorescence increase upon binding Ca2+ than Calcium Green-1

Fura Red

490

660

420/480 lndo-1 Mag-Fura-2 Mag-Fura-5 Mag-indo-1 Magnesium Green Oregon Green 488 BAPTA-1 Oregon Green 488 BAPTA-2 Oregon Green 488 BAPTA-5N

350

>550 405/485

340/380 340/380

510 510

350 490

530

Quin-2 Rhod-2

340 540

405/485

490

0.02-2

+

NA

0.03-3 2.5-250 2.8-280 3.5-350 0.6-60

+ + + +

NA NA

10, 70 10, 70 NA

0.02-2

10, 70

More efficiently excited than Fluo-3 or Calcium

0.06-6

NA

Green-1 More efficiently excited than Calcium Green-2

520 490

Fluorescence decreases on binding to Ca2+; fluorescence weaker than with other Ca2+ dyes

520 490

NA 520

2-200

495 570

0.01-1 0.06-6

+ +

NA NA

Magnitude of Ca2+dependent increase greater than Oregon Green 488 BAPTA-1 First generation Ca2+ dye

* Adapted from information in Haugland (17). All dyes shown are available from Molecular Probes. All dyes are available in free acid and esterified forms except Calcium Green-Texas Red. b Other wavelengths close to the values quoted will usually produce satisfactory results. Using different wavelengths may be desirable in some cases to minimize problems of autofluorescence, or cross-talk between channels when more than one dye is used and where emission spectra of the dyes overlap (Sections 4.3, 6.4, and 12.4). c The approximate ranges over which free Ca2+ concentration can be measured using each dye have been calculated from their Kd values (Section 4.1), taking into account that Ka values are usually higher in vivo than in vitro. Kd values used in this table were those obtained in vitro by Molecular Probes and are given in Haugland (17). Note that Kd values vary between different batches of dextran-conjugated dyes. d Fluorescence of the AM ester can be a major source of error in Ca2+ measurement. ' NA = not available.

a Table 2. pH-sensitive fluorescent dyes

pHdye

Typical excitation X(nm)b

Typical emission X(nm) b

Useful pH range

Dextran conjugate (Mr, kDa)c

Notes

ACMA CarboxySNAFL-1

410 505/540 514 505/540 514 505/540 514 450/405 (470/380 dextran dye) 488 or 514 488 or 514

470 600 540/630 620 540/630 600 540/630 550 530

7.6-9.6 7.2-8.2

NA 10, 70

Apparently binds to membranes

7.2-8.2

NA

7.2-8.2

NA

7-8

NA

580/640 580/640

6.5-8 6.5-8

NA NA

375

425

6.5-8

NA

440

505

6.5-8

NA

488 or 514 490/450 490/450

580/640 530 520

6.5-8 6.5-7.5 6-7.2

490/450 490

520 530

6-7.2 6-7.2

NA 10, 40, 70 3, 10, 40, 70, 5-40 million NA NA

490/450

520

6-7.2

NA

CarboxySNAFL-2 SNAFL-calcein HPTS (pyranine)

CarboxySNARF-1 Chloromethyl SNARF-1 LysoSensor Blue DND-192 LysoSensor Green DND-153 SNARF-calcein BCECF Fluorescein Carboxyfluorescein Carboxydichlorofluoresceind Carboxydimethyl fluorescein

Better retained within cells than carboxy SNAFL dyes Low cost; no membrane permeant form of HPTS available; caged form of dye free acid available Better retained with cells than carboxy SNARF-1 Fluorescence increases on binding H+ Fluorescence increases on binding H+ Better retained with cells than carboxy SNARF-1 Has been most widely used pH dye Rapidly leaks out of cells; caged form of dye free acid available Better retained within cells than fluorescein

Better retained within cells than fluorescein

Cell tracker green CMFDA Fluorosenic sulfonic acid LysoSensor Blue DND-167 LysoSensor Green DND-189 Oregon Green 488 carboxylic acid Oregon Green 500 carboxylic acid Oregon Green 514 carboxylic acid CI-NERF DM-NERF Rhodol Green carboxylic acid LysoSensor Yellow/ Blue DND-160

490/450

520

6-7.2

NA

Better retained within cells than fluorescein

490/450

520

6-7.2

NA

Better retained within cells than fluorescein

375

425

4.5-6

NA

Fluorescence increases on binding H+

445

505

4.5-6

NA

Fluorescence increases on binding H+

490/440

520

4.2-5.7

10,70

500

525

4.2-5.7

NA

510/450

530

4.2-5.7

10,70

510/450 510/450 490

540 540 520

4-6 4-6 4-6

10, 70 10, 70 NA

360

540/440

3-5

NA

More photostable than fluorescein More photostable than fluorescein More photostable than fluorescein

' Adapted from information in Haugland (17). All dyes shown are available from Molecular Probes. All dyes are available in free acid and cell permeant forms, with the exception of HPTS. b Common excitation and emission wavelengths for both free and dextran-conjugated dyes (except where stated). Dyes with two excitation or two emission wavelengths presented in the form x/y are ratiometric dyes. (See footnote b in Table 1.) c NA = not available. d A range of polyfluorinated fluoresceins are also available (17).

Richard M. Parian and Nick D. Read serious problem (23). The effects of different environmental conditions upon dye response can be examined in vitro (Figure 2) (24). Dye retention within cells and sequestration within subcellular organelles (Figure 3) is also included amongst the problems of intracellular dye behaviour and depends upon the dye, the cell type, and the method of loading (Section 5). This, together with toxic dye effects and photobleaching, is largely responsible for defining the 'time window' over which useful data from dye imaging may be collected. Recently the use of pressure injected dextran conjugates of dyes (Section 5.7) has improved dye retention and reduced sequestration (17, 25-30), although the fluorescence of dextran dyes is still influenced strongly by the intracellular environment (27).

4.3 Single wavelength dyes, ratiometric dyes, and ratiometric dye-pairs Two major classes of ion-sensitive dyes exist: single wavelength dyes and dual wavelength (ratiometric or ratio) dyes (17, 31, 32). With single wavelength dyes, the intensity of the fluorescence emission spectrum increases in proportion to the free ion concentration (Figure 1A). The problem with using a single wavelength dye is that it is difficult to distinguish between differences in ion concentration and variations in dye brightness caused by factors such as the dye concentration, dye photobleaching, and dye leakage from a cell (see Section 7.2). This makes absolute measurement of ion concentration with single wavelength dyes difficult (33). Ratiometric dyes provide a solution to these problems because they exhibit a spectral shift upon binding to the ion of interest (31, 32) (Figures IB, 1C, 1E-G). In ratio imaging, two fluorescence images of a cell loaded with a ratiometric dye are detected at appropriate wavelengths and the ratio of fluorescence intensities for the image pair calculated (Sections 11 and 12). In principle, this ratio is independent of the amount of dye measured and proportional to the free ion concentration, allowing for improved calibration of ion concentration There are three types of ratiometric dyes: (a) Dual excitation ratiometric dyes. These dyes are usually excited sequentially at two wavelengths and two images are sequentially collected at a single emission wavelength. With Fura-2, the two excitation wavelengths specifically excite the ion binding form of the dye at the shorter wavelength and the ion free form of the dye at the longer wavelength. As the free Ca2+ concentration increases there is a shift in the excitation to shorter wavelengths (Figure 1C). With BCECF, the excitation wavelengths commonly used correspond to the pH-sensitive and the pH-insensitive (isosbestic point) parts of the emission spectrum (Figure IF). (b) Dual emission ratiometric dyes. These dyes are excited at one wavelength and fluorescence emissions detected at two longer wavelengths. With Indo-1 and cSNARF-1, the two emission wavelengths correspond to the 230

6: Calcium and pH imaging in living cells ion binding and free forms of each dye. As the concentration of free Ca2+ or protons increases there is a shift in the emission spectra of Indo-1 or cSNARF-1, respectively, to the shorter wavelengths of the ion binding forms of these dyes (Figures 1B and 1G). (c) Dual excitation-dual emission ratiometric dyes. These dyes are excited simultaneously or successively at two wavelengths and their fluorescence detected (again simultaneously or successively) at two longer wavelengths. An example of such a dye is SNAFL-calcein for pH (34). Ratio imaging can also be performed with a 'ratiometric dye-pair' (34, 35) composed of two fluorochromes. Different dye-pair combinations are currently used for Ca2+ imaging: (a) Two different Ca2+-sensitive dyes, e.g. Fluo-3 and Fura Red, (Figure 1D) which provide a combined emission spectrum that exhibits an overall shift to shorter wavelengths as the free Ca2+ concentration increases. (b) Ca2+-sensitive dye and a Ca2+-insensitive 'volume marker' dye, e.g. Calcium Green-Texas Red (Figure 1E), as a single dextran conjugate or as independent dyes; Fluo-3 and cSNARF-1 imaged at its pH-insensitive isosbestic wavelength (Figure 1G). The ratiometric dye-pair strategy is adopted because of the lack of Ca2+sensitive ratiometric dyes which can be excited at visible wavelengths (see Table 1). Ratiometric dye-pairs are not used for pH imaging because of the wide availability of ratiometric pH dyes excited with visible light (Table 2).

5. Introducing calcium and pH dyes into living cells 5.1 General considerations The introduction of dye into cells is fundamental to dye-based imaging techniques and is often the major stumbling block that decides whether or not the technique can be applied to a particular organism or cell type. The important points to consider when selecting a loading method are given below. The first four points may be taken as the basis for assessment of the success of any loading procedure: (a) Introduction of sufficient dye into cells and the degree of control over the amount of dye introduced. (b) Dye localization within the cell compartment of interest (e.g. the cytosol or specific organelles). (c) Dye retention within the cell compartment of interest. (d) The level of perturbation of cells by the dye itself and the loading procedure. (e) The number of cells that can be loaded with dye in a given time. (f) The relative ease of different loading procedures and the type of equipment required. Loading cells with dyes is not an exact science and generally involves some 231

Richard M. Parton and Nick D. Read compromising of the above requirements. So far there is no single loading method which can be successfully applied to all cells and no absolute way of performing any loading method. Dye loading methods can be divided into two basic types: (a) Methods based on cell permeant dye forms (e.g. ester loading and low pH loading). (b) Methods based on cell impermeant dye forms (e.g. electroporation and microinjection). In general, methods involving permeant dye forms are easier to perform but pose more of a problem with dye sequestration within organelles. Methods which involve cell impermeant dye forms are more difficult but often afford better control over dye delivery. The most commonly applied methods are discussed in Sections 5.1 to 5.7, and all have their relative merits and drawbacks. Making use of loading techniques and conditions described by other researchers can make life easier and at least one general reference for each method has been given below. However, small differences in the loading conditions, age and treatment of the dye, the state of the cells or tissue used, and the specific cell type, can critically affect loading success. Generally, the best loading method and conditions must be determined empirically for one's particular organism and cell type. Protocol 1 is a description of ester loading, the most commonly used dye loading method, although many of the points raised can be applied to the other methods.

5.2 Ester loading Ester loading (37) generally employs the cell permeant acetoxymethyl (AM) esterified form of dyes (Protocol 1). The ion-insensitive, esterified form of the dye is lipophilic allowing it to cross membranes. Once in the cytoplasm the ester groups are cleaved off by endogenous esterases releasing the ion-sensitive free acid form of the dye. Whilst the technique has been applied to most cell types of animals, plants, and fungi, success is very dye-, cell-, and conditionspecific. For instance AM esters of Ca2+ dyes load poorly or not at all into walled plant and fungal cells (33) whilst the AM esters of many pH-sensitive dyes are rapidly taken up by these cell types and cleaved to release intracellular free dye. Animal cells are generally easier to ester load with Ca2+ dyes. Several problems with the ester loading method have been recognized (21, 38): (a) Incomplete ester hydrolysis within the cytoplasm leading to the loading of organelles. (b) Active uptake of dye free acid from the cytosol into organelles (39) (Figure 3). (c) Poor retention within cells of the dye released by ester hydrolysis. (d) Cytotoxicity of AM esters and/or of the formaldehyde and acetic acid released within cells by ester hydrolysis. 232

6: Calcium and pH imaging in living cells (e) Altered intracellular spectral properties of ester loaded dye because of incomplete cleavage of the ester groups. (f) Background fluorescence of uncleaved dye ester (with some dyes only; see Table 1) or extracellular free dye produced by the action of extracellular esterases. Various ways have been devised to improve ester loading: (a) Using a non-cytotoxic detergent such as Pluronic-F127 (Molecular Probes, Inc.) to disperse dye ester evenly. (b) Loading at low temperature to reduce sequestration within organelles (21). (c) Continuous loading throughout an experiment (30). (d) ATP-induced permeabilization of membranes in some cell types (40). (e) Using dye forms such as cSNARF-calcein (17) and Fura-PEl (39) which can be ester loaded but have improved properties of localization and retention within the cytosol. Protocol 1. Loading cells with the fluorescent dye cSNARF-1 by the ester methoda Equipment and reagents Either an epifluorescence microscope with low light camera or a confocal microscope, equipped with a suitable filter set for imaging cSNARF-1 (excitation at 488 nm, emission > 540 nm). Alternatively, for ratio imaging the following filter set may be used: excitation 514 nm; emission-1 580 ± 15 nm; emission-2 640 ± 20 nm. Ultrapure HPLC water

A culture chamber allowing access to cells (41) cSNARF-1 AM ester (Molecular Probes Inc.): 20 mM stock in DMSO (dimethyl sulfoxide, Sigma) stored at -70°C Pluronic F-127 (Molecular Probes Inc.): 2% (w/v) aqueous stock stored at -10°C Appropriate cell culture media

Method 1. Prepare the dye solution by mixing a 2 ul aliquot of 20 mM stock cSNARF-1 AM ester in DMSO and 198 ul of Pluronic F-127 (0.04%, w/v) made up with ultrapure HPLC water. The concentration of the solvent DMSO is kept as low as possible, usually < 1% (v/v). Generally, aqueous dye ester solutions are best made up fresh and used immediately. However, the 200 uM cSNARF-1 AM stock can last for a week if kept at -10°C. 2. Prepare cells. In some cases it is desirable to hold cells down and a simple method is to use polylysine coated coverslips (polylysine applied to coverslips as a 1:100 dilution of the 1% (w/v) stock solution for 30 sec, then washed off for 5 min in running water) (see Section 9). 3. Apply the dye solution (or culture medium containing dye solution) to the culture chamber to give a final dye concentration of 2-20 uM.

233

Richard M. Parton and Nick D. Read Protocol 1.

Continued

4. Observe uptake of dye into cells by fluorescence microscopy. Simple examination of total fluorescence emission is of initial value but it is better to carry out an analysis of dye uptake using the same imaging system which will be later used for ion imaging. Examination of fluorescence at each of the two cSNARF-1 emission wavelengths allows loading conditions to be optimized for subsequent ratioing. It is important to make observations of the time taken for dye uptake into cells, the location of dye within cells, and any changes in dye location with increasing loading time. It is also important to examine cellular autofluorescence at the imaging wavelengths prior to assessment of loading (Section 6.4). 5. Where required, terminate loading by dilution with fresh medium, or by washing out excess dye by perfusion or exchange of medium in the chamber, (However, continuous loading with a low dye concentration may sometimes be found to be the best method.) 6. Assess variation in loading within the cell population (Section 5.8). 7. Assess the viability of individual dye loaded cells (see Section 15). •'Note that this protocol, with the exception of the precise equipment set-up and dye used, would be generaIly appropriate for ester loading all Ca21 and H1 dyes.

Figure 3, Confocal images of cSNARF-1 ester loaded into a growing Dryopteris affinis rhizoid. (A) 10 min after a 15 min loading period with 3 uM cSNARF-1 AM (see Protocol 1), Vacuoles excluding dye appear dark. (B) The redistribution of dye 45 min after ester loading. The vacuoles and cytoplasm are hard to distinguish from each other because dye is present in both. (C) Three hours after ester loading most of the dye has been accumulated within the vacuolar system. Bar = 15 um. From ref. 30 with permission.

234

6: Calcium and pH imaging in living cells

5.3 Low pH loading In low pH loading (42, 43) the pH of the loading medium is lowered to a value (typically between pH 3-5) at which the acid groups of the free dye become protonated (and hence uncharged) and the dye is able to directly cross the plasma membrane. Within the cytosol the pH is higher, the dye becomes dissociated, and the charged molecules are trapped in the cell. The method is most commonly used with walled plant cells. However, certain potential drawbacks of this method are: (a) Loading is usually very slow and the amount of dye loaded is often low compared with other methods. (b) Cells must be tolerant of a low external pH. (c) Sequestration of free dye within organelles is common. (d) Washout of excess extracellular dye before imaging is critical. (e) Dye often becomes irreversibly bound to the walls of plant or fungal cells.

5.4 Scrape loading Scrape loading (44) is applied to adherent cell cultures and involves transiently permeabilizing cell membranes by physical perturbation to allow membrane impermeant dye free acid or dye dextran conjugates to enter cells. The dye is trapped once membrane integrity has recovered. The main problems of the method are: (a) Long recovery times required. (b) Loss or sequestration of dye free acid (avoided when using dextran conjugates). (c) The need to remove excess dye from around cells before imaging.

5.5 Electroporation Electroporation (45, 46) involves applying short electrical pulses to cells which, by temporarily permeabilizing cell membranes, can allow dyes to leak in. It is very important to optimize electroporation conditions for the cell type used. Variables which need to be controlled include: concentration of dye in the surrounding medium, distance between the electrode and cells, voltage applied, number of electrical pulses, pulse frequency, capacitance, and the composition of the 'poration' medium. Careful attention should be paid to the composition of the 'poration' medium because of its influence over the electrical treatment. Standard culture media will often have to be modified (46). The method has been mostly used with animal cells (46), other non-walled cells such as plant cell protoplasts (33), and also some walled cells such as pollen grains and yeast cells (47, 48), Cell suspensions are most commonly used, 235

Richard M. Parton and Nick D. Read although the technique has been extended to adherent cells (46). The main problems of the method are: (a) Electroporation equipment required. (b) The potential for cell damage which necessitates caution in its application (45). Checks on cell viability are critical (e.g. by assessing membrane integrity with a membrane impermeant dye such as propidium iodide). There is evidence that cells are able to reseal their membranes within five seconds if treated carefully (46).

5.6 lonophoretic microinjection lonophoretic microinjection (49, 50) is the easier to perform of the two microinjection methods. The cell is impaled with a glass micropipette and the negatively charged free acid of the dye driven in directly by applying a negative current. Some cell types are better suited than others for microinjection. Large, round cells (e.g. oocytes) are the easiest to inject whilst small, turgid, and vacuolate walled cells (e.g. plant guard cells) are the most difficult. In many cases microinjection is the only way to load plant and fungal cells effectively. Problems with the method are: (a) Microinjection requires substantial apparatus (micromanipulators, a micropipette puller, a suitable power supply, needle holder, and capillary glass for making micropipettes). With more difficult cell types (e.g. walled plant and fungal cells), microinjection can be a very laborious and painstaking procedure requiring a high degree of dexterity and patience. (b) One of the greatest difficulties with microinjection is to obtain the 'ideal' injection needle and then to generate multiple needles with the same characteristics. This is crucial with cells which are difficult to inject. Programmable, multistage pipette pullers are useful to maintain reproducibility (e.g. the Sutler P-97 micropipette puller; Sutler Instrument Co.). (c) Damage to cells caused by impalement or, more often, when removing the needle. (d) The ability of many cells to expel or sequester dye free acid after it is introduced into the cytosol (50). (e) Cells may be perturbed by the applied current. (f) Only cells larger than 5-10 um in diameter can be injected. (g) Only charged molecules up to — 10 kDa can be injected.

5.7 Pressure microinjection Unlike ionophoretic injection, pressure injection involves forcing a volume of dye into cells by applying pressure (38, 51, 52) (see Chapter 10). Most of the points covered in the previous section on ionophoretic microinjection also 236

6: Calcium and pH imaging in living cells apply here. Pressure injection is commonly more difficult to perform than ionophoresis, although it is usually much easier to use in animal cells (where the technique can be automated) than plant or fungal cells. The micropipettes used for pressure injection tend to have wider apical apertures than for ionophoresis, especially for injecting the more viscous dextran-conjugated dyes, and this can be more damaging to cells. An important advantage over ionophoresis is that, in principle, pressure injection allows the delivery of virtually any size of charged or uncharged molecule into cells. It is particularly useful for introducing ratiometric dye-pairs (Section 4.3) at specific relative concentrations. Several different pressure injections systems exist and some use compressed gases whilst others involve liquid compression. Crude systems have little fine control over the pressure applied whilst more complex systems, such as those employing a pressure probe (51), provide very precise regulation of the application of pressure, and a digital readout of the pressures involved. An advantage of the pressure probe for use with walled, turgid plant, and fungal cells is that the system can be pressurized before injection, with the needle held against the cell wall, to prevent cell turgor pressure forcing cell contents into the injection needle upon impalement. Although in many ways pressure microinjection would appear to be the desired method for introducing dyes into cells, the technical difficulties involved and the small numbers of cells which can be loaded are often significant justification for the use of the other methods.

5.8 Quantifying the extent of dye loading The concentration of intracellular dye is important because it determines the brightness of the fluorescence signal from the cell. However, dye toxicity and possible intracellular buffering effects of the dye on the ion of interest, also need to be considered. A means of assessing the extent of loading, at least in relative terms, is important, especially as there is usually significant cell-to-cell variation in the amount of dye loaded. Absolute quantification on the basis of fluorescence is difficult because of differences in dye brightness in vivo and in vitro even when the effect of ion concentration has been taken into account (21). Nevertheless, comparing the relative fluorescence brightness of cells loaded with dye under defined imaging conditions is a useful measure. With pressure microinjection intracellular dye concentration can be estimated from the injected volume (20).

6. Equipment for fluorescence microscopy 6.1 Fluorescence microscopes The main microscope manufacturers provide a range of fluorescence microscopes suitable for ion imaging. The points to look for in a microscope are: (a) An epifluorescence microscope with appropriate dye excitation source (Section 6.3). 237

Richard M, Parton and Nick D. Read (b) Transmission optics which allow correlative microscopy with bright-field, phase-contrast, and/or differential interference contrast optics. (c) A filter arrangement which allows the rapid (0.1 sec or faster) selection of different excitation and/or emission wavelengths, and which should ideally be automated. (d) Suitable ports for the attachment of one or more image detectors (typically low light cameras). (e) A specimen stage providing good access to the specimen to allow for micromanipulation and experimental treatments. Two main types of microscope are used: upright microscopes, in which the objective is above the stage; and inverted microscopes, in which the objective is located below the stage. For the majority of ion imaging applications, the inverted configuration is preferable because it: (a) Allows the use of high magnification, high numerical aperture, short working distance objectives, in conjunction with slide culture/perfusion chambers. (b) Provides good access for microinjection. (c) Avoids the requirement for placing a coverslip on top of the sample which can result in living cells being deprived of oxygen during the course of an experiment. Upright microscopes are particularly useful when cells on the surface of a large opaque specimen (e.g. a leaf) needs to be microinjected but this requires the use of long working distance lenses.

6.2 Objectives Important considerations when selecting an objective for fluorescence work are: (a) Numerical aperture (NA): important for the efficiency of light collection and in determining the axial and lateral optical resolution. (b) UV transmission: important for use with UV excited Ca2+ dyes and for photoactivation of caged probes (Section 14). Many high NA plan apo objectives do not transmit light at wavelengths below 360 nm sufficiently well for these purposes. (c) Correction for axial chromatic aberration (particularly important for confocal microscopy). Specialized lenses corrected for chromatic aberration at UV wavelengths are necessary for UV confocal microscopy. (d) Working distance and coverslip thickness correction. The most appropriate objective to use will vary depending upon whether an inverted or upright microscope is used and whether single cells or thick tissues are examined. For further discussion about objectives refer to Chapters 1 and 2. 238

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6.3 Dye excitation sources for ion imaging The main illumination sources for exciting fluorescent dyes in ion imaging are: (a) Mercury arc lamp. Has an uneven spectral output with emission high enough for dye excitation only at specific wavelengths: 313, 334, 365, 405, 436, 546, and 577 nm. Note that these wavelengths are suboptimal for dual excitation ratioing of the important Ca2+ dye, Fura-2. (b) Xenon arc lamp. Provides a much more even emission over the 300-700 nm range than mercury lamps and is, therefore, more suitable for imaging work requiring dye excitation at both UV and visible wavelengths. A problem of xenon lamps is the high emission in the IR which can lead to heating of the specimen unless a heat filter is used in the optical path. (c) Lasers. Used as an excitation illumination source primarily in confocal and multiphoton imaging (Chapter 2). Although lasers only provide excitation at discrete wavelengths, a range is available which covers most of the useful visible and UV wavelengths for ion imaging. The most common lasers used for Ca2+ and pH imaging are: krypton-argon (emission at 488, 568, and 647 nm); argon (emission at 488, 514, and sometimes 457 nm); UV-argon (emission at 351, 364, and sometimes 457, 488, and 514 nm); and helium-cadmium (emission at 442 nm). There is an increasing trend for confocal microscopes to be fitted with multiple lasers to compensate for the limited lines available from individual lasers.

6.4 Filters for ion imaging A filter set comprises one or more excitation filters, emission filters, and dichroic mirrors (Figure 4) arranged to isolate the most appropriate excitation wavelength(s) and collect the required fluorescence emission wavelength(s). The three main types of filter used are: (a) Band width (band pass or barrier) filters. A variety of abbreviations are used for these filters including DF (discriminating filter), BP (band pass), NB (narrow band), and WB (wide band). They are characterized by only transmitting light within a defined spectral range and are used as both excitation and emission filters. (b) Long pass (LP) filters transmit all wavelengths above the stated value. (c) Dichroic mirrors or beam splitters (DRLP) reflect light of shorter wavelengths than the stated value and transmit all wavelengths longer than that value. They are used to separate excitation light from the higher wavelength spectrum of the fluorescence signal, or for separating different parts of the emission spectrum to two detector channels (e.g. for emission ratio imaging). Complete filter sets are commercially available for most Ca2+ and pH dyes from microscope manufacturers and Molecular Probes. Individual filters can 239

Richard M. Parton and Nick D. Read be purchased to assemble a customized set for particular needs from various manufacturers including Omega Optical, Ditric Optics, Baling Electro-Optics, Glen Spectra, and Oriel. When assembling a customized filter set from individual filters it is useful to start by examining the excitation and emission spectra of the dye (17) (Figure 1). It is particularly important to maximize the collection of useful dye signal whilst discriminating against contaminating cellular autofluorescence or background fluorescence. It is also important to check the literature for information on dye spectral changes which have been reported to occur within the intracellular environment (Section 4.2). For example, imaging Ca2+ with Fura-2 is significantly influenced by the nature of the intracellular environment when examined with 340/380 nm excitation ratioing (emission 510 nm). However, these problems may be reduced by 340/365 nm excitation ratioing (20).

7. Fluorescence imaging systems 7.1 General requirements The main components of the fluorescence equipment required for imaging ion-sensitive dyes are as follows (Figure 4): (a) Fluorescence microscope set-up as described in Section 6. (b) Fluorescence detector (video camera or photomultiplier tube). (c) Computer hardware and software for image capture, processing, analysis, and storage. (d) Hardware for producing hard copies of images (e.g. a slide maker or suitable printer; see Section 13.2). Selecting the most appropriate imaging equipment depends on one's specific imaging applications and budget. Of critical importance for ion imaging is the image detector used (53-55). Features of the image detector which need to be considered are: (a) Quantum efficiency. The number of the photons arriving at the detector which are actually detected (i.e. recorded as a voltage readout). (b) Sensitivity and noise. Sensitivity is related both to the quantum efficiency of the detector and the degree of noise which it generates. It may be thought of as the lowest level of photons which can be detected with an acceptable signal-to-noise ratio (Rs/n) (see Section 12.2 for further discussion). (c) Linearity. The relationship between input signal (photons) and output signal (voltage). (d) Spatial resolution. The extent to which this is limited by the detector usually depends on the number of scan lines in video-based systems or the 240

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Figure 4. Conventional fluorescence imaging system allowing simultaneous bright-field (transmitted light) and epifluorescence imaging. BF-IS bright-field illumination source, TF-620LP = transmitted light imaging filter, DIC-0 = DIC optics, C condenser, SC = specimen chamber, MS = microscope stage, OBJ = objective, P = Wollaston prism, EXIS = dye excitation light source, DM - dichroic mirror, EX-FC = excitation fitter changer, F = filter, EM-FC = emission filter changer, AN analyser for DIC, VS = video signal, VDU = video display unit. * Each camera requires a frame grabber card; for some cameras a specific, compatible type of card is required.

size and arrangement of CCD detector elements (which relates to pixels in the image). (e) Temporal resolution. This is limited by the rates of: (i) image detection, (ii) readout from the detector, (iii) digiti/ation lime for the video signal from the camera.

7.2 Conventional fluorescence imaging Ion imaging commonly combines the use of a conventional fluorescence microscope with a low light camera detector. The main disadvantage of this approach is that there is little discrimination between fluorescence from within the focal plane and 'out-of-focus' (including stray) fluorescence which 241

Richard M. Parton and Nick D. Read originates from above and below it (Figure 5). The inclusion of iiul-of-focus information and the detection of the average fluorescence over the full width of the focal plane limit spatial resolution and image quality (see Section 4.3). Confocal imaging (Section 7.3) or multipholon imaging (Section 7.4) overcome these limitations (see Chapter 2). However, attempts to reduce these problems by deconvolution procedures (56) are not to be recommended for quantitative ion imaging because of the way in which the mathematical transformations used alter numerical data within images. With imaging systems employing low light camera detectors, light from the whole imaged field is detected simultaneously, although the data may be subsequently read off the detector serially. This can allow very rapid image detection or averaging of several successive frames to produce less noisy images. Images may be captured on video or, more commonly, digitally in a computer framcstore. Camera-based systems are of two main types: tube cameras and CCD (charged coupled device) cameras (Chapter 3) (53, 54). Currently the best camera for most ion imaging applications is a cooled CCD. Recent examples of such cameras are able to collect images at close to video rate (e.g. 18.9 frames per second for a 512 X 512 pixel image size digitized to 12-bits per pixel using the AstroCam Model UltraPix FE250; AstroCam, personal communication).

Figure 5. Comparison of dye excitation, photobleaching, and collection of fluorescence emissions at the specimen in conventional, confocal, and multiphoton imaging (see Section 7). The region of excitation shown for conventional microscopy is with the excitation iris closed down to confine irradiation to only the cefl of interest. For confocal and multiphoton imaging only a single scanned spot is shown. This spot is scanned across the image to construct the final image.

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7.3 Confocal imaging Confocal imaging is a technique in which fluorescence from only a thin optical plane within the specimen is detected. This 'optical sectioning' ability eliminates the out-of-focus information normally detected by conventional imaging and provides improved spatial resolution, especially in the z axis, over conventional fluorescence microscopy. Confocal microscopy is dealt with in detail in Pawley (57) and Chapter 2. Briefly, the optical arrangement for confocal imaging is such that only fluorescence from the specimen originating in the focal plane of the excitation light passes to the detector (Figure 5). Light originating above and below the plane of focus is eliminated by having the fluorescence signal pass through a small aperture in front of the detector. Confocal microscopy has found wide application in quantitative ion imaging from single cells to tissues and intact organs (58-60). For ion imaging, a confocal laser scanning microscope (CLSM), which encompasses 'point scanning', a pin-hole aperture, and one or more photomultiplier tubes, is usually used. The important advantages of confocal microscopy with respect to ion imaging are: (a) Increased 3D resolution. This allows easier identification of dye sequestration within organelles and better interpretation of localized differences in intracellular ion concentration. It also minimizes problems of uneven dye distribution and uneven specimen thickness which can cause serious artefacts in conventional fluorescence imaging where light is detected, and hence the recorded signal averaged, over a much greater thickness of the cell. (b) The ability to image intracellular ion concentration from thin optical sections within thick tissue samples and intact organs. Unfortunately confocal optical sectioning comes at the cost of: (a) The requirement for complex and expensive equipment. (b) The potentially damaging effects of irradiating living specimens with an intense laser beam (especially significant at UV wavelengths). In confocal microscopy, regions of the sample above and below the focal plane, as well as the plane of focus, are irradiated (Figure 5). (c) The high level of fluorescence generally required to obtain reasonable image quality. This is mainly because fluorescence is only detected from a thin optical section (see Section 12.2) and additionally, the result of signal loss along the CLSM light path. (d) The slow rate of imaging due to the low numbers of photons which may be collected in a given time. For this reason theRs/nof confocal systems is often considered poor (see further discussion in Section 12.2). Accumulating or averaging successive images greatly improves the Rs/n by increasing the number of photons collected at the cost of temporal resolution. 243

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7.4 Multiphoton imaging Multiphoton imaging (which includes two-photon imaging) (61) is a recent technology which, like confocal imaging, provides improved 3D spatial resolution over conventional fluorescence imaging. However, in contrast to confocal microscopy, multiphoton imaging does not rely on any special optical arrangement to achieve this, but on the way in which the fluorescent dye is excited. The technique is performed using a laser scanning microscope system, usually a modified CLSM. The only commercial system currently available is supplied by Bio-Rad. In multiphoton imaging a dye molecule is excited by light at wavelengths which are usually > 2 X of those used for conventional fluorescence or confocal microscopy. Excitation by such low energy, long wavelength light is possible only at extremely high photon fluence so that multiple (usually two or three) photons are absorbed simultaneously and their energies roughly 'add up'. Thus in principle, dyes normally excited at 350 nm can be excited by simultaneous absorbtion of two photons at approximately 700 nm or three photons at approximately 1050 nm. Focusing the excitation laser beam into the specimen produces a cone of excitation illumination but only at the point of focus is there a high enough fluence of light to achieve simultaneous two (or three) photon absorption. This highly selective excitation means that all fluorescence emission must originate from the narrow region of the specimen focus plane and hence high 3D spatial resolution is achieved (Figure 5). The currently favoured excitation source for this type of work is a mode-locked TiSapphire laser producing 100 fsec duration pulses with a frequency of about 80 MHz over the range 700-1050 nm. Advantages of multiphoton imaging are: (a) Avoidance of UV radiation damage to cells because low energy excitation wavelengths are used and cells are only irradiated at high fluence in a narrow focal plane. (b) Reduced dye photobleaching and phototoxic cell damage (Section 9) because dye is only excited in a small region of the specimen. (c) Imaging at greater depths within thick samples due to deeper penetration by the long wavelength light used. (d) Improvement in the axial spatial resolution of caged probe photorelease (see Section 14). Although a complete multiphoton imaging system is commercially available, the technology is still under development and there are a number of problems and questions which remain to be answered, including: (a) The high cost and complexity of Ti-Sapphire lasers. (b) Dye absorbtion characteristics are very different in multiphoton excitation compared with that for single photon excitation (62). Two-photon 244

6: Calcium and pH imaging in living cells absorption spectra have been difficult to measure and as yet cannot be predicted. Emission spectra are, however, similar for conventional and two-photon excitation (62). (c) Possible harmful effects of high intensity, long wavelength irradiation. Specimen heating, particularly due to single photon absorbance of IR irradiation by water, is of major concern. This problem is reduced by using pulsed rather than continuous laser irradiation. Multiphoton imaging has been successfully employed to image free Ca2+ in living cells using Indo-1 (61, 63) and, in this respect, seems to have significant potential as an alternative to UV laser scanning confocal microscopy.

7.5 Imaging with multiple detectors Many imaging systems are designed with two or more detectors (either multiple cameras or photomultiplier tubes) and allow simultaneous detection of multiple, separate signals (64). Important applications in ion imaging are: (a) Simultaneous collection of perfectly registered bright-field (transmitted or reflected light) and fluorescence images (Figure 4). This is important for comparative analysis of dye fluorescence (or ion concentration) and cell morphology, especially in fast moving or growing cells where sequentially collected images would not necessarily correspond. (b) Simultaneous, dual emission ratio imaging where the emission signal is split by a dichroic mirror into two beams, each covering a different range of emission wavelengths. This provides the best temporal resolution and image registration for ratio imaging (see Section 12.4). With further detector channels simultaneous bright-field imaging and ratio imaging is possible. (c) Simultaneous Ca2+ and pH imaging. Simultaneous ratio imaging of Indo1 (for Ca2+) and cSNARF-1 (for H + ), co-loaded into cells, can be performed using a four channel imaging system (such as described in ref. 64) with simultaneous excitation at 350 nm (Indo-1) and 540 nm (cSNARF-1) and simultaneous image collection at 405 and 475 nm (Indo-1) and 575 and 640 nm (cSNARF-1).

8. Optimizing the performance of imaging systems The correct set-up and use of equipment is essential for achieving good, reproducible imaging data, especially in relation to quantitative analysis. The most important aspects of the imaging system to pay attention to are: (a) Optical alignment of both the excitation illumination and emission light. In confocal systems alignment can be particularly critical. Refer to manufacturer's instructions for alignment procedures. 245

Richard M. Parton and Nick D. Read (b) Routine performance checks should be carried out on equipment, particularly optical alignment and detector performance. A range of fluorescence standards for checking equipment are available (Table 3 and Protocol 2). Other test samples are often available from microscope manufacturers. Detectors may become damaged or give false readings if they become saturated with light. Detector sensitivity and 'dark signal' levels are good indicators of detector performance. (c) Care of objectives. Dirty or scratched objectives reduce fluorescence signals and can introduce optical distortions. Objectives should be checked regularly under a dissecting microscope and occasionally given a thorough cleaning with isopropanol using a new, fine, hair bristled brush (e.g. artist's sable, size 000). Objectives should be used with No. 1.5 coverslips (0.16-0.19 mm thick). Objectives with a coverslip correction collar should be correctly adjusted for the coverslip thickness used. (d) Detector settings. The correct gain and black level settings should be used to avoid detector saturation (photons arriving at the detector above the level registered as maximum signal output) and detector underflow (the level of photons reaching the detector but registering as zero signal output). A voltmeter may be used to monitor voltage output from some Table 3. Fluorescence standards Standard

Supplier

Focal Check Molecular Probes

Properties

Applications To check fluorescence image registration for multiple wavelength imaging. To determine the p.s.f.'' which allows estimation of the optical resolution and checking of microscope alignment. To determine the p.s.f.b which allows estimation of the optical resolution and checking of microscope alignment. Fluorescence intensity reference source for testing system performance.

PS-Speck

Molecular Probes

15 um spheres.a Combines two stains: one throughout the centre of the bead; the other as an outer ring. 175 nm (subresolution) fluorescent beads.a

63 nm latex beads

Bangs Laboratories

63 nm (subresolution) fluorescent beads.c

InSpeck

Molecular Probes

Beads available in a range of fluorescence intensities (0, 0.5, 1, 3, 10, 30, and 100%).a

aAvailable in a range of excitation and emission wavelengths to cover most applications. b p.s.f. = point spread function (see Section 8, Protocol 2). c Available in three forms: excitation wavelength (nm)/emission wavelength (nm) = 360/420; 555/570; 660/690.

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6: Calcium and pH imaging in living cells detectors to gain an idea of the output dark current (in the absence of light) and the magnitude of the output signal with different levels of sample fluorescence. Note that gain settings do not increase the number of photons detected but increase the output signal for a given photon level; increasing the gain increases the noise level. Protocol 2. Assessing the optical performance of a confocal imaging system using subresolution fluorescent beads Equipment and reagents • 63 nm fluorescent microspheres (polystyrene Y-G 570) from Bangs Laboratories « Glass coverslips (No. 1.5) . Silicone grease (Dow Corning, obtained from BDH Laboratory Supplies)

• Confocal imaging system (e.g. the Bio-Rad MRC600 CLSM): the system should have an electronic stepper motor to precisely contro1 focus position • 1% polylysine solution

Method 1. Coat coverslips with polylysine by placing them in a 0.1% polylysine solution for c. 10 min and then rinsing briefly with water. Remove excess water before use. 2. Circle an area of the coverslip with a thin line of silicone grease and add 50 ul of a 1:2000 aqueous dilution of the bead stock solution. Allow to settle for a few minutes before pouring off excess solution and replacing with a further 100 ul water. Cover with a second coverslip. 3. Place the sample on the microscope stage and move the required lens into position. Set up the CLSM with the largest pin-hole size, high gain and high zoom (20), and begin scanning while focusing slowly up and down until the beads are found. 4. Close down the confocal aperture to the desired level and focus on the median focal plane of the bead (i.e. where the bead is widest). Set the laser neutral density filter and photomultiplier gain to provide a bright, but not saturated signal, which does not cause excessive photobleaching of the dye. 5. Switch from normal (x-y) scanning mode to line scanning (x-z), Kalman collection filter (n = 3). Perform line scanning across the centre of the bead using the stepper motor to 'section' in 0.1 um steps from below the focal plane to above the focal plane of the bead. The resulting image is the point spread function (p.s.f.) (57). Using a Kalman collection filter makes the resulting image easier to interpret. Collect x-z images for five to ten beads.

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6. For each image plot fluorescence intensity against distance along a line through the centre of the image along both the x and z axes, respectively, and measure the distance between half-maximum intensity on each side of the maximum values. These are the full width halfmaximum (FWHM) values and provide an estimate of the axial and lateral resolution, respectively. The values obtained from five to ten images should be averaged.

9. Handling experimental material on the microscope stage One of the principle requirements for intracellular ion imaging is the maintenance of cells in a healthy state yet be able to examine them under the microscope with appropriate access for dye loading or experimental manipulation. Several things need to be considered: (a) Culture chambers. To provide the specialist culture requirements of some cell types, and to also allow experimental manipulation, a suitable culture chamber is essential. Chambers are available, or can be custom-built (41), to allow control of the medium composition (usually by perfusion of fresh medium through the chamber), temperature, and aeration. A problem with culture chambers is that they can restrict access to cells, especially for microinjection purposes. The contamination of chambers (particularly those made of plastics) with chemicals such as ionophores, dyes, and inhibitors is a common problem, even after thorough washing. This is best avoided by using disposable chambers or by keeping the same chamber for the same chemical treatments. (b) Movement of experimental material within the field of view. For cells which do not adhere inherently to the substratum, keeping a sample anchored down to reduce the problems of vibration and movement during imaging is a major issue. Methods for doing this include: adhering cells to coverslips with polylysine (from Sigma) or Cell-Tak (from Collaborative BioMedical Products); embedding cells in low temperature setting agar; or keeping cells in a minimal volume of medium. The use of a vibration isolated workstation is also an important precaution to reduce specimen movement. Holding cells in place can be a particular problem for microinjection where direct access to the cells is required. Often cells may be trapped against a surface or held still with a suction pipette. (c) Irradiation of cells. Cell irradiation for dye excitation is a serious cause of phototoxic cell damage. Although irradiation (particularly from UV light) may directly damage cells, the major problem is often photobleaching of the dye because this can result in the formation of toxic by-products 248

6: Calcium and pH imaging in living cells and free radicals. This becomes apparent when the cytotoxic effects of irradiation are compared for loaded and unloaded cells. The best ways to minimize this problem is to reduce the duration and intensity of irradiation, and the dye concentration. (d) Temperature control. This can be achieved by several means, including: (i) monitoring and controlling room temperature with an air-conditioner, (ii) perfusing medium at a specific temperature through a culture chamber, (iii) using a culture chamber with a hollow 'jacket' through which water is circulated from a temperature controlled water-bath, (iv) having a plastic hood mounted over the microscope or microscope stage which allows the air temperature surrounding the sample to be controlled (hoods of this type are available from several microscope manufacturers). (v) using a stage or culture chamber containing a resistive heater or Peltier device for temperature control. Changing sample temperature in a controlled way for experimental purposes is even more difficult and this additionally requires measurements of stage or sample temperature (usually with thermocouples). It should be noted that even small changes in the temperature of the microscope stage can result in changes in the focal position at the specimen.

10. Digital image processing The image processing and analysis involved in quantitative ion imaging can be separated into the four main categories listed below. Most commercial image analysis software packages are able to perform functions in the first two categories (see Protocol 6). (a) Numerical processing steps performed on the whole image, such as subtraction of dark or background signal or division of paired images (ratioing). (b) Sampling regions of interest (ROIs) to obtain average pixel values. (c) Relating pixel values to ion concentrations by reference to calibration data. (d) Statistical analysis of image data, including: analysis of random noise in pixel values within ROIs; comparisons of pixel values, or calibrated data, within or between individual images or populations of images.

11. Ratioing Ratioing in ion imaging refers to one or other of the following: (a) The division of successive fluorescence images, recorded at the same wavelength, by the first image in the sequence (36). 249

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(b) The division of paired fluorescence images, recorded either siimiluincously or sequentially at d i f f e r e n t e x c i t a t i o n or emission w a v e l e n g t h s (32, 34, 35). In the first case, ratioing can provide a useful visual guide for q u a l i t a t i v e imaging w i t h single wavelength dyes in order to i d e n l i f v changes in ion concentration, it usually involves collecting a sequence of images of a cell over lime and then ralioint; each image auainst the first by a pixel-by-pixel division of each image pair (Protocol 3). Problems w i t h this approach arise if there are changes in the dye d i s t r i b u t i o n , dye concentration, fluorescence i n t e n s i l y (due to photoblea.ehim;). or cell shape between successive images. The second ease is the more i m p o r t a n t and is what is most commonly referred to as 'ratio imaging (Figure 6). R a t i o imaging of this type i n v u l v e s the use of ratiometric dyes or raliomelric dye-pairs (Section 4.3). R a t i o imaging with nitiometric dyes or ratiomelrie dye-pairs can overeoine the problems of v a r i a t i o n s in fluorescence brightness which are not due lo differences in ion c o n c e n t r a l i o n . These problems arise because of v a r i a t i o n s in: (a) Dye c o n e e n l r a l i o n w i t h i n a cell or between d i f f e r e n t cells in a cell population.

Figure 6. Manipulation of cytoplasmic pH in the apical regions of growing Neurospora hyphae (by weak acid and bast; treatments) and examinee) by dual emission confocal ratio imaging of cSNARF-1. Celts were ester loaded with r.SNART-1 AM. Ratio images correspond to median confocal optical sections through colls. (A) Hyphae ot Neurospurs treated with sodium benzoale 50 mM at pH 6. (B) Hyphae of Neurospora treated with trimethylamine 50 mM al pH 7.8. Bar 10 um. Kindly provided by Sabine Fischer,

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6: Calcium and pH imaging in living cells (b) Cell thickness or thickness of the cell compartment loaded with dye. (c) Dye leakage from cells or dye redistribution between subcellular compartments. (d) Dye photobleaching. These variables are cancelled out by division of images collected at two appropriate wavelengths. Ratioing, therefore, allows considerable improvement in the quantification of ion concentration over that which can be achieved using single wavelength dyes (Sections 4.3, 12.4, and 12.5). However, ratioing does have its drawbacks, which are discussed in detail in Section 12.4. Protocol 3. Producing a ratio image Equipment • A suitable fluorescence imaging system (see Sections 6 and 7) .For image collection a Bio-Rad MRC600 CLSM was used

• A microcomputer and associated software for image analysis: the procedure described here was performed using TCSM (version 7) software from Bio-Rad; all image processing was performed off-line

Method 1. Collect paired image data for ratioing (Sections 4.3 and 7.5) and save as digital image files. 2. For each image of the image pair collect a 'background' image. In some cases it is sufficient to interrupt the light path and collect the 'dark' image produced by the detector in the absence of light. In other cases autofluorescence from the medium needs to be taken into account so an image is collected from a nearby cell-free region. The most difficult situation is where cellular autofluorescence has to be corrected for. Since imaging a cell before and after loading is not usually possible then an average or approximate autofluorescence value should be determined. 3. Correct each image of the image pair for background by performing a pixel-by-pixel subtraction of either the corresponding dark or autofluorescence images. Alternatively subtract a standard value ('offset') from each pixel based upon the average autofluorescence. 4. Remove any remaining 'speckle' (resulting from random noise) from outside the imaged cell and any regions of low signal by either a 'thresholding' or 'masking' step. Thresholding is performed by setting all pixels below a defined value to zero. The thresholding level is dictated by how noisy the fluorescence signal is; a thresholding value of three times the noise level in the true fluorescence signal (3 x standard deviations) is often used (see Section 12.2). Masking is performed by multiplying the fluorescence images by binary mask images in which

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Continued

pixel values are either zero or one. Masks are usually produced by setting all pixels in the fluorescence image which are above a threshold value to one and all below to zero. An alternative, less rigorous mask can be produced from a bright-field image according to the cell outline. 5. Define the relationship between ratio value and pixel value for the ratio image. For 8-bit image data a ratio range of 0-2 corresponding to pixel values of 0-255 is a useful starting point. However, wider ratio ranges may be needed depending upon the range of ratio values expected for the cell and the level of noise. The aim is to avoid excessive overflow where actual ratio values lie outside the defined range (some software programs give an indication of the extent to which this occurs). 6. Perform pixel-by-pixel division of the image pair. Programs for ratioing should interpret the division of a number by zero as zero rather than infinity. In some cases correction of minor misalignment in the collected image pair may be made by shifting the position of one of the images relative to the other. 7. Perform any desired visual image enhancements such as pseudocolouring, median, or smoothing filters, or conversion to 3D surface plots (Section 13.1).

12. Quantitative ion imaging 12.1 Image quality and quantitative imaging Image quality for quantitative ion imaging may be considered in terms of the useful numerical data which digital images contain. Important aspects relating to quantitative ion imaging which need to be considered are: (a) Precision of ion concentration measurement: the smallest differences or changes in ion concentration which can be reliably detected. This is mainly limited by the random variation or 'noise' of the imaging system (Section 12.2). Precision may be increased by averaging the pixels within a ROI, or averaging or accumulating successive images over time. Determination of the precision of ion concentration measurement for individual images allows meaningful statements about ion concentration to be made at the individual cell level and facilitates comparison of data sampled from within the same image or pairs of images. (b) Spatial resolution of ion concentration determination: the smallest areas over which ion concentration differences can be distinguished. This is limited by the areas over which image data is sampled in order to take 252

6: Calcium and pH imaging in living cells into account random noise rather than the limits imposed by the optics of the system (41) (Sections 12.2 and 12.3). (c) Temporal resolution of ion measurement: the speed with which images can be collected. Temporal resolution is largely dependent upon the detector system used (Section 7) and upon the speed with which image data can be stored. If a video recorder is not employed then the speed of data storage is determined by the speed of the computer set-up used. The quality and quantity of image data need to be balanced against temporal resolution. To improve temporal resolution, the quality and quantity of image data can be sacrificed by reducing image size, and thus the number of pixels sampled, and decreasing the time over which photons are counted. The highest temporal resolution can be achieved by fast line scanning in which a single line of pixels is recorded. This allows temporal resolution in the millisecond range (35). (d) Reproducibility of ion concentration determination: the degree of agreement in ion concentration measurement for independent repeats. Reproducibility depends upon day-to-day and cell-to-cell variation in the imaging system and imaging data, and how well this is taken into account. The reproducibility of calibration procedures (which includes variation resulting from the preparation of ion concentration standards) is also a factor. Reproducibility may be improved at the level of image capture by standardizing imaging settings and procedures. (e) Accurate determination of ion concentration: the certainty in absolute ion concentration measurement. The degree of accuracy depends upon image calibration and the reproducibility of calibration (Section 12.5). This dependence upon calibration makes ratio imaging much more reliable than single wavelength imaging (Sections 4.3, 11, and 12.5). Independent verification of calibration data with an alternative technique of ion measurement (e.g. ion-sensitive microelectrodes) can be an important aid in demonstrating the accuracy of measurement (49, 69, 70). Note that accuracy and precision are different. Results may be recorded with a high degree of precision but, because of a systematic error in the measurement technique, still be inaccurate. In this respect all measurements of ion concentration made by ion imaging methods should be considered as estimates of the absolute ion concentration. There are many factors that operate both during image capture and subsequent processing, which can corrupt the integrity of image data (i.e. the relationship between pixel intensity and ion concentration or pixel position in the image with respect to the region of the specimen actually sampled) resulting in image 'artefacts'. Awareness of such potential problems is essential and care is required in image capture, processing, and interpretation in order to avoid or take into account potential artefacts. Important sources of artefacts in quantitative ion imaging and suggested solutions are given in Table 4. 253

Richard M. Parton and Nick D. Read Table 4. Summary of common ion imaging artefactsa Problems

Possible solutions

Optical distortions (in 2D and 3D) which can distort the width, shape, and depth of the imaged field within a sample, and can misalign the two images used to obtain a ratio image (65-67).

1. Use objectives appropriately corrected for chromatic and spherical aberration (Section 6.2). 2. Use correct lens collar settings for coverslip thickness. 3. Reduce refractive index mismatch along optical path by modifying refractive index of culture medium or by using appropriate objective immersion medium (ideally, for cells in aqueous medium use a water immersion objective). 4. Use correction algorithms. 'Inner filter' and 'shading' effects 1. Avoid imaging deep within a sample. (excitation and emitted light attenuated or 2. Use multiphoton imaging. altered by passage through sample). Limited spatial resolution preventing 1. Use high numerical aperture objectives accurate determination of intracellular (Section 6.2). dye distribution. 2. Use confocal or multiphoton imaging (Sections 7.3 and 7.4). Poor Rs/n as a result of random variation in 1. Maximize fluorescence signal detected by: pixel values within images (68). (a) increasing dye concentration; (b) increasing intensity of excitation light; (c) using longer image integration periods or 'accumulation' of successive images; (d) averaging successive images; (e) using wider band width filters; (f) opening confocal microscope pin-hole; (g) increasing detector gain. 2. Use appropriate sampling strategies and statistical analysis to take account of random noise (Sections 12.3 and 12.6). a Note that imaging artefacts associated with properties of the fluorescent dyes are not included in this table and are discussed in Sections 4, 5, 12.4, and 12.5.

12.2 Signal-to-noise ratio There are different interpretations of the term 'signal-to-noise ratio' (Rs/n) in the current literature. Here we define the Rs/n according to Sheppard (68), as the ratio of the 'genuine' or 'useful' fluorescence signal and the variation (which may be determined as the standard deviation) (s.d.) in that signal. The Rs/n, provides a valuable measure of image quality in terms of the useful data which can be obtained. The Rs/n of an imaging system indicates its sensitivity (i.e. the smallest signal which can be reliably detected above the noise) and also the level of precision of measurement possible (i.e. the smallest differences which can be reliably distinguished). Much of the confusion about the Rs/n arises from denning 'noise' in imaging. Noise is considered here to be the random variation within images. Noise 254

6: Calcium and pH imaging in living cells originates from a variety of sources including: inherent, random, Poisson distributed statistical noise (shot noise) associated with 'counting' photons; amplifier noise; digitization noise; detector dark current (signal in the absence of light); cellular autofluorescence; stray background fluorescence; and even out-of-focus fluorescence signal. It should be noted that the contributions of the detector dark current, cellular autofluorescence, and stray background fluorescence to the overall recorded signal should be subtracted to leave the 'genuine' dye signal (Protocol 3). These quantities themselves should not, therefore, be considered as noise. However, each is itself variable (due to shot noise, amplifier noise, and digitization noise) and so subtracting the dark or background correction image from the overall recorded signal increases the variation (noise) in the corrected signal. This variation is additive for the different sources, degrading the Rs/n with respect to the true signal, thus:

and

where SR = overall recorded signal; Sg = 'genuine' signal; Sd = dark signal; Sb = background fluorescence; Sc = corrected signal = Sg; V = variation in, i.e. VSC = variation in corrected signal. It is interesting to note that subtracting an average value, for example an average value of background signal, does not increase the variation in the corrected image, whilst subtracting a variable background image does. In an ideal system the only source of noise would be the Poisson distributed statistical noise or 'shot' noise, which is inherently associated with 'counting' photons and sets the fundamental limit for the Rs/n (21): Rs/n = useful photons counted/s.d. in photons counted = n/Vn = Vn. As can be seen from this expression, with increasing numbers (n) of photons counted, Rs/n improves. More complex expressions for the overall Rs/n, which include estimates of the effects of the other sources of noise, have been derived (68). Experimentally, an estimate of the noise in an image may be obtained by capturing the same image twice in rapid succession and determining the deviation between corresponding pairs of pixels as described in Moore et al. (21). Alternatively, an even field of dye fluorescence may be imaged and, after correction for background or dark signal, the standard deviation of pixel values calculated (note, for large numbers of pixels the distribution of the noise approximates to normal). As a general guide, the magnitude of the minimum detectable signal (or difference in signal) needs to be at least three times that of the noise level. The relationship between noise and precision of ion determination is considered further in Sections 12.3 and 12.6. 255

Richard M. Parton and Nick D. Read In confocal systems, because of the often reduced numbers of photons counted, the Rs/n is often considered to be poor in relation to conventional fluorescence imaging. However, if for non-confocal imaging only fluorescence photons originating from the plane of focus are considered (i.e. the out-offocus information (Section 7.2) is not included as contributing to useful photons counted) then confocal imaging compares more favourably with conventional imaging.

12.3 Numerical data extraction The extraction of meaningful numerical data from images is a critical part of quantitative ion imaging. Digitized images are two-dimensional arrays of quantitative values organized in discrete pixels (or voxels—the 3D equivalent of a pixel). It is recommended that the actual pixel size (with respect to the specimen) should be set at a third of the optical resolution of the imaging system, according to the Nyquist sampling theory (71). The values of pixels take the form of 'grey levels' and are generally between 8-bit (256 grey levels or 28) and 32-bit (232 grey levels). The number of grey levels determines how much and how accurately information can be recorded in the image. In principle, the value of each pixel relates to the number of photons detected from that region of the specimen and, in the case of ion imaging, to the ion concentration. However, because of the random variation between pixels (Section 12.2), each pixel in an image does not accurately represent ion concentration and examining numerical data at the individual pixel level can be misleading. To take account of this noise, ROIs are 'extracted' and an average value obtained (41). Two important considerations for ROIs are: (a) The size of the ROI (number of pixels) sampled. The size of area from which an average value is extracted imposes a limit upon the spatial resolution of numerical data. It is this rather than optical resolution limits which determine spatial resolution in ion imaging. The number of pixels sampled determines to what extent random pixel variation can be taken into account and so affects the precision with which changes or differences in ion concentration can be determined. In this way the spatial resolution limit, the degree of noise in the image, and precision of ion measurement are all interdependent. Dye sequestration within subcellular compartments is another factor to consider when deciding upon sample area size. Where dye sequestration occurs some averaging may be unavoidable even with small sample areas. (b) Deciding where to place ROIs within the image. In addition to the problem of dye sequestration within different subcellular compartments described above it is also important to avoid regions of ambiguous signal, such as occur at the extreme periphery of cell boundaries or regions of low signal and high noise. Many software packages offer the possibility of defining hand-drawn or regular-shaped sample areas over one's images. 256

6: Calcium and pll imaging in living cells Although the former may he convenient for s e l e c t i n g precise regions, the l a t t e r arc heller lor slandardized. repeated sampling at a known spatial resolution (n X n p i x e l s ) . It is i m p o r t a n t to siatc the optical resolution, the pixel dimensions (determ i n e d by the optical magnification. electronic zoom, and the detector). and the numbers of pixels sampled over when reporting data extracted from images.

12.4 Quantitative ratio imaging A d i s t i n c t i o n should be made between qualitative and q u a n t i t a t i v e ratio imaging. A l t h o u g h ratio images, formed by pixel-by-pixel division of the e n t i r e fluorescence image pair, arc often displayed, t h e i r importance is visual (Figure 6, Figures 7A and 7B. Protocol 3) and they should not he used for subsequent numerical analysis. Instead iiumerical data should be extracted from corresponding ROls in the individual lluoresccnce images of ratio pairs and the average values of the sampled ROls divided (Figures 7A and 7B' Protocol 4). The reason for this is that there is a noise-dependent difference in the average r a t i o value when this is determined directly from a ratio image relative to the division of two average fluorescence values from the fluoresccnce image pair. As the noise in the fluorescence images increases the difference between these average r a t i o values also increases. This difference

Figure 7. Quantitative analysis of cytoplasmie pH in the apical regions of a growing Neurospora hypha and Dryopteris rhizoid performed by simultaneous dual emission confocal ratio imaging of cSNARF-1. (A) Ratio image of a hypha pressure injected with 10 kDa cSNARF-1 dextran. (B) Ratio image of a Dryopteris rhizoid ester loaded with cSNARF-1 AM. (A') and (B r ) Graphs of pH v. along a inidline through the cells shown in the corresponding (A) and (B). A pseudocolour scale is shown with corresponding pH values from an invitrocalibration (MES/Hepes buffer). Bars = 10 um. Modified from ref, 30 with permission.

257

Richard M. Parton and Nick D. Read becomes more significant when attempting to obtain very precise measurements of ion concentration with high spatial resolution (30). Protocol 4. Quantitative ratio analysis Equipment • A computer, associated software, and frame store for image capture, processing, and analysis. The power of the computer, the imaging software, and frame store used determine the rate at which ratioing can be performed. The procedures described are performed using Optimas 5 (Optimas Cor-

poration); and Excel 7 (Microsoft) on images captured on a Bio-Rad MRC600 CLSM (BioRad PIC format files). All image analysis described is performed off-line. All procedures may be performed using appropriate alternative programs,

Method 1. Follow Protocol 3, steps 1-4. Equivalent processing may be done directly in Optimas 5 after converting the Bio-Rad PIC format images to TIFF format. 2. Extract average fluorescence values from defined ROIs at corresponding positions in the paired fluorescence images using Optimas 5 and calculate the ratio of these average values. (How to choose the size of ROIs and where they should be placed within the fluorescence images are discussed in Section 12.3.) 3. Extract individual pixel values from the defined ROIs within the paired fluorescence images (a macro to perform this can be provided by Optimas user support) and import into a spread sheet (e.g. Excel 7). 4. Determine the degree of variation (i.e. the standard deviation) in the pixel values within ROIs using the facilities available in Excel. From this the signal-to-noise ratio can be determined (Section 12.2). More complex statistical analysis can also be performed, as discussed in Section 12.6, to quantify the effect that variation in the fluorescence images has on the precision of determining the final ratio value. 5. Relate the ratio values to ion concentration according to the experimental procedure described in Section 12.5 and Protocol 5. Analysis of the variation in ratio value for ROIs of different sizes (n x n pixels) provides estimates of the precision and spatial resolution limits to which ion concentration can be measured (see Figure 8 and Section 12.6).

Although ratio imaging potentially provides the most accurate and precise means of quantitative ion imaging, it is not without its problems which can lead to serious errors in absolute ion concentration measurement if they are not taken into account. In addition to the detrimental effect of noise in the individual fluorescence images used to generate the ratio data (see above), 258

6: Calcium and pH imaging in living cells the main problems affecting the relationship between ratio value and ion concentration are: (a) Temporal misalignment of the fluorescence image pairs.'This is generally a problem associated with the sequential capture of fluorescence images for ratioing. Problems arise when cell movement or growth occurs between capture of the first and second image of a ratio pair resulting in mismatch of the two fluorescence images. This places significant demands upon the rate of image capture, necessitating: (i) Rapid changes in excitation filters (for dual excitation dyes) or in emission filters (for dual emission dyes). (ii) Rapid sequential image capture and saving to computer disk. Simultaneous dual wavelength excitation imaging and/or dual wavelength image capture is possible in some cases (Sections 4.3 and 7.5; Figures 6 and 7) and can provide a solution to this problem. (b) Positional misalignment of the image pair. This problem can occur with two images acquired simultaneously by dual channel imaging if the two detectors are misaligned, causing misregistration of the two fluorescence images in the x-y plane. Additionally, chromatic aberration can cause misalignment in the z plane of both sequentially and simultaneously captured images where the two wavelengths differ significantly. To avoid these problems, precise optical alignment of the detector system, and correction for the different wavelengths used, are necessary. (c) 'Bleed-through' ('cross-talk') of fluorescence between detector channels. Bleed-through occurs when there is incomplete discrimination between the emission spectra of the two dye forms (ion bound and ion free) of a ratiometric dye or the two dyes of a dye-pair. This problem is most significant in the latter case when using two dyes for ratioing (Section 4.3). The extent of contamination between imaging channels should be checked by measuring the fluorescence signal in both imaging channels from each dye imaged on its own. Changing the filter set, changing the relative proportion of each dye in the mixture, or the detector gain settings of each imaging channel may correct the problem. Alternatively, a correction equation may be derived empirically by examining the degree of bleed-through at various dye concentrations (64). (d) Autofluorescence differentially contaminating different channels. The problem may be resolved by a similar approach to that discussed in the last point by using appropriate filters or gain settings. (e) Differential rates of photobleaching of the two forms of a ratiometric dye or the two dyes of a ratiometric dye-pair. Although this would be expected to be more of a problem with ratiometric dye-pairs it has been shown to be a problem with the ratiometric dye Indo-1 (19). Dye photodegradation should be minimized by reducing excitation illumination. 259

Richard M. Parton and Nick D. Read (f) Differential rates of dye leakage from the cell or sequestration within organelles of the two dyes of a ratiometric dye-pair. This has been overcome in the case of Calcium Green-Texas Red (Table 1; Figure 1E) where the two dyes are co-conjugated to a 70 kDa dextran.

12.5 Calibration of dye response One should be aware that, although the ultimate aim of quantitative ion imaging is to obtain absolute values of ion concentration, the actual values obtained are, in reality, only estimates. How well these estimates reflect the true ion concentration values is limited primarily by how the dye behaves in the intracellular environment, and how well the dye response is calibrated. The relationship between fluorescence emission and ion concentration may be described in terms of the equilibrium equation for the binding of dye molecules to free ions. The following equations apply for 1:1, dye:ion interaction, as occurs with Fluo-3 or Fura-2 (31, 72). For a single wavelength dye: [ion] = Kd (dye-ion complex)/(free dye) = Kd (F - Fmin)/(Fmax - F) Similarly, for a ratiometric dye: [ion] = Kd F2free/F2bound (R - R m in)/(Rmax - R)

where Kd = dissociation constant; F = measuredfluorescence;Fmin= the fluorescence when all dye is uncomplexed, i.e. at zero [ion]; Fmax = the fluorescence when all dye is complexed with the ion, i.e. at saturating [ion]; R = the ratio of fluorescence intensity at wavelength 1 divided by the fluorescence intensity at wavelength 2; Rmin = fluorescence ratio at zero [ion]; Rmax = fluorescence ratio at saturating [ion]; F2free = dye fluorescence at wavelength 2 with zero [ion]; F2bound = dye fluorescence at wavelength 2 with saturating [ion]. For derivations see Grynkiewicz et al. (31) and Thomas and Delaville (72). Calibration is primarily concerned with the determination of the .Kd value of the dye and the values of Fmin and Fmax (or Rmin and Rmax for a ratio dye). These values must be determined experimentally; published Kd values should be used with caution (20, 73). The Kd values of ion-sensitive dyes are usually higher in vivo than in vitro (17). With single wavelength dyes, Kd values are usually determined in vitro by imaging a series of dye buffer solutions over a range of known ion concentrations. Often a modified medium is used with ion concentrations (e.g. Mg2+, K+, Na+, etc.), viscosity, and hydrophobicity adjusted in an attempt to mimic the intracellular environment (24, 72). The Kd is rarely determined in situ as this requires that there is no change in intracellular dye concentration during the calibration procedure, a situation which is difficult to guarantee. As a result of the differences in the dye content of individual loaded cells (Section 5.8), meaningful calibration for a particular cell necessitates the determination of Fmin and Fmax for that individual cell, in 260

6: Calcium and pH imaging in living cells situ. These difficulties often restrict the use of single wavelength dyes to the qualitative analysis of changes in ion concentration. Kd values for ratiometric dyes can also be determined by in vitro experimentation. However, with such dyes the problem of variations in intracellular dye concentration is less significant. Kd values are, therefore, commonly determined from a three point in situ calibration (Protocol 5). This involves recording ratio values at zero and saturating ion concentrations (Rmin and Rmax, respectively) and at an intermediate ion concentration and then fitting the data to the equation above. Kd(F2free/F2bound) is sometimes determined experimentally as a combined constant. Note that when wavelength 2 is the ion-insensitive wavelength (isosbestic point) then F2free equals F2b0und so Fatree/F2bound = 1. The calibration of ratiometric dye-pairs which consist of two ion-sensitive fluorochromes (Section 4.3) is complicated because two different Kd values for Ca2+ binding are involved. This means that the standard equations for calibration are no longer valid, although experimentally derived calibration curves can be used. 12.5.1 Calibration strategies for calcium and pH imaging The ability to obtain good calibration data is severely hampered because dye behaviour varies under different environmental conditions (see Section 4.2). Due to the lack of knowledge about the dynamic physical and chemical nature of the intracellular environment, it is very difficult (if not impossible) to reproduce such conditions in vitro. As a result, experimenters often strive to calibrate dyes in situ by, for example, using ionophores in conjunction with externally applied solutions of known ion concentrations to clamp the intracellular ion concentration to defined values (Table 5 and Protocol 5). Detailed protocols for in situ calibration procedures may be found in Thomas and Delaville (72). Problems with these methods include: (a) Unstable clamping of cytoplasmic ion concentration, often because of the activities of cellular homeostatic mechanisms. Procedures sometimes incorporate metabolic or ion pump inhibitors to combat this. (b) Rapid loss of dye or sequestration within organelles which can cause apparent shifts in cytoplasmic ion concentration. (c) Cellular disruption and alteration of the intracellular environment. Although in situ calibrations are sometimes referred to as in vivo calibrations, this is not always an appropriate term as the effects of ionophore treatment and disruption of the intracellular ion concentration tend to be toxic with effective clamping. Because of the difficulty in interpreting the results obtained by in situ methods is in genuinely fixing the cell at the desired ion concentration the best calibration data incorporates an independent method of ion concentration measurement such as ion selective microelectrodes (49, 69, 70). Multiple point calibrations, which allow accurate curve fitting to the data, 261

Table 5. Summary of techniques used for calibrating ion imaging data Types of calibrations

Result

Advantages/problems

Ease

Multiple point response curvea

A good standard for comparison but the Kd of the dye in vitro often varies from that in vivo (Sections 4.1 and 4.2). A basic estimation of absolute [ion]1.b

Very easy, routine Relatively easy. routine

Intheory, the best estimation of absolute [ion],. Problems with incomplete clamp of [ion]i, cellular disruption/toxicity, loss of dye, and dye compartmentalization. Problems as above, especially incomplete clamp of [ion],. Only useful for pH.

Difficult to achieve good results; set-up is not routine Easy to perform but generally unsatisfactory Technically demanding; not routine

In vitro Buffer solutions: - MES/Hepes for pH -Ca2+/EGTA for Ca2+ 'Pseudocytosol' buffer solution (24)

Multiple point response curve

In situ lonophore permeabilization of cells (47, 68; Protocol 5)

Single pointto a few stepped changes

Cell permeant weak acids and bases (e.g. propionic acid and trimethylamine) (13) Combined use of dye imaging and ion-sensitive microelectrodes (Sections 12.5.1 and 14)

As above

Single point to a few stepped changes

Requires complex apparatus and technical expertise but potentially the best method—[ion], need not be clamped to specific values.

a Response curve = dye fluorescence (or ratio) versus ion concentration. b (ion]( = intracellular ion concentration.

6: Calcium and pH imaging in living cells are desired. Where a full dye response curve is obtained, calibrated values may be read directly without calculating the Kd, Rmax, and Rmin- Often, however, two or even one point calibrations are all that are possible during an experiment. In such cases the dye response over the rest of the ion concentration range may be inferred by reference to in vitro data. Protocol 5. In situ calibration Note that this protocol is a generalized description of the use of ionophore or protonophore methods for in situ calibration. The procedure is of most value with ratio imaging techniques where the result may be applied to the entire population of cells if standard imaging settings are used throughout. With single wavelength imaging calibration must be performed for each cell imaged. Equipment and reagents • Incubation chamber allowing access to cells, preferably with provision for perfusion of medium (Section 9) • Equipment for dye loading, if necessary (see Sections 5.5-5.7) • Fluorescent dye • lonophore or protonophore: ionomycin (Sigma) or Br A23187 (Molecular Probes Inc.) for Ca2+ and nigericin (Sigma) or nigericin/valinomycin/K+ for pH (17, 72, 74). Different cell types exhibit varied responses to different ionophores so it is advisable to try different ionophores and various concentrations (generally over the range 10 uM up to mM concentrations). • MES (Sigma)

> A suitable fluorescence imaging system (see Sections 6 and 7) • Hepes (Sigma) • EGTA (Sigma) • Calibration media (which may be based on the standard cell culture medium) with the ion of interest buffered at known concentrations. Where the ionophore used is known to allow the transport of other ions, the concentration of those species should be fixed at the concentration expected within the cell. It is also useful to regulate the ionic and osmotic strength of the calibration media over the range of different ion concentrations used.

Method 1. Prepare calibration media. pH can be buffered over the range 5.5-8.5 using 10 mM MES/Hepes whilst Ca2+ concentration can be controlled with 10 mM EGTA mixed with varying concentrations of CaCI2 (75). Temperature can affect both the buffers used to control ion concentration and the dyes used for imaging and should also be taken into account. 2. Prepare cells in the culture chamber for loading and treatment. Cells should be adhered to the substratum or held in position (Section 9) to prevent movement during the changes in calibration medium. 3. Load cells with dye (Section 5). 4. Collect the required experimental imaging data. 5. Apply calibration medium buffered at the first ion concentration in the calibration series in the presence of ionophore and allow cells to equilibrate. Images should be collected at intervals until a steady

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Richard M. Parton and Nick D. Read Protocol 5. Continued state is reached. Experience will reveal how long equilibration should take and whether a reliable steady state can be reached before declining cell health and image quality cause the observed values to change. 6. Apply the second calibration buffer and repeat the imaging time course. Continue the procedure with further ion concentration increments covering at least minimal (effectively zero) free ion concentration, dye saturating ion concentration, and an intermediate value. 7. If cells lose fluorescence or become significantly disrupted during the clamping procedure it may only be possible to impose one ion concentration change. This is a severe limitation when using single wavelength dyes. However, in the case of ratio imaging, the calibration may be continued by repeating the experiment with fresh cells for each ion concentration buffer in the calibration series. 8. Finally, if possible, return cells to the initial ion concentration calibration buffer. The value obtained should be the same as that recorded originally. 9. Perform a simple in vitro calibration with free dye in the media used for in situ calibration. Use the appropriate form of the dye in the in vitro calibration (i.e. free acid or dextran conjugate). This provides a reference to compare calibrations on different days with different instrumental settings. 10. Construct calibration curves (see Figure 8} by plotting data points or fitting the data to the theoretical curve described by the equations given in Section 12.5. The ion concentration may be derived directly for the calibration curve or calculated using the appropriate equation.

12.6 Statistical analysis of image data Statistical analysis of quantitative image data is often confined to the analysis of variation in measured ion concentration (after suitable calibration), within populations of cells or between cell populations given different experimental treatments. This generally involves using simple methods such as the students t-test (for confidence intervals and comparison of means). Such an approach tends to ignore the variation in the pixel data within the individual digital images. However, as the limits of quantitative imaging are pushed to their extreme (in order to make more precise measurements of ion concentration with better spatial resolution) the need for adequate analysis at the pixel level becomes more pressing. Statistical analysis of pixel value data has important applications in: (a) Examining the inherent variation in image data (particularly in relation to the Rs/n) in order to define the limits of precision and resolution for ion 264

6: Calcium and pH imaging in living cells

Figure 8. Example of an in vitro calibration curve for cSNARF-1 free acid with error bars indicating the precision of measurement. Spatial resolution was determined by the 1 4 x 1 4 pixel sampling area size (ROD. Error bars in all graphs are the 95% Bayesian interval for the mean ratio of the small sample areas based on variation between individual ratio values of each pixel (see Sections 12.3 and 12.6 for further explanation). Modified from ref. 30 with permission.

(b)

(c) (d)

(e)

measurement (see Sections 12.2 and 12.3; Figure 8) (30). Image collection parameters and image sampling can subsequently be optimized with respect to the precision and resolution of ion measurement necessary to allow meaningful biological interpretation of the data. Analysis of spatial inhomogeneities in ion concentration within individual images (of individual cells). A good example of this is in investigating the presence of intracellular ion concentration gradients associated with polarized cell growth (30, 76). Analysis of ion concentration changes between images of individual cells collected sequentially in a time course. Comparison of differences in ion concentration in individual cells of a cell population. This allows the heterogeneity in individual cellular responses to be recorded. If data is averaged across cell populations rather than being considered at the single cell level such heterogeneity is lost. Distinguishing between pixel variation within images (a fundamental limitation imposed by the imaging system), cell-to-cell variation within a 265

Richard M. Parton and Nick D. Read uniformly treated cell population, and differences between experimentally treated and untreated cells. Statistical methods (e.g. analysis of variance testing or non-parametric tests) (77) are available to analyse the contribution of each source of variation to the overall observed variation. Image data can pose particular problems for statistical analysis: (a) Extraction of individual pixel values from digital files, which is not available as a standard function in many image analysis software packages (Protocol 4). (b) Problems related to sampling pixel data from defined ROIs within digital images of cells for analysis of pixel variation. Images of cells, unlike 'ideal' in vitro test images, are subject to additional sources of pixel variation including regions of different dye concentration and different ion concentrations. How such variation is recorded is highly dependent upon both the size and placing of ROIs within an image (Section 12.3). (c) The statistical nature of the inherent variation in photon counting ('shot noise')—which is Poisson distributed (Section 12.2). This dictates that the variation in the fluorescence signal is proportional to the mean fluorescence intensity and so, due to the relationship between fluorescence intensity and ion concentration, the precision of measurement itself varies with ion concentration. (d) The propagation of variation initially present in a captured image during subsequent image processing steps, such as image subtraction and division (Section 12.2). Standardizing image collection with respect to average fluorescence intensity (and Rs/n) by using standard imaging conditions and image collection parameters, simplifies statistical analysis and allows more rigorous comparison of data. The inherent variation between pixels due to noise generated by the imaging procedure may be investigated with different image collection parameters by examining test solutions (uniform dye/buffer mixtures) of known ion concentration in vitro. These results can be compared with the between pixel variation observed in vivo, within individual images of dye loaded cells. This variation is likely to increase with uneven dye distribution and specimen thickness, dye sequestration within organelles, and local domains of different ion concentration. Ratio imaging poses a significant problem with regard to statistical analysis. .In ratio imaging, the average ratio must be calculated by dividing average fluorescence intensity values of images collected at the two wavelengths (see Section 12.4 and Protocol 4). Thus, statistical analysis cannot be performed directly on the ratio image pixel data. The pixel variation within the fluorescence images should be considered and for this to be done correctly the relationship between individual corresponding pairs of pixel values (covari266

6: Calcium and pH imaging in living cells ance) needs to be determined. This requires more advanced statistical analysis, for example the Bayesian approach employed by Parton et al. (30). With more complex problems such as these, expert statistical advice should be sought.

13. Visual presentation of image data 13.1 Visual image enhancement There are various ways in which digitized images may be visually enhanced. The most important for ion imaging are pseudocolouring and 'smoothing' (using the various smoothing, averaging, and median filters available with many image processing software packages). Pseudocolouring, assigning pixels different colours according to their numerical value, is particularly useful in drawing attention to small or localized differences and is routinely applied to images of ion distribution, usually ratio images. A pseudocolour scale with a specific 'look-up table' (LUT) can be easily custom-designed with most imaging software. However, pseudocolouring should not be over-interpreted. Reference to the pseudocolour scale may give the impression that individual colours or hues always represent specific values of ion concentration. This is usually incorrect due to the random variation in pixel values inherent in images (Section 12.2) and the fact that LUTs are often stepped such that a range of pixel values correspond to a given colour. An alternative to pseudocolouring which is commonly used to draw attention to spatial differences in images is three-dimensional graphical representation, in which pixel intensity is coded as height to give a threedimensional surface (36). This is, however, only available with some software packages (such as NIH Image). Smoothing, averaging, and median filtering makes noisy images look more even and is useful for reducing distracting random 'speckle' for qualitative examination. However, caution should be used because of the way in which filters can affect boundaries, such as the extreme periphery of a cell, or distort highly localized changes in pixel value. Filtering should not be applied before extraction of numerical data for quantitative analysis unless one is fully aware of how the pixel values are mathematically transformed and the likely effect on quantitative results.

13.2 Preparation of digitized figures and plates for publication As ion imaging deals almost exclusively with digital image files it is convenient to avoid traditional photographic processing of figures for publication and work digitally. Protocol 6 describes the preparation of figures and plates in digital form for submission to journals either as hard copy or in digital format saved on disk. When submitting images or composite plates on disk it is 267

Richard M. Parton and Nick D. Read essential to also submit top quality hard copies. This is because variation between the calibration of the monitor/printer of the publisher and the system on which plates were initially prepared can cause deviations from the expected colour shades and the brightness and contrast of grey scale images. It is also useful to supply individual TIFF files of the images. If problems arise with the composite file format or with reproduction of either images or text, the publisher can then remake the plate from the TIFF files and add labelling by reference to the hard copies. Where possible a modular design in composite plates should be used (see Figures 6 and 7) because this can be more easily dissected by the Publisher if rearrangement is necessary for space in the journal. Printers for generating hard copies of figures/plates (e.g. dye sublimation printer, video printer, ink jet printer, or laser printer) vary considerably in the prices of hardware and media and the quality of printouts. We recommend the latest generation of ink jet printers as both the hardware and media are extremely cheap relative to other systems and both grey scale and colour reproduction is good enough for most applications except where the highest quality is required (Protocol 6). Protocol 6. Preparation of digitized figures and plates for publication Note that although Macintosh systems are most commonly used by Publishers, PC compatible computers are still more common amongst researchers. This protocol is based on the use of PC compatible software packages and equivalent Macintosh software packages for particular procedures are given. Equipment • Digital image files for processing (e.g. BioRad PIC files collected using the Bio-Rad MRC600 CLSM and COMOS 7.0 software) • Personal computer with sufficient RAM (> 8 Mb) and processor power (pentium 120 MHz or better) to handle image data. A large hard drive (500 Mb-1 Gb) or archiving system (e.g. a magneto optical disc drive or writable CD ROM) is advisable for storing images. • Hewlett Packard Desk Jet 850 colour ink jet printer and Hewlett Packard (HP) Premium Glossy paper

• Software for image processing and additional functions. The packages used routinely by us are: Microsoft PowerPoint 7.0; PaintShop Pro 4.0 (JASC Inc.); Core/draw 3.0 (Corel Corp.); Confocal Assistant 4.02; Optimas 5.0 (Optimas Corp.); Fig. P 2.7 (BioSoft). (Roughly equivalent Macintosh programs are: Adobe Photoshop 3.0; Adobe Acrobat Pro 2.0; Adobe Pagemaker 6.0; Adobe Illustrator 6.0; NIH (National Institutes of Health) Image and Sigma Plot (Jandel), respectively.)

Method 1. Open the Bio-Rad PIC file in Confocal Assistant 4.02, then open the desired colour look-up table (LUT), resave and convert to tagged image file format (TIFF) which is the preferred format for image handling (use the RGB variety of colour TIFF).

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6: Calcium and pH imaging in living cells 2. Import the TIFF file into PaintShop Pro 5.0. The TIFF file may then be cropped to size, rotated, or resized as desired. Scale bars may be conveniently added into the TIFF file at this stage. Digital images are generally collected at a limited resolution of (between 512 x 768 to 1024 x 1024). To improve this, for high quality reproduction, TIFF files should be saved at high resolution in Paint Shop Pro 5.0 (save as: options; set 'dots per inch' DPI to 1400). For submitting digital images on disk to a journal the resolution required is usually specified by the Publisher. 3. Import the processed TIFF file(s) into Powerpoint 7.0. 4. To produce a composite plate which contains line drawings and graphs, these should be prepared using suitable software (e.g. Coreldraw 3.0 for line drawings and Fig. P 2.7for graphs) and imported into Powerpoint 7.0. 5. Arrange images, diagrams, and graphs and add labelling. 6. Produce high quality hard copies using the Hewlett Packard Desk Jet 850 colour ink jet printer on HP Premium Glossy paper. The best grey scale and colour reproduction is obtained when the printer has been calibrated specifically for the attached computer and monitor.

14. Combining ion imaging with other experimental techniques Besides combining imaging with simple treatments involving the external application of different environmental stimuli, agonists, inhibitors, and ion chelators it is often highly desirable to also combine these approaches with more sophisticated experimental techniques or other experimental methods for the measurement of ion concentration or ion channel activity. Important examples include: (a) The use of caged compounds (78). A caged compound is usually a biologically important ion or molecule (e.g. Ca2+, H+, cAMP, inositol triphosphate, Ca2+ buffers, or neurotransmitters) bound to a photolabile 'cage' which masks its biological activity. Irradiation of the cage with UV light (photolysis) releases the bound species in its active form. Cage photolysis is usually achieved with UV light irradiation from the fluorescence excitation source or with specialist UV flash photolysis systems (18). Alternatively, irradiation with the UV or IR laser of a UV CLSM or multiphoton laser scanning microscope (Sections 7.3 and 7.4) can allow highly localized photorelease of caged compounds. Caged molecules may be loaded into cells by similar means as used for fluorescent dyes (Section 5): microinjection or electroporation of cell impermeant forms; 269

Richard M. Parton and Nick D. Read ester loading of the AM ester forms (e.g. nitr-5-AM or diazo-2-AM); and, in the case of the cell permeant caged cyclic nucleotides, by direct diffusion. Caged compounds provide a relatively non-invasive means to manipulate intracellular signalling processes in a very specific and controlled manner which is often highly localized in time and space within living cells. Although intracellular Ca2+ or H+ are often imaged in combination with caged probe release, UV excited ion-sensitive dyes (e.g. Fura-2 and Indo1) are generally not used for this purpose because of the possibility of dye photobleaching during cage photolysis. (b) Microinjection of molecules (such as calcium ion buffers) (25). Pressure injecting the calcium buffer BAPTA has been used to collapse intracellular calcium gradients to investigate their role in tip growth (25). A range of other molecules can also be introduced into cells by pressure microinjection in order to experimentally manipulate signalling pathways, including proteins such as calmodulin. (c) The manganese quench technique (79). Manganese ions (Mn2+) can act as a surrogate for Ca2+ and are taken up into by cells through calcium channels. Unlike Ca2+, however, Mn2+ causes fluorescence quenching of the Ca2+-sensitive dye Indo-1 upon binding to it. Non-inhibitory concentrations of Mn2+ (~ 200 uM) are applied externally to Indo-1 loaded cells and changes in fluorescence are imaged at the Ca2+-insensitive emission wavelength (450 nm) over time. This allows the occurrence of localized Ca2+ channel activity in the plasma membrane to be mapped and measured. (d) Simultaneous electrophysiological measurements (69). It is possible to simultaneously measure membrane potential and/or ion channel activity and intracellular ion concentrations whilst imaging intracellular pH or Ca2+ ion concentration. This combination of electrophysiological and imaging techniques can provide invaluable information on the interrelationship between signalling processes. The main problem with such an approach is in the elimination of electrical interference, which originates from the imaging system and external sources, and which adversely affects electrophysiological measurements. A wire mesh Faraday cage needs to be fitted round the microscope for this purpose (35). Autofluorescence from the microelectrodes or patch pipettes used for electrophysiological recordings can also cause problems and needs to be taken into account.

15. Critical controls for intracellular ion imaging It is appropriate to conclude by summarizing the most critical controls which should be carried out routinely as part of any ion imaging procedure: (a) Intracellular dye distribution. Bright-field, phase-contrast, or DIG microscopy should be used to image dye loaded cells allowing dye distribution 270

6: Calcium and pH imaging in living cells to be mapped and correlated with cytological features. Ideally, confocal fluorescence microscopy should be used to assess the extent to which dye is sequestered within organelles during the time over which ion imaging is performed. (b) Cell health. Although cells may remain viable throughout an imaging procedure, their normal activities may be significantly perturbed. It is, therefore, important to determine whether cells behave normally over the duration of an imaging experiment. Following cellular morphology, growth rates, or response times in dye loaded cells during imaging and comparing these with the same parameters examined in unloaded cells provide the easiest measure of cell health. Possible perturbatory effects of culturing cells on the microscope stage (Section 9), the dye loading procedure (Section 5), the presence of intracellular dye, and the cytotoxic effects of dye excitation (especially with laser or UV light) all need to be assessed. (c) Intracellular dye response. Imposing changes on the intracellular ion concentration, as with in situ calibration (Section 12.5 and Protocol 5) is the most common method of checking the responsiveness of intracellular dye. In some cases the dye may become chemically altered (e.g. by photodamage) or irreversibly bound to cell walls or membranes such that ion concentration in the cell compartment of interest is no longer reported. (d) Routine performance checks on the imaging equipment (see Section 8 and Protocol 2). In addition, performing a simple in vitro calibration after each experiment allows systematic errors (such as incorrect setting up of the equipment on a particular day) to be detected.

Acknowledgements Thanks to Dr Mark D. Fricker for informative discussions and advice and also to Sabine Fischer for her help in preparing the manuscript.

References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

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6: Calcium and pH imaging in living cells 43. Legue, V., Blancaflor, E., Wymer, C, Fantin, D., Perbal., G., and Gilroy, S. (1997). Plant Physiol., 114, 789. 44. McNeil, P. L., Murphy, R. F., Lanni, F., and Taylor, D. L. (1984). /. Cell Biol., 98, 1556. 45. Mir, L. M., Banoun, H., and Paoletti, C. (1988). Exp. Cell Res., 175, 15. 46. Bright, G. R., Kuo, N. T., Chow, D., Burden, S., Dowe, C., and Przybylski, R. J. (1996). Cytometry, 24, 226. 47. Obermeyer, G. and Weisenseel, M. H. (1995). Protoplasma, 187, 132. 48. Pena, A., Ramirez, J., Rosas, G., and Calahorra, M. (1995). J. Bacterial., 177, 1017. 49. Nett, W. and Deitmer, J. W. (1996). Biophys. J., 71, 394. 50. Oparka, K. J. and Read, N. D. (1994). In Plant cell biology: a practical approach (ed. N. Harris and K. J. Oparka), p. 27. IRL Press, Oxford. 51. Oparka, K. J., Murphy, R., Derrick, P. M., Prior, D. A. M., and Smith, J. A. C. (1991).J. Cell Sci., 98, 539. 52. Correa, A. and Hoch, H. C. (1993). Exp. Mycol, 17, 1. 53. Tomkins, P. and Lyons, A. (1993). In Fluorescent and luminescent probes for biological activity (ed. W. T. Mason), p. 264. Academic Press, London. 54. Aikens, R. (1993). In Fluorescent and luminescent probes for biological activity (ed. W. T. Mason), p. 277. Academic Press, London. 55. Cinelli, A. R., Neff, S. R., and Kauer, J. S. (1995). J. Neurophysiol., 73, 2017. 56. Keating, T. J. and Cork, R. J. (1994). In Methods in cell biology (ed. R. Nuccitelli), Vol. 40, p. 221. Academic Press, London. 57. Pawley, J. B. (ed.) (1995). Handbook of confocal microscopy, 2nd edn. Plenum Press, New York. 58. Lemasters, J. J., Chacon, E., Ohata, H., Harper, I. S., Nieminen, A. L., Tesfai, S. A., et al. (1995). In Methods in enzymology (ed. G. A. Attardi and A. Chomyn), Vol. 260, p. 428. Academic Press, London. 59. Helm, P. J., Franksson, O., and Carlsson, K. (1995). Pflugers Arch. Ear. J. Physiol, 429, 672. 60. Czymmek, K. J., Whallon, J. H, and Klomparens, K. L. (1994). Exp. MycoL, 18, 275. 61. Piston, D. W., Kirby, M. S., Cheng, H. P., Lederer, W. J., and Webb, W. W. (1994). Appl. Opt., 33, 662. 62. Xu, C. and Webb, W. W. (1996). J. Opt. Soc. Am. B, 13, 481. 63. Gryczynski, I., Szmacinski, H., and Lakowicz, J. R. (1995). Photochem. Photobiol., 62, 804. 64. Morris, S. J. (1993). In Optical microscopy: emerging methods and applications (ed. B. Herman and J. J. Lemasters), p. 177. Academic Press, London. 65. Bolsover, S. R. and Silver, A. (1991). Trends Cell Biol., 1, 71. 66. Silver, A. R., Whitaker, M., and Bolsover, S. R. (1992). Pflugers Arch., 420, 595. 67. White, N. S., Errington, R. J., Fricker, M. D., and Wood, J. L. (1996). J. Microsc., 181, 99. 68. Sheppard, C. J. R., Gan, X., Gu, M., and Roy, M. (1995). In Handbook of confocal microscopy (ed. J. B. Pawley), 2nd edn, p. 363. Plenum Press, New York. 69. Jaffe, D. B. and Brown, T. H. (1994). Microsc. Res. Tech., 29, 279. 70. Felle, H. H. and Hepler, P. K. (1997). Plant Physiol., 114, 39. 71. Webb, R. H. and Dorey, C. K. (1995). In Handbook of confocal microscopy (ed. J. B. Pawley), 2nd edn, p. 55. Plenum Press, New York. 273

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7

Reflection-contrast microscopy J. S. PLOEM, F. A. PRINS, and I. CORNELESE-TEN VELDE

1. Introduction 1.1 Methodology Faure-Fremiet (1) used reflected light in cell biology studies half a century ago. Curtis (2) and Izzard and Lochner (3) performed the first fundamental studies of interference images of living cells on glass surfaces using reflectedlight microscopy. Reflection images were obtained with scanning reflecting microscopy (4, 5). Reflected-light microscopy for the study of living cells (2, 3, 6, 7) has been described as interference reflection, reflection-interference contrast, surface-contrast, and surface-reflection interference microscopy (8). Ploem (9, 10) investigated further optical methods to improve the image contrast in reflected-light microscopy. In collaboration with Leica (11-13), an improved microscope system was developed for reflected-light microscopy. It used an aperture diaphragm with a central stop (creating an annular aperture) in the epi-illumination light path at an aperture plane conjugate with the back focal (aperture) plane of the objective (14). This was combined with epipolarization microscopy using immersion objectives equipped with a quarter lambda plate in their front lens (15). New high aperture objectives were developed for high resolution biological microscopy. This optically improved version of reflected-light microscopy was named reflection-contrast microscopy (RCM).

1.2 Applications With RCM, a considerable improvement of image quality has been obtained which allowed the observation of weakly reflecting microscopic structures. Most of the early applications of RCM were directed towards the observation of living cells (16). A significant improvement of the image contrast in RCM was obtained in 1982. Bonnet observed (unpublished) that oxidized polymerized diaminobenzidine (DABox), the end-product of an immunoperoxidase staining, showed very strong reflectance. This finding was used for a nonradioactive in situ hybridization with peroxidase-DAB as label and RCM as the microscopic detection technique (17, 18). It resulted in the first successful

J. S. Ploem, F. A. Prins, and I. Cornelese-ten Velde detection of a unique gene on a human metaphase chromosome using a nonradioactive method. Hoefsmit (19) described early applications of RCM after immunogold labelling. Routine histological specimens with a thickness of more then one micron cause complex reflections. Thinner sections are not generally cut for routine histological applications because they do not absorb sufficient light to produce satisfactory image contrast in transmitted light microscopy. Cornelese-ten Velde (20, 21) achieved a significant widening of applications in RCM by examining ultrathin (35-70 nm) sections in biology applications. Reflection can be very strong in thin specimens if sufficient difference in refractive index exists between the stained cellular components and their environment. Cornelese-ten Velde and Prins (22) and Prins et al. (23, 24) made the significant observation that many (immunological) markers like (peroxidase generated) diaminobenzidine polymer products (DABox), immunogold silver, and the alkaline phosphatase staining products show strong reflectance in thin layers. Due to the strong reflection of most immunological markers they can be simultaneously visualized with most conventional histochemical stains. This often allows good morphological localization. Recently multicolour RCM detection was developed for in situ hybridization studies with multiple immune markers (25). For some chemical substances (for example biomaterials) introduced into cells no histochemical stains are yet available. RCM can, however, sometimes visualize such substances if they have a refractive index that differs sufficiently from that of their immediate environment in the cell.

1.3 Review articles Westphal (26) discussed the fundamentals of image formation in classical reflection microscopy for biological applications. Early developments of RCM are reviewed by Pluta in his handbook 'Advanced light microscopy' (8). Verschueren (16) reviewed reflected-light microscopy in studies on live cells. An extensive analysis of RCM methodology and image formation has been published by Cornelese-ten Velde (20), and combined with applications of RCM in a thesis (21). More immunocytochemical applications using ultrathin sections were made by Prins et al. (23, 24). Ploem et al. (27, 48) published recent (1995, 1997) general reviews of RCM.

2. Optical systems for RCM 2.1 Early developments in reflected-light microscopy In conventional reflected-light microscopy, scattered light arises at the upper and further lens surfaces of the objective. This causes glare resulting in a reduced image contrast. In transparent biological specimens, light reflects also from the underside of the microscope slide if this slide is not oiled to a condenser. To reduce glare due to scattered light in reflected-light microscopy of 276

7: Reflection-contrast microscopy opaque objects like coal, Stach (14) designed an aperture diaphragm provided with a central stop creating an annular aperture. These are inserted in the incident light path at a plane conjugate with the back focal (aperture) plane of the objective. Piller (15) introduced the 'antiflex' method for reflected-light microscopy of coal. To that purpose such an 'antiflex' objective is provided with a quarter wave plate in the front lens of the objective. Linearly polarized light is led into a vertical illuminator and is then deflected downwards by a semi-transparent beam-splitter mounted above the objective at 45° to the optical axis. Scattered light reflected upwards from the lens surfaces is blocked by an analyser inserted above the objective, which is crossed with regard to the polarizer. However, light reflected downwards by the beamsplitter transmits the objective towards the surface of the microscope specimen and is reflected back into the objective, passing the quarter lambda plate twice. The polarization plane of this light changes 2 X 45° = 90°, and thus light originating from reflective surfaces in the specimen can pass the analyser and go on towards the eyepieces. Zeiss (Oberkochen and Jena, Germany) manufactured the first 'antiflex' (low power) oil immersion objective for opaque specimens (e.g. coal). Ploem (9, 10) in collaboration with Leica (11-13), integrated both these optical methods—used in coal petrology—into a system for high resolution reflected-light microscopy in biological applications (Figure 1 a). To this purpose, new high power oil immersion objectives provided with quarter lambda plates in the front lenses (suited for high resolution biological and medical microscopy) were developed. For practical purposes, the optical set-up for RCM was incorporated in an epifluorescence microscope equipped with a high pressure mercury lamp for incident illumination. An extra relay lens was mounted in the incident illumination light path (directly in front of the lamphouse) creating an extra aperture plane conjugate with the back focal (aperture) plane of the objective. In this plane, special diaphragms with a central stop creating an annular aperture (precisely adapted to the aperture of the chosen objective) could be inserted. One of the fluorescence filter blocks of the fluorescence epi-illuminator is then replaced by a polarizing block enabling epipolarization microscopy, which is needed as a part of the RCM optical system. Such a microscope can then be used for both fluorescence, epipolarization, and reflection-contrast microscopy. Oil immersion objectives provided with quarter lambda plates in the front lens were made for RCM by Leica (X 40, later X 50 and X 100 oil immersion RCM objectives). Zeiss made an X 63 antiflex oil immersion objective for RCM. These objectives, constructed as non-infinity optics are no longer manufactured.

2.2 New developments in reflection-contrast microscopy Recently (1997, Ploem and Prins, personal communication) have developed another optical method for RCM. A 'normal' X 100 oil immersion objective 277

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Figure 1. (a) Schematic representation of light path in RCM using an RC objective with a quarter lambda plate in the front lens, (b) Schematic representation of light path in RCM using a 'normal' objective and a Wollaston prism above the objective, (c) ZeissAxioskop research microscope for incident-light microscopy, (dl Leica DMR research microscope for incident-light microscopy. (e) RCM diaphragm module with different annular aperture diaphragms inserted in Leica microscope stand, (f) Polarizer block.

(without a quarter lambda plate in its from lens) is used, hut a suitahlc Wollaston prism corresponding to the x 100 objective has to be inserted directly above the objective. In practice a Wollaston prism as used for differential interference contrast (DIC) can be used. In principle the Wollaston prism in an epi-illumination set-tip achieves a similar function to that of the quarter lambda plate, i.e. turning the polarization direction of the rays reflected from

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/. S. Ploem, F. A. Prins, and I. Comelese-ten Velde the specimen is such a way that this light can pass the analyser towards the eyepieces. As with using the special RCM objective, polarizer block and a central stop in the diaphragm module in the the epi-illumination light path are required (see Section 2.3.1). In principle (Figure 1b) the light, linearly polarized by the polarizer, and reflected towards the specimen by the optically flat reflector (beam-splitter), is split by the Wollaston prism into pairs of rays vertically polarized to each other. The image splitting effect is smaller then the resolving power of the objective. After reflection from the surface of the specimen both part-rays are recombined in the Wollaston prism. They then can pass the analyser towards the eyepieces. The results with oil immersion objectives with lower magnifications (X 63, X 40) in combination with their corresponding Wollaston prisms do not give satisfactory RCM results. Since RCM is especially directed at maximal optical resolution, a X 100 objective is the best choice. With most of the very thin and very flat (plane) specimens selected for RCM, practically no inclined surfaces are present and therefore interference phenomena due to phase differences in the reflected beams do not dominate the microscope image. Of particular interest for RCM is the possibility of a better differentiation between metallic tracers used in immuno marking like gold and silver enhanced gold particles and a cytochemical counterstaining used for morphological orientation in the specimen. In research microscopes the Wollaston prism is mounted in a turret above the objective, and can be shifted by turning the turret slightly. As a result reflectance of a counterstain can often be reduced (or enhanced) about twofold. In RCM applications where immunogold and silver enhanced immunogold staining are used in combination with (non-depolarizing) morphological counterstaining or immunostaining (peroxidase, phosphatase), the visibility of gold and silver particles (especially when only a few particles are present) can be significantly enhanced. Shifting the prism quickly back permits again morphological orientation based on the counterstaining. The effect is especially seen with rather strong counterstaining. With weaker counterstaining the effect is less marked. Moving the (DIC) turret to a position for bright-field microscopy (no Wollaston prism in the path) the gold and silver enhanced gold particles can be selectively visualized. There may however be a loss of intensity (of about twofold) of the reflectance signal but weakly depolarizing stains such as immunoperoxidase, immunophosphatase, or morphological counterstaining are not clearly visible. Since the RC central stop remains in the illuminating light path the image background remains relatively dark.

2.3 Modern reflection-contrast microscopes 2.3.1 Upgrading a fluorescence microscope for RCM It is quite simple to equip a modern epifluorescence research microscope (Figures 1c and 1d) with the few extra parts needed for RCM: 280

7: Reflection-contrast microscopy (a) An oil immersion RCM objective with a quarter lambda plate in the front lens; or a 'normal' X 100 oil immersion objective with a corresponding Wollaston prism mounted above the objective. Usually a Wollaston disk corresponding to the X 100 objective (as available for differential interference contrast) is accommodated in a turret in the objective nosepiece. (b) A diaphragm module for RCM (Figure 1e) to be inserted (Figure Id, arrow) in the incident illuminating light path. This module allows the introduction of a sliding set of annular aperture diaphragms. (c) A polarizer block is placed in the epifluorescence illuminator (Figure 1 f ) . When thus equipped for reflection-contrast microscopy, the microscope can also function for transmitted light (absorption) and fluorescence microscopy. These three microscope methods can be used sequentially and partly simultaneously for the same specimen (Protocol 1). (d) A protective filter in front of the lamp is required to protect the polarizer (consult manufacturer's manual), if the microscope is exclusively used for RCM. If the microscope is mainly used for fluorescence microscopy and only infrequently for RCM such a protective filter will interfere with fluorescence excitation light and should not be mounted. Protocol 1. Adaptation of a fluorescence microscope for reflection-contrast microscopy Equipment • Fluorescence incident-light research microscope stand with potential for insertion of a RCM module: Leica DMR research microscope for incident-light (fluorescence) microscopy (Figure 1d) or Zeiss Axioskop research microscope for incident-light (fluorescence) microscopy (Figure 1c) (arrows indicate position of RCM module insertion) • Lamphouse with a high pressure mercury or xenon lamp as used in fluorescence microscopya • Protection filter for the polarizer (the polarizer can be damaged by long exposure to intensive light) • Polarization block, Leica or Zeiss, contains polarizer, neutral reflecting mirror, analyser • RCM diaphragm module, Leica (Figure 1e) or Zeiss, adapting sliding sets with central stops and/or aperture diaphragms to be inserted in the incident light path

Fluorescence illuminator with three to five exchangeable filter blocks for epifluorescence microscopy; one block must be replaced by the polarization block (Figure 1f) • RCM oil immersion objective equipped with a quarter lambda plate in the front lens system: Leica n-plan oil immersion x 100/1.25 RCM objective (infinity corrected optics) or Zeiss plan-neofluar oil immersion x 63/1.25 antiflex objective (infinity corrected optics) • Alternatively a 'normal' oil immersion objective: Leica x 100 PL, fluotar 1.30. The corresponding Wollaston prism is often mounted in a turret above the objective (as available for differential interference contrast). Lower power oil immersion objectives with their coresponding Wollaston prisms do not give satisfactory RCM results.

Method Note: be careful to observe manufacturer's safety warnings. 1. After removing the lamphousing (Leica 105z or 106z), slot the protection filter for the polarizer into the light exit window of the lamphousing from the front. 281

/. S. Ploem, F. A, Prins, and I. Cornelese-ten Velde Protocol 1. Continued 2. Exchange a filter block in the epifluorescence illuminator with a polarization block. 3. Introduce the RCM module into the space reserved for it in the epiillumination light path of a research microscope stand with provisions for incident illumination (research epifluorescence microscope stands of recent design have such provisions). 4. Introduce the sliding units with the various central and aperture stops into the RCM module and adjust the central stop and the aperture stop according to the manufacturer's instructions for RCM. Centre the field diaphragm. 5. Choose the size of central stop in the RCM module in accordance with the RCM objective. 6. Centre the incident illumination carefully (see Protocol 2), by using the adjusting screws on the lamphousing (see the manufacturer's manual for high pressure lamps). Adjusting is easier when a Bertrand lens is used. aMercury high pressure lamps (150 or 100 W) emit strong emission lines dominating the reflectance by the specimen. Xenon lamps (75 W) emit an almost continuous spectrum and provide 'true' colours.

2.3.2 Light sources In epifluorescence microscopy high pressure mercury and xenon lamps are used and these are also suitable for RCM. In fluorescence microscopy, the light source has to provide light in the wavelength range absorbed by the fluorochrome. The wavelength of the emitted fluorescence light differs from the wavelengths of the absorbed light. In RCM, however, light upon reflection at the specimen does not change its wavelength. High pressure mercury lamps have a spectrum with the emitted energy concentrated into peaks. These strong mercury lines at, for example 365, 405, 436, 546, and 578 nm, dominate the reflection by the specimen. Xenon lamps emit a more continuous spectrum. With this light source the colours reflected by the specimen in RCM are then not dominated by strong emission peaks. Therefore, they more truly represent the typical spectral reflecting properties of a particular histochemical stain (see Section 3.1.3). This is especially important if multiple immunohistochemical markers are used in the same specimen. They can only by recognized by differences in the colour they reflect. For specimens stained with multiple stains, a xenon lamp can be of advantage. If only one marker is used for the staining of a specimen, the mercury emission peaks do not cause a problem. Often the great intensities of such emission peaks result in a stronger reflection signal. Most epifluorescence microscopes are usually equipped with high pressure mercury lamps. When such microscopes are upgraded for RCM, one 282

7: Reflection-contrast microscopy usually continues to use the mercury lamp. The microscope can then be effectively used for both fluorescence and reflection-contrast microscopy. 2.3.3 RCM module for the insertion of central aperture stops Modern research microscope stands for reflected-light microscopy allow the insertion of an RCM module in the incident illumination light path. The RCM module is inserted at an (aperture) plane, conjugate with the back focal (aperture) plane of the objective (Figure 1e). The Leica RCM module allows the insertion of two sliding holders close to each other. One with central stops of various sizes and one with different (aperture) diaphragms. Depending on the aperture of each type of objective, a certain size of central stop and aperture must be inserted, to obtain an optimal image contrast. The aperture diaphragm determines the filling of the objective aperture (back focal plane of the objective) and thus the angle of the cone of light projected by the objective onto the specimen (28). With biological objects, which are generally quite transparent, most of the incident light traverses the slide. It then reflects at the lower side of the glass microscope slide, where a large change in refractive index occurs (if no condenser is oiled to the slide). Central stops reduce this unwanted scattered light considerably. A detailed discussion of optical conditions for reducing unwanted scattered light in reflection-contrast microscopy is given by Cornelese-ten Velde et al. (29). Protocol 2. Alignment of the lamp and annular aperture diaphragms in RCM module Note that exact alignment of the lamp and the central stop are critical for RCM. 1. Centre the lamp (XBO-75) or (HBO-100/150) according to the manufacturer's fluorescence microscope manual. 2. The projected image of the central stop must be centred in the middle of the image of the arc of the lamp using the Bertrand lens of the research microscope. 3. The image contrast of the specimen can be optimized by adjustment of the collector of the lamphousing. 4. Always realign the arc and the central stop when the lamp is replaced.

2.3.4 Polarizer block in fluorescence epi-illuminator Most fluorescence microscopes are equipped with an epi-illuminator containing four or more exchangeable filter blocks (cubes). Each block contains an excitation filter, a dichroic mirror, and a barrier filter for excitation with a certain wavelength for fluorescence microscopy. Blocks for excitation with 283

/. S. Ploem, F. A, Prins, and I. Cornelese-ten Velde ultraviolet light, blue, or green excitation light are most frequently used. A fourth position may contain a filter block for excitation with violet light or is left empty to facilitate transmitted-light microscopy. In this position of the illuminator, a polarizing block containing a polarizer, a semi-transparent beamsplitter, and an analyser, can be inserted. With this choice of blocks the microscope can be used for either fluorescence or reflection-contrast microscopy of the same specimen. (The protection filter for the polarization block absorbs some excitation light wavelengths needed for fluorescence microscopy and must be removed from the lamphousing. For short periods, however, the polarization block would not be damaged and could be used unprotected by this filter.) 2.3.5 Wollaston prism in a DIC turret above the objective For RCM with 'normal' objectives without a quarter lambda plate in the front lens (or Wollaston prism in the objective) a Wollaston prism, corresponding to the oil immersion objective chosen must be turned in the light path for vertical illumination. An aperture diaphragm with a central stop must be inserted in the RCM diaphragm module in the incident illuminating light path, corresponding to the aperture of the objective (see Section 2.3.3). The polarizer block in the vertical illuminator must be turned into its position in the epiillumination light path. Shifting the Wollaston prism by slightly turning the DIC turret will enable some differentiation between depolarizing and nonpolarizing stains (see Section 2.2). 2.3.6 RCM oil immersion objectives When using RCM oil immersion an aperture diaphragm with a central stop must be inserted in the RCM diaphragms module in the incident illuminating light path, corresponding to the aperture of the objective (see Section 2.3.3). A rotating collar around the objective allows the turning of the quarter lambda plate in the front lens of the objective. For RCM, the quarter lambda plate should be rotated to maximize the reflection from the specimen, at which position the polarization angle of the reflected light is at its optimum for passing the analyser, and RCM is performed. Epipolarization microscopy is obtained when the quarter lambda plate is set to minimal reflectance. Protocol 3.

Focusing on small and thin specimens on microscope slides

Note: because the specimens selected for RCM are often thin and small, focusing in RCM is a very delicate procedure. An additional difficulty is that all areas outside the specimen boundaries are observed as dark. For this reason the instructions below should be followed step for step. Marking the site of the specimen is necessary. The use of a low power dry objective (X 2.5 magnification) for locating the specimen is essential for easy RCM.

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7: Reflection-contrast microscopy Equipment • Microscope stand (Leica or Zeiss) with accepts the appropriate central stops and aperture diaphragms for RCM. On the revolving nosepiece an RCM objective and a very low power dry objective (for example x 2.5) should be available.

• A marking pen with dark permanent ink

Method 1. Circle marking with a pen closely around the specimen on the nonspecimen side of the slide. 2. Put a drop of immersion oil on the marked area of the specimen. 3. Mount the microscope slide with the specimen on the stage. 4. Swing the low power objective into observing position (keep free of the immersion oil on the specimen). 5. Search for the marking made on the lower side of the microscope slide and focus on this marking. The field diaphragm should be simultaneously visible in the field of view. If you cannot see it clear enough, move the handle of the field diaphragm in the RCM module up and down till you notice the field diaphragm. 6. Adjust the field diaphragm in the RCM module to about 75% of the microscope field by moving its handle. 7. Lower the microscope stage quite a bit by using the coarse stage movement. 8. Swing in the RCM oil immersion objective into observing position without, however, touching the immersion oil on the specimen (since no coverglass is used the objective might damage the specimen by swinging in at too low a level). 9. Raise the stage until the RCM objective just dips into the immersion oil (look from the side of the microscope stage). 10. Move the objective further into the immersion oil with the fine focus control (but avoid the spring-loaded front lens of the RCM objective being pressed inwards). Carefully focus, through the eyepieces onto the projected image of the field diaphragm (narrowed before to a size of 75%). If no image of the field diaphragm is observed, lower the microscope stage (but without losing contact with the immersion oil), using the fine focus controls until an image of the field stop is obtained. 11. After obtaining an image of the field diaphragm, stop focusing. You are now in the plane of the specimen. If step 1 of this protocol is carried out properly, you are within the marked area of the slide and should find the specimen by moving the stage slowly in various directions.

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/. S. Ploem, F. A. Prins, and I. Cornelese-ten Velde Protocol 3.

Continued

12. After microscopic examination the slides should be removed and stored vertically in a microscope slide box, to allow the oil to drain away from the slide. (Prevent dust collecting on the slides since this might strongly disturb the RCM examination of the specimen on the next occasion.) 13. For microscopic re-examination of stored slides, first remove the immersion oil still remaining on the specimen with 100% ethanol and let dry. Bring a fresh drop of immersion oil on the specimen before RCM.

3. Image formation 3.1 General 3.1.1 Reflection of light waves For a detailed discussion of image formation in reflection-contrast microscopy, the reader should consult Cornelese-ten Velde (20). The reflection of light at an interface with a given refractive index n and an absorption coefficient k is described by Fresnel equations. The physical properties n and k of stains can be influenced by the pH, the salt concentration, their conformation (crystalline or amorphous precipitates), and by chemical binding or electrostatic forces that may occur during the preparation. In image formation of (stained) microscopic structures in reflection microscopy, three main types of optical phenomena and their combination can be observed: (a) Interference of reflected light waves due to the presence of two or more reflecting interfaces. (b) Selective reflection of certain wavelengths by a substance, due to its high refractive index for certain wavelengths. Both interference of reflected light (only differences in n) and selective reflection (also contribution of k) may occur in the same specimen. (c) Depolarization of reflected light from metal surfaces (high value for k). 3.1.2 Interference of reflected light waves By reflection at an interface (Figure 2a), a part of the light wave (I0) is reflected (Ir) and the remainder is refracted (It) on transmitting the interface (I). Figure 2. (a) Diagram of multiple reflections in a thin layer system. It illustrates the computation of reflectance. $i = angle of incidence; or = angle of refraction; I0 incident light; lr reflected light. (For other symbols see Section 3.1.2.) (b) Schematic drawing of light rays reflecting at a cell attached to a glass substrate in an aqueous medium (after Pluta). Rgm: reflection at glass medium interface. Rgc: reflection of glass-cell contact. (c) RCM image of living cells viewed through the coverglass (the objective is beneath the coverglass) to which cells of the kidney tumour cell line SK-RC-52(Gural) are attached. Bar = 10um.

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287

/. S. Ploem, F. A. Prins, and I. Cornelese-ten Velde At the following parallel interface (II) the transmitted (refracted) component reflects a part (Itr) and transmits (Itt) the rest again. The reflected part of the component ray (Itr) at interface (II) will also reflect and transmit (Itrt) interface (I), and so on. The first reflected part (Ir) of the incident wave can interfere with the transmitted part of Itr ( = Itrt) and so on, because they are components of the same wave (same frequency), they have a phase difference, and vibrate in the same plane (20). Interference of reflected light waves gives information about distances between reflecting layers (e.g. glass-cell surface) if they are in the wavelength range of visible light. Interference of reflected light plays a dominant role in the RCM of unstained specimens such as living cells. Interference fringes mostly dominate these images. In thin tissue sections generally only a few fringes are visible. In unstained thin sections, when using a xenon lamp a weak colour image can often be observed. This is caused by interference of reflected light waves and depends on the thickness of the section. Interference phenomena can also occur simultaneously with selective reflection, for instance when a dye is present in a layer that produces interference of selective reflected light rays. Both account for image formation by the DABox immunocytochemical marker (20) (see Section 3.3). There are two methods to discriminate between these two phenomena, namely to use incident light of various wavelengths or to use different illuminating numerical apertures (28) (see Section 3.1.3). 3.1.3 Selective reflection Selective reflection can highlight stains (substances) with special reflecting properties in the image (mostly more intensively than in transmitted-light microscopy). The resulting enhanced image contrast can simplify the location of the cytochemical staining. It helps the localization of exceptionally fine structures and very low density in antigen binding sites. Selective reflection of light can be defined as reflection of certain wavelengths by a substance. It is due to high absorption and a high refractive index of the substance for those wavelengths. In reflection microscopy, stained specimens exhibit generally the complementary colour of the conventional transmitted-light image, e.g. a blue stained structure in transmitted light (due to the absorbance of red wavelengths), is seen as red in the reflected light image. This is caused by the reflection of a part of the red incident light at the interface with the stained structures. Selective reflection is the more dominant reflection mode with stained specimens (30). Because of the relative complex image formation in reflected-light microscopy, it is always important to investigate the nature of the reflected light, to avoid erroneous conclusions such as supposing fluorescence instead of reflection (30, 31). By changing the wavelength of the incident light and the illuminating numerical aperture (28), Opas and Kalnins (32) could, for example, verify that the stain Coomassie brilliant blue showed mainly selective reflection. 288

7: Reflection-contrast microscopy 3.1.4 Depolarization Depolarization of reflected light can occur with metals. Metal objects in cytochemical applications (e.g. silver enhanced immunogold particles) have up till now usually been visualized with reflected-light polarization microscopy (epipolarization microscopy). The quarter wave plate of an RCM objective can be rotated to minimal achieved brightness of the image. In this situation the optical configuration of epipolarization microscopy is set. When, however, the quarter wave plate of the RCM objective is turned to maximal brightness of the image, the optical setting for RCM is obtained. About a twofold increase in intensity of the light reflected by the silver enhanced immunogold particles is then achieved (29).

3.2 Image formation in RCM of living unstained cells The image formation with cells growing on a glass slide has been reviewed by Pluta (8). In Figure 2b, a schematic representation is given of a vertical section through a living cell on a coverglass. When a thin medium layer (e.g. under the cell) or a thin layer of cytoplasm is present, reflections at both the upperand underside of these layers occur. When the thickness of the layer is in the order of the wavelengths of visible light, dark and light bands are seen in the microscope image. These are caused by interference of reflected light waves. Relatively bright areas are observed at the glass-medium interface where a large difference in refractive index exists resulting in a strong reflectance of light. Darker areas are observed at glass-cell surface contacts. A smaller difference in refractive index exists here, causing only a weak reflectance of light (Figure 2c) (2, 3, 6, 7, 33-35). The influence of conical illumination and the size of the aperture diaphragm has been investigated to estimate the thickness of cellular structures in living cells (28).

3.3 Image formation in RCM of stained specimens Before about 1985, mostly non-immunological stains were investigated. Pera (36, 37) studied haematoxylin/eosin stained histological and cytological sections. He noted that when the quarter wave plate of the RCM objective is turned to obtain maximum brightness of the image—as should be done for RCM—an enhanced staining contrast was obtained. Weakly stained cellular structures, which were only just recognizable in conventional microscopy were clearly visible with RCM. With histochemically stained thick histological sections the image information can be quite complex. With RCM of cytochemically stained thin specimens often a strong image contrast is obtained. They are often observed against a dark background. Sometimes this may mimic fluorescent images. Van der Ploeg and van Duijn (30, 31) conclude that some observations interpreted as fluorescence of stained structures reported in the literature (38), were due to selective reflection of light. Cornelese-ten 289

/. S. Ploem, F. A. Prins, and I. Cornelese-ten Velde Velde et al. (20) used a model system to study the image formation of DABox. They showed that the reflection of DABox depends primarily on its thickness, which was estimated at 50-100 nm for sensitive detection of DABox by RCM. From this fact new applications arose: specimens with the stain (DABox) present in a very thin layer would yield optimal images with RCM. These images were obtained from ultrathin epon sections containing DABox and immunoperoxidase stained Lowicryl sections (20). Pre-embedding is nowadays often used for the detection of surface antigens (or extracellular matrix components) with the PO-DAB as detection method (23). It may be useful to compare the image contrast obtained with RCM with that obtained by conventional transmitted-light microscopy of the same tissue section. The inverted contrast in RCM (bright label against a dark background) cannot however be easily compared with the transmitted-light image (dark label against a light background). Many microscopes are however linked to a TV camera and connected to a PC with graphic software for the treatment of digitized microscope images. It is then possible to invert (make negative) the RCM image. When such an inverted RCM image is then compared with the original transmittedlight image of the same (semi-thin) tissue section, it becomes clear how large the gain obtained with RCM is in detecting the immunolabel (Figure 3). Inverting the image is also possible by photographic manipulating, by making prints from black and white reversal film or colour diapositives.

4. Applications 4.1 General Cyto- and (immuno)histochemistry are important applications in biology and medicine. Immunohistochemistry studies using transmitted-light microscopy are sometimes hampered by poor morphological detail and low contrast of labels in paraffin or cryosections. Bright-field routine sections for histopathology are generally cut relatively thick (3-6 um) to obtain sufficient image contrast. When relative thick specimens are used some layers of the specimen are, however, not in focus when a high power objective is used. Thinner sections would have been desirable, since practically no out-of-focus layers of the section would then perturb the image. However, thinner sections lack sufficient image contrast in transmission microscopy. Although relatively thick specimens can be examined with confocal microscopy since this optical method eliminates out-of-focus information and examines only a thin layer in the specimen. Very thin sections can be examined with reflection-contrast microscopy since reflection is a surface phenomenon. The microscope images obtained with RCM from ultrathin (30-70 nm) sections have a large image contrast and are totally in-focus even with high power objectives (Figure 4a). Semi-thin sections (0.25 u.m), however, can provide satisfactory RCM images (Figure 4b). This is important for the routine application of RCM. Most laboratories will 290

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Figure 3. Localization of immune deposits in nephrology. IgG deposits in the glornerulus of a mouse with graft versus host disease-induced lupus nephritis, visualized by an indirect immunoperoxidase-DAB labelling. (a-d) The same 1 um thick section counterstained with Light Green seen by (a) bright-field microscopy (BFM), (b) combined transmitted and incident light (BFM/RCM), (c) only RCM, and (d) inverted computer image of (c). The digitized image of (c) was only inverted, therefore no colour and brightness enhancements were performed. Bar - 10 um.

only have standard microtomes available and only a few laboratories will have ultramicrotomes. Most modern motorized microlomes (especially when equipped with a glass or diamond knife) are very suited to cut semi-thin sections for RCM. Ultrathin sections, however, give superior images. Like fluorescence microscopy, RCM provides images with a large image contrast between the immune label and the tissue background. However, unlike fluorescence microscopy, it also shows tissue background reflections (Figures 5 and 6). Since fluorescent markers can fade, the lifetime of such specimens can be limited. With RCM, fading of the cytochemical label is 291

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(b) Figure 4. RCM image of (a) ultrathin section and of (b) semi-thin. (a) Ultrathin LR White section of a rat kidney stained with haematoxylin. (b) 0.25 urn thick LR White section of a rat kidney stained with haematoxylin. Bar = 10 uM.

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Figure 5. RCM image of binding of PO labelled cMOv 18 on HeLa cells. After development of PO with DAB, the cells are embedded in epon (pre-embedded) and ultrathin sectioned. Bar = 10 um.

Figure 6. Localization by RCM of immune deposits in nephrology. IgG deposits in glomerulus of a mouse with graft versus host disease-induced lupus nephritis, visualized by an indirect immunogold labelling, Ultrathin Lowicryl section mounted on a gelatin coated slide, stained by rabbit anti-mouse IgG, followed by 15 nm colloidal goldconjugated goat anti-rabbit IgG. Bar = 10 um.

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/. S. Ploem, F. A. Prins, and I. Cornelese-ten negligible, so that specimens can be stored and re-examined several times. With a high image contrast and no deterioration of the image by out-of-focus image formation, RCM of ultrathin sections fulfils the theoretical optical criteria of optimal light microscopic resolution, For some substances (e.g. drugs, particles of various materials as for example ceramic material) introduced into cells, no cytochemical staining methods have yet been developed. Detection of such substances by RCM is sometimes possible when such substances have a refractive index that differs sufficiently from their environment in the specimen (Figure 7), In many applications it is important to visualize not only, for example, a specific antigenic site in the cell by using an immunolabel, but also to verify the exact morphological site of such labelled objects in relation to other subcellular structures. Fortunately RCM visualizes all materials in the specimen that have a sufficiently different refractive indices. In addition counterstaining can be carried out to obtain further cytochemical or histochemical information. If such a counterstaining would identify certain other cellular organelles, the precise location of the immune label would then often be easier. The possibility of using multiple simultaneous stains in one specimen depends on: • the differences in the colour of the various stains • the microscopic image contrast obtained The greater the microscopic image contrast, the easier it is to use the difference in reflectance of a label to distinguish that label from another stain.

Figure 7. RCM image of an degradation of implanted biomaterials. Phagocytosed and degradated spheres of injected hydroxyapatite in a giant cell. Ultrathin section. Bar = 10 um.

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7: Reflection-contrast microscopy RCM of most immunolabels result in strong signals. Many immunocytochemical labels can therefore be combined with conventional histo- and cytochemical stains. These should, however, be used in a low concentration to give a reflectance signal that is just sufficient to observe the cellular structures for which the stain is intended. In this way the RCM image looks like an inverted low magnification EM image. The same cell components are visible, but, often with a higher image contrast. Interestingly several fluorochromes, such as the DNA/RNA stain. Acridine Orange, can be also used as a counterstain in RCM, since Acridine Orange, in low concentrations and in a thin layer, gives sufficient reflection of light. With transmitted-light microscopy, it is not always possible to examine very thin layers and small amounts of biological material, or cells grown on, for example, (non-transparent) filters. Such specimens often require EM methods for examination. Since RCM is an incident light method it is possible to examine a great variety of ultrathin sections of specimens (e.g. tissues or cells on a substrate, cultured cells on filters, small biopts).

4.2 Special applications 4.2.1 In situ hybridization studies One early application of RCM, as mentioned earlier, was the non-radioactive in situ hybridization using pcroxidase-DAB as a label, in genetic studies of human chromosomes. It resulted in the first successful detection of a unique gene on a human metaphase chromosome using a non-radioactive method (17, 18). RCM is well suited for in sim hybridization studies (Figure 8) and is

Figure 8. Non-radioactive in situ hybridization of mouse metaphase chromosome spread with biotinylated pUC 1.77 detected with peroxidase-DAB method. Bar = 10 um.

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/. S. Ploem, F. A. Prins, and I. Cornelese-ten Velde Table 1. Comparison of the colour of various enzyme precipitates as revealed by different types of microscopy after enzyme cytochemical detection of DNA probes in situ (25) Enzyme reagentsa

Microscopy Bright-field

APase APase

N-ASMX-P + Fast Red TR Red BCIP + NBT Blue/purple

APase APase PO PO PO PO

N-ASMX-P + New Fuchsin NBT/BCIP/INT H 2O2 + AEC H2O2 + chloronaphthol H 2O2 + DAB H2O2 + TMB

GO

PMS + NBT

Red Brown Red Purple Brown Greenc,d/ purpled Blue/purple

Reflectioncontrast

Fluorescence

Yellowb Red Orange/ yellowb Yellow Red Whiteb Yellow" White/yellow" Whited Redc/yellowc Yellow"

a APase = alkaline phosphatase; PO = horseradish peroxidase; GO = glucose oxidase; AEC = aminoethyl carbazole; BCIP = bromochloroindolyl phosphate; DAB = diaminobenzidine; N-ASMX-P = naphtholASMX-phosphate; NBT = nitroblue tetrazolium; PMS = phenozine-methosulfate; NBT/BCIP/INT substrate, product name (Dako A/S);TMB = tetramethylbenzidine. bFixation of enzyme precipitates in a protein matrix is essential; mounting in immersion oil. c Fixation in a protein matrix is not essential; mounting in immersion oil. d Colour of the reaction product in air dried slides or after mounting in PBS/glycerol.

especially of advantage if the specimens have to be stored to be re-examined later. In applications where RCM in situ hybridization methods are applied to cells, nuclear counterstaining may be of help to provide information about the localization of the specifically stained structures. Multiple reflecting immune labels for in situ hybridization studies have been developed by Speel (25) (Table 1). 4.2.2 Rare event detection with RCM The detection of infrequent objects in medical specimens (e.g. urine and blood) is especially difficult if similarly stained artefacts are also present on the slide. A careful (and time-consuming) visual inspection is needed of all these objects which often show only minor differences in image contrast, colour, or signal intensity between specific and non-specific staining. An example may illustrate this. Microscopic detection of infrequently occurring leptospires in clinical blood samples is difficult. Detection of only a few leptospires, using immunofluorescence and transmitted-light microscopy after immunoperoxidase staining, is complicated by the fact that some artefacts resemble leptospires. With immunogold silver or PO-DAB staining a very strong reflectance signal from the leptospires can be seen with RCM (Gravekamp and Prins; personal communication). Discrimination between leptospires and artefacts is easy because of the large image contrast. In addition the 296

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Figure 9. PO-DAB stained leptospires in a blood-smear. Blood was experimentally infected with leptospires (serovar grippotyphosa). Note the characteristic coiling, normally observed with EM. Bar = 10 um. With routine transmitted-light microscopy such coiling is difficult to observe and rarely occurring leptospires are difficult to find because of a lack of image contrast.

resolution of the microscope image is excellent and characteristic coils of the PO-DAB stained leptospires, (which arc very difficult to distinguish with conventional light microscopy) can be seen in RCM images (Figure 9). 4.2.3 RCM detection of substances with differences in refractive index RCM has been used for example in studies of biomaterials (material for bone implants). The degradation of calcium phosphate ceramic particles was studied in a mouse model system. For RCM ultrathin sections are directly laid on glass slides, strong reflection signals and optimal RCM images can be obtained from biomaterials having sufficiently different refractive indices (Figure 7). This avoids the loss of biomaterial during staining, a problem encountered in processing this material for EM. 4.2.4 Comparison and combination with other microscopic methods RCM has been used in comparison and combination with other microscopic techniques such as: differential interference contrast microscopy, high voltage EM, immunofluorescence microscopy, immunoelectron microscopy, scanning EM (39, 40). phase-contrast microscopy, and total internal reflection aqueous fluorescence (41). 297

/. S. Ploem, F. A. Prins, and I. Cornelese-ten Velde 4.2.5 RCM in combination with electron microscopy A methodology in which an overview of a specimen with a resolution close to the optimal for LM, can be followed with EM examination of the next sequential section cut from the same tissue block, provides promising possibilities. The large image contrast of DABox and immunogold (silver) in RCM makes an overview and statistical analysis of stained structures in a large part of the specimen relatively easy and efficient. RCM images can be directly compared with low magnification EM images (22-24). EM can, of course, extend the RCM observations with higher resolutions in smaller selected areas of the specimen. Such a continuum in observations of the same specimen with a very large range of resolutions, was up till now not easily possible. This is a practical proposition since often the same pre- and post-embedding methods can be used both for light and electron microscopy and can then be processed for either RCM or EM (Figure 10).

5. Specimen preparation 5.1 General Due to the enhanced detection sensitivity of RCM for many substances, this technique is very sensitive to all kinds of contaminants. Contamination by, for example, dust particles or other unwanted substances must be avoided at all cost (Protocol 3). Microscope slides must therefore be carefully cleaned and their surface not touched after they have been cleaned. Precipitates must be removed from staining solutions. All solutions should therefore be filtered before handling the specimens. Coverglasses are not generally used on the microscope specimens for RCM. After microscopic inspection the specimens should be stored in such a way that no dust or dirt can accumulate on them.

5.2 Immunohistochemistry There are three main types of methods for antigen localization in ultrathin sections by RCM. These methods are common in electron microscopy and the protocols are (minor) adaptations of these EM methods. Table 2 gives a scheme of the protocols that could be followed for different immunohistochemical applications. The choice for a particular method depends on the type and location of the antigen. If the antigen is unstable (i.e. antigenicity is lost after fixation), ultracryo methods are preferred. When the antigen is stable, its intra- or intercellular localization determines the accessibility of the antibody and by that the choice for pre- or post-embedding methods. Surface antigens are best detected by pre-embedding methods, whereas matrix antigens (such as collagen, fibronectin) and intracellular antigens are best detected by post-embedding methods. Tissue processing for pre-embedding is described in Protocol 4. Before this procedure is carried out, it is advised to test the 298

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Figure 10. (a) RCM and (b) EM image of linear deposition of rabbit anti-laminin antibodies in rat kidney by indirect immunoperoxidase-DAB staining, Ultrathin epon section of preembedded tissue, (a) Bar = 10 um. (b) Bar = 5 um.

Table 2. Scheme of protocols: immunohistochemical methods for thin sections in RCM (appropriate number of protocol in brackets) Pre-embedding

Post-embedding

Ultracryo sections

Pre-fixation and irnmunostaining [5] Embedding [5] Ultrathin sectioning [8] Counterstaining [8] RCM [2]

Fixation and embedding [6]

Fixation and freezing [7]

Ultrathin sectioning [8] Immunostaining (9] Counterstaining [9] RCM [2]

Ultrathin cryosectioning [7] Immunostaining [91 or Counterstaining [9! RCM [2]

299

J. S. Ploem, F. A. Prins, and I. Cornelese-ten Velde antigenicity remaining on 2 um cryosections to determine the concentration of glutaraldehyde that can be tolerated. Protocol 4. Tissue processing for pre-embedding All solutions must by filtered before use with a 0.2 p,m filter (Schleicher & Schuell, FP030/3) Equipment and reagents Vibratome (Oxford Laboratories, USA) Beem capsules Size OO (Polaron, England) Fixative solutions: 4% paraformaldehyde, 0.1-0.2% glutaraldehyde, 0.1 M phosphate buffer pH 7.3 1% glutaraldehyde, 0.1 M phosphate buffer pH7.3 PBS: 150 mM NaCI, 10 mM phosphate pH 7.3 Quench buffer: 0.5 M NH4CI in 0.1 M phosphate buffer pH 7.3

Wash buffer: 0.1 M phosphate pH 7.3, 3% sucrose PO-conjugated antibody 1% bovine serum albumin (BSA) in PBS Diaminobenzidine medium (DAB): freshly prepared 0.05% diaminobenzidine in 0.05 M Tris-HCI pH 6 1% osmium tetroxide in 0.1 M phosphate buffer pH 7.3 30% hydrogen peroxide

Methoda 1. Fix tissue blocks (c. 4 mm cube) for 3 h at 4°C by immersion in, or perfusion with, fixative solution. 2. Cut 70 n.m vibratome sections of the tissue blocks and store the sections in wash buffer. 3. Incubate the sections with quench buffer for 60 min at 4°C. 4. Pre-incubate 1% BSA, PBS for 30 min at 4°C. 5. Incubate sections for 16 h with antibody in 1% BSA, PBS at 4°C. 6. Wash five times for 10 min with wash buffer at 4°C. 7. Incubate for 2 h in PO-conjugated antibody in 1% BSA, PBS. 8. Wash five times for 10 min in wash buffer at 4°C. 9. Fix with 1% glutaraldehyde for 10 min at room temperature. 10. Wash three times for 5 min with PBS. 11. Add 10 ul hydrogen peroxide to 10 ml filtered DAB solution. 12. Develop for 30 min in the dark with diaminobenzidine medium. 13. Wash twice with PBS for 10 min. 14. Post-fixate with 1% osmium tetroxide for 30 min. 15. Dehydrate and embed in epon 812 (see Protocol 5). For sectioning see Protocol 8. ' Modified from the method given in ref. 42, with permission.

300

7: Reflection-contrast microscopy Ultracryo methods can be chosen if the antigen is unstable (i.e. antigenicity is lost after fixation). The accessibility for the immunological reagents to reach their targets is excellent. Ultracryo methods are also performed when antigen location or antigenicity is not precisely known. These methods have not been developed for conventional light microscopy. RCM, however, offers this possibility. The excellent RCM image definition of ultrathin cryosections, is comparable to that of ultrathin plastic sections. Ultracryo and post-embedding methods are quite similar techniques. However, each has different advantages. Handling of resin sections is easier than cryosections, larger sections can be cut, and sequential sectioning is easier. Material embedded in resin blocks and resin sections can be stored for long periods. Biological material embedded in resin is easy to counterstain. RCM of ultracryo sections is relatively easy to perform and the whole ultracryo procedure is the fastest of the three methods described above. RCM of ultrathin cryosections, in addition, enables the possibility of direct comparison with EM.

5.3 Fixation and embedding Paraffin wax embedding and sectioning are not suitable for RCM. Many routine biological and medical laboratories, however, increasingly employ specimen embedding in plastics such as GMA, LR White, and Lowicryl. For such laboratories only a relatively small additional effort is needed to make extra sections for RCM (see Section 5.4). For embedding in plastics similar— but slightly modified techniques—as those required for electron microscopy are used. A selection of the methods used for the preparation of specimens for RCM is given in Protocols 4-10. For different types of specimens, e.g. cell cultures, biopsies, a variety of fixation methods, pre- and post-embedding in plastic or ultracryo techniques are employed. For further details the reader is referred to practical handbooks on EM techniques. Protocol 5 describes fixation and embedding in epon (43) for mainly morphological studies by RCM. One significant application of RCM of ultrathin sections is the morphological quality control of cell or tissue cultures. Protocol 5. Fixation of tissue and embedding in epon for RCM Equipment and reagents • Razor blade, wax plate, pair of tweezers, Beem capsules, size 3 (Polaron, England) • Fixative solution: 0.1 M cacodylate buffer pH 7.3, 1.5% glutaraldehyde, 1% paraformaldehyde (Merck) . Post-fixation solution: 0.1 M cacodylate buffer pH 7.3, 1% osmium tetroxide (Agar UK, R1017) . Propylene oxide (Fluka, 82320)

• Wash buffer: 0.1 M cacodylate pH 7.3, 3% sucrose • 30%, 50%, 70%, 80%, 90% ethanol in distilled water, and 100% ethanol . DDSA (Fluka, 45345) .NMA(Fluka, 45347) * DMp-30 (Fluka, 45348) • Oven (temperature range 30°C to 100°C)

301

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Continued

a

Method 1. Prepare mixture of 80 ml epon 812, 50 ml DDSA, and 45 ml NMA. Add to 10 ml of the mixture 0.15 ml DMP-30 (accelerator) and mix. 2. Cut the tissue into small pieces << 2 mm2) with a razor blade. 3. Fix for 2 h at 4°C with fixative solution. 4. Wash at 4°C overnight with wash buffer. 5. Fix for 1 h at room temperature with post-fixation solution. 6. Wash three times for 5 min with distilled water. 7. Dehydrate with 50%, 70%, 80%, 90% ethanol in distilled water, and 100% ethanol for 30 min each. 8. Immerse in propylene oxide for 5 min. 9. Infiltrate with epon:propylene oxide (1:1) mixture overnight. 10. Infiltrate with epon:propylene oxide (2:1) mixture for 2 h. 11. Embed in Beem capsules in epon and polymerize at 60°C overnight. * Modified from the method given in ref. 43, with permission.

Protocol 6. Fixation and post-embedding of tissue or cell cultures for RCM Equipment and reagents • Razor blade, wax plate, pair of tweezers, rubber policeman, centrifuge • Conical tip capsules or 1.5 ml Eppendorf tubes and Beem capsules, size 3 (Polaron, England) • Ultraviolet polymerization lamp (Polysciences Europe, No. 8778) • 10% gelatin in PBS at 37°C (for embedding fixed cells) • Fixative solution: 4% paraformaldehyde and 0.1-0.2% glutaraldehyde in 0.1 M phosphate buffer pH 7.3

• Aldehyde residue blocking solution: 0.05 M glycine in PBS . Wash buffer: 0.1 M phosphate buffer pH 7.3 with 3% sucrose LR White (medium 2031) resin (The London Resin Co. Ltd.) • Lowicryl: mix 2 g cross-linker mixing with 13 g monomer and 0.075 g initiator (Lowicryl K4M kit No. 15923, Polysciences Inc.) . 30%, 50%, 70%, 80%, 90% ethanol in distilled water, and 100% absolute ethanol

A. For tissuea 1. Cut the tissue into small pieces (< 2 mm cube) with a razor blade. 2. Fix tissue for 1 h at 4°C with fixative solution. 3. Store tissue for 16 h at 4°C with 2% paraformaldehyde in 0.1 M phosphate buffer pH 7.3. 4. Wash for 2 h at 4°C with wash buffer. 5. Dehydrate with 30% ethanol for 30 min at 4°C. 6. Dehydrate for 45 min through 50%, 70%, 80%, 90%, and 100% ethanol at-20°C. 302

7: Reflection-contrast microscopy 7. 8. 9. 10. 11.

Infiltrate for 45 min with Lowicryl:ethanol (1:1). Infiltrate for 2 h with Lowicryl:ethanol (2:1). Infiltrate in Lowicryl (pure resin). Embed in Beem capsules in Lowicryl (pure resin). Irradiate for 24 h with the lamp-to-capsule distance at 30 cm at -20°C.

LR White embedding (alternative to Lowicryl K4M) 1. Follow part A, steps 1-4 as for the tissue treatment. 2. Dehydrate in ethanol 30%, 50%, 70%, 100%, 45 min each, at room temperature. 3. Infiltrate in LR White:ethanol (1:1) for 1 h. 4. Infiltrate in pure LR White three times for 1 h each. 5. Embed in Beem or gelatin capsules and polymerize at 4°C for 24 h. B. For cell cultures 1. Pour off the culture medium and fix cell cultures for 10 min at 4.C. 2. Scrape from bottle with a rubber policeman. 3. Centrifuge (5 min, 100 g) in conical tip capsules, and mix pellet in 10% gelatin, PBS 37°C. 4. Centrifuge (5 min, 260 g) again and cool to 4°C for 10 min. 5. Treat as for tissue (part A, steps 1-11). aModified from the method given in ref. 44, with permission.

5.4 Sectioning Because for RCM and EM the same specimen preparation method is used, it is possible to obtain subcellular information from the RCM image of a particular site in the ultrathin section (of almost the same tissue structure) by putting a consecutive section on a grid and examining this by EM. Ultracryo microtomy (Protocol 7) in combination with RCM is a recent procedure for optimal immune localization (24). Protocol 7. Fixation and ultracryo freezing and sectioning Equipment and reagents • Ultracryo microtome (Leica) and glass or dry cryo diamond knife . Dako pen (S 2002) and (3 mm) wire loop 9 (Figure 77) • Aminosilane or gelatin coated glass slides (see Protocol 10) . Hotplate (37 °C)

• Fixation solution: 4% paraformaldehyde, 0.1% glutaraldehyde in 0.1 M phosphate buffer pH 7.3 . Wash buffer: 0.1 M phosphate pH 7.3 with 3% sucrose . Cryoprotection solution: 2.3 M sucrose in PBS

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Continued

A. For tissue (fixation and freezing)a 1. Cut the tissue into small blocks (less than 1 mm3) with a razor blade. 2. Fix tissue for 1 h at 4°C with fixative solution. Or store for longer periods in 2% paraformaldehyde at 4°C. 3. Infiltrate blocks in cryoprotection solution for 1 h. 4. Freeze in liquid nitrogen on special (cryo) specimen holder. Sectioning and mounting 5. Cut sections (60-90 nm) on an ultracryo microtome at -120°C with a glass or diamond knife. 6. Transport sections from the knife edge with a wire loop containing a drop of 2.3 M sucrose to a coated slide. 7. Circle the drop of sucrose with a Dako pen, mark the reverse side of the slide with water-resistant ink. B. For cell cultures 1. Fix cell cultures with fixation solution for 10 min at 4°C, remove from bottle with a rubber policeman, embed pellet in 10% gelatin, PBS, and cool for 10 min at 4°C. Fix at 4°C overnight with 4% paraformaldehyde. 2. Cut pellet into small blocks (less than 1 mm cube). 3. Treat further as for tissue (part A, steps 3-6). a Modified from the method given in ref. 46 with permission.

Figure 11. Picking up resin sections from a water-bath with a wire loop and placing them in a droplet of clean water on a cleaned (coated) slide. Cryosections are picked up with a droplet of sucrose (Protocol 7} and are transferred to a dry slide. 304

7: Reflection-contrast microscopy Semi-thin (0.25 um) sections are well suited for RCM, although ultrathin sections have a superior image definition. Routine plastic embedded tissue can be sectioned using a glass knife or diamond knife on a routine motorized microtome (Reichert-Jung 2050), that makes cutting of sections down to a thickness of about 0.25 um possible. Ultrathin sections made with an ultramicrotome can be brought directly from the diamond or glass knife to a clean microscope slide (see Protocol 10), using a wire loop (Figure 11) (Protocol 8). The sections can then be examined with RCM. Ultrathin cryosections (46) can be brought directly on the microscopic slide and stretch very well without significant artefacts (Protocol 7). Ultrathin sectioning with an ultracryo microtome is now a routine procedure.

Protocol 8.

Sectioning of resin embedded tissue

Equipment and reagents • Ultramicrotome or a motorized microtome (Reichert-Jung 2050) • Glass or diamond knife • Wire loop 3 mm

routine

• Coated or clean glass slides (see Protocol 70) • Hotplate • Millipore filtered distilled water

Method 1. Cut the sections (250 nm) on a motorized routine microtome, or on an ultramicrotome (70 nm sections), using a glass or diamond knife. 2. Collect the sections from the trough, with a wire loop (3 mm) and place on a water drop on the slide (Figure 11). 3. Dry slide on a hotplate for 45 min at 60°C. (For immunohistochemistry collect separately on coated slides and dry on a hotplate for 45 min at 37°C.) 4. Mark the ultrathin sections on the reverse side of the slide with a water-resistant ink. 5. When pre-embedded specimens are sectioned (see Protocol 4), counterstain with 0.01% toluidine blue for 5 min, and wash with distilled water. 6. Store slides in a dust-free slide container.

5.5 Immuno- and histochemical staining for RCM For a theoretical description of these methods the reader is referred to Cornelese and Prins (22), and Prins et al. (23, 24). Immunocytochemistry is done on resin embedded and cryosections (see Section 5.2 and Protocol 9). 305

/. S. Ploem, F. A. Prins, and I. Cornelese-ten Velde Protocol 9. Immunomarking of cryo or resin embedded sections Equipment and reagents All solutions must by filtered with a 0.2 um filter (Schleicher & Schuell, FP030/3) • Hotplate • Incubation buffer: 1% bovine serum albumin (BSA, Sigma, A-9647), 0.1% gelatin (Merck, 4070), in PBS • Aldehyde quench buffer: 0.05 M glycine in PBS . Protein blocking buffer: add 5% serum (in accordance with the second antibody) to incubation buffer • Immunogold reagents: IgG conjugated to 1, 5, or 10 nm gold (Aurion, The Netherlands) . DAB, H2O2 solution: add 10 ul H 2 O 2 to 10 ml DAB

• 1% glutaraldehyde in PBS • Silver enhancement reagents: developer and enhancer solutions (Aurion, The Netherlands) • Diaminobenzidine medium (DAB): freshly prepared 0.05% diaminobenzidine in 0.05 M Tris-HCI pH 6 • 30% hydrogen peroxide . Light Green (BAH/Gurt, 34204) solution 0.2% in distilled water . Haematoxylin (Merck, 15938) 0.2% in distilled water

A. Peroxidase staining 1. Carry out all incubations in a moist chamber. 2. Quench free aldehyde groups with quench buffer for 15 min. 3. Pre-incubate with protein blocking buffer for 5 min. 4. Incubate for 1 h at room temperature with serum according to secondary antibody in a suitable dilution in incubation buffer. 5. Wash four times for 5 min with PBS. 6. Incubate for 1 h with a peroxidase-conjugated secondary antibody in a suitable dilution of incubation buffer. 7. Wash four times for 5 min with PBS. 8. Fix for 5 min with 1% glutaraldehyde in PBS. 9. Wash four times for 5 min with PBS. 10. Develop for 10 min in the dark with DAB, H2O2 (see Protocol 4). 11. Counterstain PO-DAB for 5 min with Light Green. 12. Wash for 1 min with distilled water. 13. Dry for 15 min at 60°C on a hotplate. B. Immunogold staining with silver enhancement 1. Follow part A, steps 1-9 (peroxidase staining procedure). 2. Develop immunogold—after a few wash steps with distilled water—for 5-10 min with silver enhancer (mix developer and enhancer 1:1 just prior to incubation), and wash five times for 3 min with distilled water. 3. Counterstain IGSS with Light Green or haematoxylin for 5 min. 4. Follow part A, steps 12 and 13 (peroxidase staining procedure). 306

7: Reflection-contrast microscopy The immunocytochemical staining efficiency of resin embedded sections (since only their surface is accessible for antibodies) could be somewhat less satisfactory than with cryosections. When antigenicity (45) is diminished by the light fixation with paraformaldehyde, the cryo substitution method described by Edelmann (47) has considerable potential. In this method freezedrying and infiltration in Lowicryl in a cryosorption freeze-dryer are used. If multiple colour immune labelling is done, for example with in situ hybridization for chromosomal studies in the same specimen, the different coloured end-products may dissolve in immersion oil. If this is likely to occur then it is advisable to cover the specimen with a thin layer of protein fixative solution (25) (see Section 5.6).

5.6 Mounting and examining sections on microscope slides In immuno applications the cleaned glass slides must be coated (commercially available coated slides can be used) (Protocol 10). To avoid introducing extra reflecting surfaces, specimens on microscope slides should generally not be covered with coverglasses. If it is necessary to mount the specimen in an embedding medium, this must have a refractive index close to that of glass and immersion oil (n = 1.518). Immersion oil is brought directly onto the specimen. Again it is sometimes useful to smear a small amount of BSA fixative solution (25) over the slides (to create a thin protective protein layer). This prevents the coloured immuno reaction products dissolving in the immersion oil. The RCM oil immersion objective is directly dipped into the immersion oil (Protocol 3). Protocol 10. Glass slide cleaning and coating Equipment and reagents • Glass slides ( Menzel-Glaser, Euroslides, Art. No. 102) • Glass slides with adhesive coating (StarFrost Adhasiv-Objekttrager, Knittel-Glaser, Germany) . Acetone (PA quality) • 96% ethanol

• Aminosilane, 2% aminopropylethoxylane (Sigma) in acetone • Gelatin, chrome alum: dissolve 0.45% (w/v) gelatin in distilled water (500 ml), filter, and add 19.5 ml of a 4% chrome alum solution in distilled water

A. For morphological studies 1. Clean slides by washing them for 1 min with 96% ethanol and 1 min with acetone. 2. Wipe dry with tissue. B. For immunostaining 1. For coating of slides with aminosilane wash twice for 10 min with acetone and coat with aminosilane overnight. 307

/. S. Ploem, F. A. Prins, and I. Cornelese-ten Velde Protocol 10.

Continued

2. Wash twice for 10 min with acetone, wash for 5 min in distilled water, and allow to air dry. 3. Alternatively use glass slides with adhesive coating (commerically available).

6. Summary The few optical parts that are needed for RCM can be easily installed on modern fluorescence research microscopes. Reflection microscopy can, in contrast to absorption microscopy, visualize very thin tissue sections. All of such a thin layer of tissue is in-focus even when an objective of high NA is used. With very thin specimens the microscope image is not disturbed by pre-focal and post-focal optical information, as seen with specimens of routine thickness. Most immunomarkers like peroxidase, alkaline phosphatase, gold, and silver enhanced gold, show a strong reflectance with RCM, resulting in a large image contrast. The detail that can be observed with light microscopy is dependent of both lateral resolution and image contrast. It is therefore not surprising that RCM provides images with a quality close to the optimal possibilities of light microscopy (48). The large image contrast obtained with RCM in immunostaining permits the use in the same specimen of classical cytochemical and histochemical (counter)stains. Most of such stains show a weaker reflectance then the immunomarker stains mentioned above, so that a simultaneous observation of both type of stains is possible. This can be very useful for fine morphological or further cytochemical orientation in cells and tissues. The combination of RCM and EM (including ultracryo sections) in examining the same specimen is an interesting new possibility. The large image contrast of DABox and immunogold (silver) in RCM makes an overview and statistical analysis of stained structures in a large part of the specimen relatively easy and efficient. EM can, however, extend the RCM observations with higher resolutions in smaller selected areas of the specimen. Such a continuum in observations of the same specimen with a very large range of resolutions, was up till now not easily possible. Another possibility of RCM is the study of substances (for example ceramic particles) in tissues, for which no specific stains are available. Such substances can sometimes be visualized only on the basis of a relatively small difference of refractive index in comparison with the tissue in which they are located. Another interesting possibility of RCM is the study of living unstained cells. The cell to substrate contacts can often be clearly seen. Such images provide an information about the attachment of cells to glass surfaces. 308

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Acknowledgements We thank Mr J. Bonnet, from the Laboratory of Histochemistry and Cytochemistry of Leiden University, The Netherlands, for his expert advice on RCM methodology.

References 1. 2. 3. 4.

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8

Histomorphometry A. J. REYNOLDS

1. Introduction The analysis of histological material has traditionally been by comparative morphological analysis. A section of tissue, suitably processed and stained, is examined under a light microscope by a skilled microscopist and a diagnosis made by observing the morphology of the tissue and cells. Whilst this approach is suitable for the majority of analyses conducted, there is a growing need for a more quantitative methodology. Visual assessment of histological preparations is poorly reproducible and measurement has several advantages such as: • objectivity • reproducibility • the ability to detect changes not immediately apparent to the naked eye (1) A problem encountered in interpreting a microscopic image is that one sees a flat image of a section cut from a three-dimensional object and the viewer is required to visualize the spatial context of the object. We are used to seeing objects as a three-dimensional projection of that object and we have a natural tendency to interpret flat sections as projections. An example is that a circular profile will generally be interpreted as being derived from a sphere but, it could also have come from a cylinder. Problems can become compounded if we want to obtain information about the actual size of spatial objects from a study of sections cut from the bulk. For instance, if a structure composed of spherical objects is cut by a section plane it is unlikely that all of the objects will be sectioned through their full diameter, thus most profiles will have diameters smaller than that of the spheres. To assist the microscopist in the interpretation of sections, mathematical relationships have been derived which relate the two-dimensional 'image' with the three-dimensional 'whole'. The French geologist Delesse proved in 1874 that the volume density of the components of rock can be estimated by measuring the relative areas of their profiles (also called the areal density) on random sections cut through the rock. This is one of the fundamentals of measurement and proved for the first

A. ]. Reynolds time that a random section can be a quantitative representation of the material from which it is derived. The techniques used for the quantitative measurement of objects are grouped together under the term 'stereology'. Histomorphometry is the quantitative analysis of histological material to obtain a set of data using stereological techniques. Stereology is defined as: • a body of mathematical methods relating • three-dimensional parameters defining the structure to • two-dimensional measurements obtainable from sections of the structure (2) Stereological methods have been widely used in geology and metallurgy for many years but it is a relatively new technique in biology being some 30 years old (3); and the widespread acceptance of these techniques is largely due to the efforts of Weibel and Loud. The basic parameters of stereology are usually the ratios of two quantities such as the number of objects per volume of tissue and a set of definitions proposed by Underwood have now become generally accepted (4). A list of these definitions, their abbreviated symbol, and dimensions are shown in Table 1 but it is by no means an exhaustive list. To make the above list more self-explanatory the reference system (the denominator of the ratio) is written as a capital subscript, for example the

number o f objects p e r unit area i n a tissue section area fraction of these objects is:

where A is the area of the objects in an area of the tissue AT When examining sections cut from a bulk material the area fraction AA is equal to the volume fraction Vv. But, whilst this is true for a single section if it is large enough, the estimates of either area fraction or volume fraction must be based on a representative sample. In practice this means that several sections should be measured and the average or cumulative totals used for reporting. Stereology thus gives the histologist a means of quantifying images of tissue sections to obtain meaningful data concerning the progression of diseases such as carcinoma. For instance; biopsies of tumours can be taken at regular intervals and the mean nuclear volume measured to monitor the progression of the disease and the effects of treatment on the tumour. Stereology is also useful in other branches of pathology such as microbiology and cytology. 312

8: Histomorphometry Table 1. Symbols used in stereology Symbol

Definition

P PP PL PA Pv L LL

Number of points Number of points on feature per point applied Number of cuts per unit length of test line Number of points per unit test area Number of points per unittest volume Length of element or test line Lineal fraction, i.e. length of line on a feature per unit length of test line Length of lineal elements per unit test area Length of lineal elements per unit test volume Area of feature or test area on micrograph Surface or interface area Area fraction, i.e. area of sectioned feature per unit test area Surface area per unit test volume, e.g. membrane area per unit cytoplasmic volume Volume of 3D features or test volume Volume fraction, i.e. volume of a feature per unit test volume Number of features Number of cuts a feature makes per unit length of test line Number of profiles of a feature per unit test area, e.g. granule profiles per (um2 of section Number of features per unit test volume, e.g. mitochondria per um3 of cytoplasm Mean linear intercept length, i.e. LL/NL Mean profile area, i.e. >AA/NA Mean surface area, i.e. Sv//Vv Mean volume, i.e. Vv/Nv Mean diameter, e.g. of a population of granules Mean diameter of a population of profiles Mean volume of an individual cell or organelle

LA L\/ A S AA Sv V Vv N NL /VA NV L A S V D d v

Dimensions

um-1 um-2 (um-3 um um-1 um-2 um2 um2 um-1 um3 um-1 um-2 (um-3 UM um2 um2 um3 um um um3

The modern trend towards computer automated analysis has made the acquisition of statistically significant data relatively fast but there is still a place for the older manual or computer interactive techniques.

2. Microscopy 2.1 Specimen preparation Histological specimens are derived from living tissue which is a dynamic system and thus prone to rapid changes in morphology, especially when deprived of nutrient and/or oxygen. To minimize these changes the tissue must be preserved in as near lifelike condition as possible to enable measurements to be made on the tissue component in question. While no system has been developed that is absolutely sure not to introduce artefacts, the traditional method for light microscopy has been to chemically fix, dehydrate, and embed 313

A. J. Reynolds the tissue in low melting point paraffin wax. A general protocol of specimen preparation is described in Protocol 1. Protocol 1. Preparation of tissue specimens for light microscopy Equipment and reagents • Automated tissue processor (Hacker) or glass beakers . Water-bath • 10% formalin in physiological saline

• 74OP IMS (BDH) . Xylene (BDH) . Low melting point wax

Method 1. 10% formol saline, at room temperature, for 0.5 h. 2. 70% alcohol, at 40°C, for 1 h. 3. 90% alcohol, at 40°C, for 1 h. 4. 74OP IMS, at 40°C, for 1 h. 5. 740P IMS, at 40°C, for 0.75 h. 6. 740P IMS, at 40°C, for 0.75 h. 7. 740P IMS, at 40°C, for 0.75 h. 8. Xylene, at 40°C, for 1 h. 9. Xylene, at 40°C, for 0.75 h. 10. Xylene, at 40°C, for 0.75 h. 11. Wax, at 60°C, for 5.5 h.

The resultant wax impregnated tissue is then placed in a suitable mould incorporating a unique label for the sample and embedded in fresh wax. A microtome is used to cut sections of desired thickness suitable for light microscopy (2-5 u.m) which are then attached to microscope slides. To generate contrast, the sections are stained with suitable polychromatic dyes and covered with coverslips using an appropriate mounting media. The number of stains available probably number in the hundreds and therefore only three will be described which have relevance for this text. The first, haematoxylin and eosin (H&E), is the commonest strain in present use for routine histological preparation (Protocol 2). Protocol 2.

Haematoxylin and eosin stain (H&E)

Reagents • Xylene (BDH) . 740P IMS (BDH) • Harris' haematoxylin staina

• Eosin stain, 1% aqueous solution . Concentrated hydrochloric acid (BDH)

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8: Histomorphometry Method 1. Take sections to water: (a) Dewax in xylene for 3-5 min. (b) (c) (d) (e)

740P for 10 sec. 90% alcohol for 10 sec. 70% alcohol for 10 sec. Running tap-water for 10 sec.

2. Stain with Harris' haematoxylin for 5 min. 3. Wash well with running water. 4. Differentiate with 1% acid alcohol. 5. Wash and blue in running tap-water for 5 min. 6. Stain in 1% eosin for 5 min. 7. Wash in tap-water for 5 min. 8. Dehydrate in graded alcohol: (a) 70% alcohol for 10 sec. (b) 90% alcohol for 10 sec. (c) 740P, twice, for 10 sec. 9. Clear in at least two changes of xylene. 10. Mount in suitable mounting medium. aThe stain, as bought from the manufacturer, is a poor dyeing agent; to obtain purposeful results it needs to be converted to its oxidation product haematein. This is achieved by the use of an oxidizing agent such as mercuric II oxide (HgO). Before conversion, haematoxylin is added to an aqueous solution of alum (usually the potassium salt—aluminium potassium sulfate). This is the mordanting step, where aluminium ions combine with the haematoxylin dye; the resultant aluminium-haematein complex then stains via the metal ion AI2+.

The second stain is a silver stain to highlight nucleolar organizing regions (NORs) which are thought to represent the level of cell proliferation (5). NORs are segments of chromosomes encoded for ribosomal nucleic acid (rRNA), and are present in specific loops of deoxyribose nucleic acid (DNA) which project into the nucleoli where they can be seen by electron microscopy as ill-defined pale staining regions within the more electron dense regions. Some of the NORs can be identified in histological sections by the use of a silver nitrate method which demonstrates an acidic protein with which some of the sites are associated. It is important to realize that these silver staining NOR associated protein (AgNOR) sites represent only some of the nucleolar organizer regions in each nucleolus. Furthermore, the spot-like silver reactions seen within the nucleoli in paraffin sections may each represent more than one AgNOR since they tend to be closely aggregated in the nucleoli of normal or benign cells (6). 315

A. J. Reynolds This is only one of a number of AgNOR stains; Chapter 4 in this volume lists some alternative stains. Protocol 3.

Silver stain for nucleolar organizer regions (AgNORs) (7)

Reagents • Solution A (50% silver nitrate): 50 g AgNO3 (BDH) in 100 ml distilled water • Solution B (gelatin solution): 2 g gelatin (BDH), in 1 ml formic acid (BDH), 100 ml distilled water

• Working solution: solution A (2 parts): solution B (1 part), mix immediately before use

Method 1. Take sections to water: (a) (b) (c) (d) (e)

Dewax in xylene for 3-5 min. 740Pfor10sec. 70% alcohol for 10 sec. 90% alcohol for 10 sec. Running tap-water for 10 sec.

2. Rinse sections in distilled water for 2 min. 3. Incubate in freshly prepared working solution for 45 min at room temperature. 4. Wash in distilled water for 1 min. 5. Dehydrate in graded alcohol: (a) 70% alcohol for 10 sec. (b) 90% alcohol for 10 sec. (c) 74OP twice for 10 sec. 6. Clear at least two changes of xylene. 7. Mount in suitable mounting medium. 8. AgNOR sites are intranuclear black dots. The background is pale yellow.

Notes: (a) Sections may be counterstained with Neutral Red or carmalum. Heavy counterstaining may obscure AgNORs. (b) Sections may be toned in 1% gold chloride. (c) Working solution deteriorates rapidly on standing. The third is a general stain called immunoperoxidase and is used, with the suitable antibody, for the staining of antigen/antibody sites. This technique is 316

8: Histomorphometry becoming increasingly popular to show the location of a specific protein within a cell or extracellular matrix such as connective tissue. Some proteins are specific to particular cell types and antibodies can be used as cell type markers. For instance there is a form of actin that marks smooth muscle cells, and the CD3 molecule marks T lymphocytes. The sites of protein stain brown and a counterstain is used to highlight nuclei. Chapter 5 in this volume gives a wider description of methods for immunostaining. Protocol 4. General immunoperoxidase method Reagents • Dako pen grease-type pencil • Trypsin (Sigma): 0.05 g trypsin, 50 ml 0.1% calcium chloride (BDH) « Tris-buffered saline: 30.285 g Tris, 16.875 g NaCI (BDH), 187.5 ml 1 M HCI, distilled water to 5 litres, adjust pH to 7.6, 0.5 ml Tweena • TBS/0.1% bovine serum albumin (BSA)/ 0.1% sodium azide: 500 ml Tris buffer, 0.05 g BSA, 0.05 g sodium azide (BDH)b

• Swine serum: 5 ml normal swine serum, 20 ml Tris buffer pH 7.6 • Diaminobenzidine tetrahydrochloride (DAB): 50 ml Tris buffer, two DAB tabletsc • Copper sulfate (BDH): 4 g copper sulfate, 7.2 g sodium chloride, distilled water to 1 litre • Primary and secondary antibody kit (Dako) • Streptavidin (Dako) • Horseradish peroxidase (Dako)

Method 1. Take sections to alcohol (see Protocol 2, step 8). 2. Ring sections with Dako pen. This is to restrict the antibody staining to the section only. 3. Take sections to water (see Protocol 2, step 1). 4. If trypsin is required, place sections in Tris buffer pH 7.6, for 5 min, then trypsin in calcium chloride 37°C, pH 7.8, for the required time according to the antibody used. 5. Rinse in distilled water 6. Block endogenous peroxide in 3% hydrogen peroxide for 5 min. 7. Rinse in buffer. 8. Immerse in swine serum for 10 min. 9. Drain off swine serum. 10. Incubate with primary antibody diluted in TBS+0.1% BSA and 0.1% sodium azide for 30 min at room temperature. 11. Incubate negatives in TBS/BSA/azide for 30 min at room temperature. 12. Rinse briefly in TBS for 1-2 min. 13. Incubate with secondary antibody diluted in TBS for 30 min at room temperature; poly-biotinylated goat anti-rabbit 1:500, monobiotinylated goat anti-mouse 1:300. 14. Rinse briefly in TBS for 1-2 min.

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A. J. Reynolds Protocol 4.

Continued

15. Poly and mono kit. To 5 ml Tris buffer add one drop streptavidin and one drop biotinylated horseradish peroxidase.d Incubate for 30 min at room temperature. 16. Rinse briefly in TBS for 1-2 min. 17. To 5 ml DAB in TBS add 20 ml of 3% hydrogen peroxidee for 10 min at room temperature. 18. Wash in tap-water for 5 min. 19. Copper sulfate solution for 1 min. 20. Wash in tap-water for 5 min. 21. Counterstain with Harris' haematoxylin (see Protocol 2, steps 2-5). 22. Complete mounting procedures as in Protocol 2, steps 8-10. 23. Antigenic sites stain brown. Nuclei stain blue. aDo not add Tween if oestrogen antibody receptors are the target antigen. b Store in fridge. c Store in freezer. dLeave to stand for 30 min before use. " Filter before use.

2.2 Obtaining an image Although it is outside the scope of this chapter to discuss the principles of light microscopy, it is essential to obtain good quality images that are in-focus and with enough contrast to allow the measurement of features. The techniques described in this chapter are all based on the observation of images in normal transmitted light and the microscopist should ensure that the microscope is correctly aligned to obtain the best quality image possible. The first step in obtaining a good image is Kohler illumination. Protocol 5. Procedure for obtaining Kohler illumination Equipment • Light microscope with transmitted light source • Phase telescope

• Slide with good contrast

Method 1. Switch on the light source and centre it if the microscope lamp has adjusters. 2. Place a slide with a tissue section on the microscope stage and obtain an image using a x 10 objective.

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8: Histomorphometry 3. Partially close the field iris until it appears in the field of view. Focus the image of the iris using the sub-stage condenser height adjustment control. 4. Centre the image of the field iris using the sub-stage condenser centring screws. 5. Open the field iris until its boundary is just outside the field of view. 6. Take out an eyepiece and replace it with a phase telescope focused on the back focal plane of the objective lens. 7. Close the condenser iris until it is about 70% of the back focal plane of the objective lens. 8. Replace the eyepiece. 9. When changing objectives the illumination will need to be rechecked.

A more detailed description of setting up the microscope with Kohler illumination can be found in Chapter 1 of this volume.

2.3 Calibration In any quantitative study, the performance of the testing apparatus must be assessed by careful calibration. In the light microscope, the calibration is performed using stage micrometers of known dimensions preferably traceable to national standards. For direct microscopical observation an eyepiece graticule is used. This is placed in the microscope in a position where the scale will appear in sharp focus and superimposed on the image. The graticule normally lies on the eyepiece diaphragm (i.e. in its focal plane, which is coincident with the primary image plane of the objective lens) (8). The eyepiece graticule is calibrated using a stage micrometer in the following manner. Protocol 6. Calibration of the light microscope Equipment • Light microscope • Stage micrometer

• Eyepiece graticule

Method 1. Set up the microscope to achieve Kohler illumination (Protocol 5). 2. Obtain an image of a contrasty specimen using the objective lens required to make measurements. 3. Replace the slide with a stage micrometer and focus onto the scale. 4. Focus the image of the eyepiece graticule by turning the focus ring of

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A. J. Reynolds Protocol 6.

Continued

the eyepiece until the image of the graticule is sharp against the image of the stage graticule. 5. Replace the eyepiece of the microscope and align the zero mark of the eyepiece graticule with the zero mark on the stage micrometer. 6. Count the number of divisions on the eyepiece graticule between the zero and the next convenient mark on the stage micrometer. 7. Calculate the distance on the stage micrometer and divide by the number of divisions on the eyepiece graticule to obtain the dimensions of one eyepiece graticule division.

When making observations from photographic images, a micrograph should be taken of the stage micrometer at the magnification used for recording the specimen image and printed to the same size as the test micrographs. This can then be used to calibrate the measuring device used. For calibrating computer-based image analysis systems the same techniques can be used depending on whether the image capture system (e.g. video camera) is attached to the microscope or free standing.

Figure 1. Micrograph of a section of renal carcinoma stained with haematoxylin and eosin on which is superimposed a simple line graticule. The graticule has been printed in white to make it stand out against the micrograph. Bar = 10 um.

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3. Linearmeasurements 3.1 Intercept measurement As the name suggests, these measurements are based on the measuring of straight lines either directly on an image or on an overlay (graticule) superimposed on the image. These are the simplest types of measurement, and the apparatus needed may be no more sophisticated than a ruler. One example of this type of measurement is the determination of mean nuclear volume, another example is the estimation of the volume fraction (V v ) of the nuclei, in the determination of prognosis in renal cell carcinoma (9). The intercept method uses an overlay graticule ruled with parallel lines. The intercept is the point where the line cuts a feature boundary, i.e. nuclear boundary. In the case of nuclei, there will be an entrance and exit intercept point and the linear distance between them is measured with a ruler, in the case of a micrograph, or with an eyepiece graticule when making the measurements directly off the image. Figure 1 is a micrograph of renal cell carcinoma stained with haematoxylin and eosin (Protocol 2) with a simple graticule of parallel lines overlaying the image. A method for measuring the mean nuclear volume is given in Protocol 7. Protocol 7. Measurement of mean nuclear volume using the point intercept method Equipment • Overlay graticule of known dimensions • Stage micrometer

• Suitable eyepiece graticule • Micrograph of known magnification

Method 1. Superimpose on the micrograph a graticule consisting of a series of equally spaced parallel lines (see Figure 7) or, if measuring directly from the microscope, superimpose the image of the calibrated eyepiece graticule on the image (Protocol 6). The light microscope must be aligned to achieve Kohler illumination (Protocol 5). 2. Measure the total length of line (intercept inside each nucleus. If more than one line intersects a nucleus, measure the length of each intercept and add the two together. 3. Raise each intercept length to the power three. 4. Calculate the mean of the cubed intercept lengths. 5. Multiply the mean value by TT/3 to obtain unbiased estimate of mean nuclear volume. 321

A. J. Reynolds In stereological terms

where V is the mean nuclear volume, L3 is the total length of lines measured raised to the power three, and N is the number of nuclei measured. A worked example of the calculation of mean Value (V) and standard deviation (SD) for the nuclear volume is shown in example 1. These parameters can then be readily used for comparison with those of other samples and for evaluation of statistical significance of differences between these samples. Example 1. Estimation of mean nuclear volume A micrograph of a section taken from a renal cell carcinoma was printed to a size representing X 1000 magnification. A simple line graticule with spacings of 10 mm (Figure 7), was overlaid on to the micrograph during printing so that the graticule is outlined in white. This makes it easier to see against the micrograph. To perform the analysis the total intercept line length in each nucleus was measured and the data obtained is shown in Table 2.

Table 2. Line length measurement from Figure 1, at the magnification of the micrograph 1 mm = 1 um Nucleus

Line

(um3)

1 2 3

4

5 6 7 8 9 10 11 12 13 14 15 16 17

Line length3 (um3)

Line len

(u-m) 5 4 3 7 7 5 4 10 3 6 g 5 g h 3 h 3 h 9 2 9 j 2 k Sum line length (um) 87 a a b c c d e e f

125 64 27 343 343 125 64 1000 27 216 125 27 27 729 8 729 8 Sum line length3 (um3) 3977

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8: Histomorphometry At X 1000 1 mm is equivalent to 1 um. Total line length3 = 3977 um. Number of nuclei measured (N) = 17. Using Equation 3:

The mean nuclear volume is 245.0 ± 307.7 um3. The results reflect the large variation in size of the nuclei typically found in carcinoma tissue. To measure the volume fraction of the nuclei (Vv) a method similar to Protocol 7 can be used except that the total length of the lines needs to be taken into consideration. A simple protocol for the estimation of Vv is as follows, again using the example of Figure 1. Protocol 8. Measurement of volume fraction by the intercept method Equipment • Overlay graticule of known dimensions • Stage micrometer

• Suitable eyepiece graticule • Micrograph of known magnification

Method 1. Superimpose on the micrograph a graticule consisting of a series of equally spaced parallel lines or, if measuring directly from the microscope, superimpose the image of the calibrated eyepiece graticule on the image (Protocol 2). The light microscope must be aligned to achieve Kohler illumination (Protocol 3). 2. Measure the total length of line (intercept) inside each nucleus. If more than one line intersects a nucleus measure the length of each line and add the two together. 3. Measure the total length of the lines on the graticule. 4. Express the total length of intercepts within the nuclei (LNuc) as a fraction of the total line length on the graticule (LT) i.e. LNUC/LT.

Again, in stereological terms:

A worked example using the data in Table 2 and a total line intercept length of 847 um using Equation 4. (The value of 847 um is obtained by dividing the total length of lines on the graticule in millimetres by the micrograph magnification (X 1000).

A. /, Reynolds Example 2. Estimation of volume fraction

Volume fraction is 10.27%.

3,2 Point counting This is similar to the previous examples except that a square graticule is superimposed over the image and the intersections arc used as points fort Vv measurement. A square lattice is convenient to use although the use of triangular and hexagonal lattices have been described (2). The relative merits of each type of graticule have been discussed (10) and the triangular lattice was thought best. F-Iowever, in practice, square lattices are normally chosen for convenience. Attention must be paid to the spacing of the lattice and also the number of points counted by this technique when considering the accuracy of the method. The spacing of the lattice (d) is important for two reasons. The first is that it plays a part in the number of points counted; the more points counted the greater the accuracy. The second is that the error inherent in point counting, when a fixed number of points are counted, depends partly on

Figure 2. Micrograph of a section of renal carcinoma stained with haematoxylin and eosin on which is superimposed a simple grid graticule. The graticule has been printed in white to make it stand out against the micrograph. Bar = 10 um.

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8: Histomorphometry the lattice spacing especially when counting objects with an apparent volume (corpuscular). When measuring Vv of a particular type of organelle, the number of test points needed to achieve the desired accuracy is inversely proportional to the square root of Vv and ideally each organelle should have no more than one point (11). In practice this idealized situation is difficult to achieve and compromise spacings are used, spacings of less than 1 cm are not generally used. Figure 2 is the same micrograph as shown in Figure 1 except that a square graticule has been superimposed onto it. The volume fraction is estimated by counting the number of test points that fall within a feature and dividing by the total number of test points. In stereological terms:

where PN is the total number of test points that fall within the objects to be measured and PT is the total number of test points. Protocol 9. Measurement of volume fraction by the point counting method Equipment • Overlay graticule of known dimensions • Stage micrometer

• Suitable eyepiece graticule • Micrograph of known magnification

Method 1. Superimpose on the micrograph during printing, a graticule consisting of a grid of squares (Figure 2), the graticule will appear white against the micrograph making measurement easier. Print an image of the stage graticule at the same magnification to calibrate the micrograph graticule. If measuring directly from the microscope, superimpose the image of a calibrated eyepiece graticule on the image (see Protocol 6). 2. Count the number of intersections that fall within the nuclei. 3. Count the total number of intersections on the graticule. 4. Express the total count of intersections within the nuclei (PNUC) as a fraction of the total intersections on the graticule (PT) i.e. PNuc/PT.

A worked example of volume fraction estimation is shown in example 3. Example 3. Volume fraction measurement by the point counting method A micrograph of a section taken from a renal cell carcinoma was overlayed with a simple square graticule during printing (Figure 2), the graticule will appear white against the micrograph making measurement easier. Print an image of the stage graticule at the same magnification to calibrate the micrograph graticule and the total points falling in each nucleus was counted. 325

A. J. Reynolds The number of points falling within the nuclei was 13 and the total number of points on the test graticule was 121. Using Equation 5:

The volume fraction is 10.74%. The volume fractions calculated by the intercept and point counting methods are very close to each other showing that either method could be used. In practice, point counting is preferred as it is easier to count the points than to measure lines. 3.2.1 Anisotropy The above paragraphs describe a lattice system that can be used when the tissue is isotropic in the distribution of the items of interest. This is not always suitable for some tissues in which the organelles are not randomly distributed (anisotropic). Estimations of Vv in these cases when using a regular square lattice result in large errors especially when the spacing of the lattice corresponds to the periodicity in the sample. The problem can be overcome by placing the lattice obliquely across the tissue pattern, but a better system to use is one where a series of random points are used. A simple graticule can be devised where a piece of transparent material such as acetate sheet is marked on two adjacent edges with 100 equally spaced points. A table of random pairs of numbers can be used to plot co-ordinates on the sheet which is then placed on the micrograph and the number of co-ordinates which lie on the features of interest counted. The area values can be calculated by the expression:

The random array often yields slightly higher statistical errors than the regular array. But, the random array, in addition to the advantages when studying anisotropic tissues, can allow more points to be applied to single micrographs since it can be applied repeatedly in different alignments without generating bias. The graticule should however be rotated between each application (12). Repeated sampling of a single micrograph with a regular array graticule is difficult to achieve with any degree of ensuring independence between the samples. 3.2.2 Estimation of the number of points to be sampled The statistical error of measurement by point counting of a feature is a function of the inverse relationship of a number of points applied to the feature, the less points counted the greater the statistical error. In any micrograph the features to be measured, e.g. nuclei, will be of different relative sizes thus each 326

8: Histomorphometry feature will accumulate a unique number of points. Therefore the precision of the technique will be dependent on the smallest feature of interest. To determine the number of points that need to be sampled, a pilot study is performed to give a crude estimate of Vv. A way to determine the number of points is to calculate the relative standard error (RSE) using the method of Hally (13) from the expression:

Therefore

and

The value of Vv is the crude value calculated from the pilot study. A worked example of this is as follows. Consider a value of Vv of 0.2 (20%) with a desired RSE of 5%:

However only 20% of the features occupy the total area of the micrograph therefore we must count at least 320/20 X 100 = 1600 points falling on the whole micrograph to maintain the desired 5% RSE. This means that several micrographs or microscope fields of view have to be measured.

4. Automated measurement 4.1 Measurement with an interactive computer system (digitizer tablet) A relatively cheap way of automating the repetitious measurement of features on a micrograph or directly from the microscope has become available due to the falling costs of personal computers (PCs), digitizing tablets, and charge coupled device black and white cameras (CCD). The simplicity of the equipment needed coupled with the operators hand-eye-brain co-ordination make these systems extremely versatile for the measurement of very complex images. A digitizing tablet or bit pad (Figure 3), is an input device in which a stylus or cursor is moved, by hand, over a flat surface. In general, most digitizing tablets employ a grid of wires embedded in the pad which carry a series of high frequency pulses. They operate on the principle of magnetostriction which is the

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A, J. Reynolds

Figure 3. Digitizing tablet and cursor. The micrograph is fixed to the tablet with sticky tape to stop it moving.

change in the dimensions of a ferromagnetic body caused by magnetization or demagnetization, A pen or cursor detects these changes and interprets them as x and y co-ordinates which are then stored in the computer's memory. The procedure for measuring objects is to first calibrate the digitizer tablet with a scale of known dimensions, such as a micrograph of a stage micrometer printed to the same magnification as the test print, and to then draw round the features of interest with the cursor. The device then records the co-ordinates of the trace and the computer calculates parameters such as area, perimeter, largest and smallest diameters. The advantage of these devices is that they measure features with a greater precision than manual point counting techniques and that one tracing can yield several parameters. A disadvantage is that the accuracy of the system is only as good as the tracing skills of the operator, however if the actual area of the objects to be measured is kept above 16 mm2 for a circular profile the errors will be of the order of 6% which is acceptable for most applications in biology. Errors will increase if the shape is more complex than a circular profile but this can be somewhat compensated for by 328

8: Histomorphometry making the image of the feature larger. Of course, the digitizer must be calibrated using a linear scale of known dimensions.

4.2 Semi-automatic image analysers Until the advent of fast personal computers with large memories, automated measurement systems relied on a hard wired computer to perform image capture and analysis with a personal computer used as a 'front end' displaying an interface program to control the system. Figure 4 shows an automated image analyser consisting of a video camera, digitizing pad, personal computer, hard wired computer (under the table), and an image display screen. The video camera can either be fitted with a lens as a free standing image capture device (as illustrated) or be attached to a microscope for direct analysis. The figure shows the Cambridge Instruments Quantimet Q520 image analyser, the lefthand monitor shows the captured image with the features of interest highlighted as a graphics overlay, and the right-hand monitor shows the software interface displaying columns of measurements. The graphics tablet for image editing is in the centre of the figure and the PC mouse is shown to the left. The software is menu-based with, amongst other options, a detection facility which allows the user to determine the range of grey levels measured (see Section 4.2.1). The volume fraction of the nuclei shown in Figure 7 was estimated using this system. The micrograph was placed under the video camera and the areas of maximum density (nuclei) were delineated using the detection function. This is done by observing the image on the display screen and using a software slider controlled by the PC mouse to fill in the areas of interest. The area fraction is determined by a software algorithm similar to the intercept method already described. In a two-dimensional image such as a micrograph the area fraction is equal to the volume fraction, i.e. AA = Vv. The value of the volume fraction obtained by this method was 10.37%. To automate the analysis of similar samples which avoids repetitious actions a sequence of commands can be written called a macro (see Section 4.2.4), but this requires a knowledge of the programming language used with the system. Modern systems now almost always include a personal computer running Microsoft Windows™ or Windows 95(™), with appropriate internal frame grabbers and video display cards. Figure 5 shows 'a state-of-the-art' image analyser consisting of a personal computer, video camera, and output device; the central processing unit (CPU) and image capture boards are housed in a remote box to free-up bench space. The relative low cost of colour cameras have also made it possible to digitize colour images which is of great importance when analysing the products of immunostaining. All of the modern image analysers incorporate a frame grabber attached to the camera so that the images can be digitized and imported into the analyser. The file formats used are normally GIF or bitmap, although most software packages provide 329

A. J. Reynolds

Figure 4. Automatic image analyser with hard wired computer. The micrograph is shown on a light box with a CCD camera above. The digitizing tablet allows for operator editing of the image. The right-hand monitor shows the software interface and the left-hand monitor shows the image with the binary digitized detected regions overlayed on top. The hard wired computer is under the bench to save desk space.

the ability to import files from numerous other sources. TIFF, which was the standard image file formal, seems to be going out of style at present. The size of the images captured or imported depends on the system used but even a relatively small image of 1024 X 1024 pixels uses approximately 1 Mb of space (one image can be stored on one high density floppy disk). With computers with small RAM (random access memory) the programs will run very slowly, a minimum of 8 Mb of RAM is normally required but an increase to 16 or 32 Mb of RAM will significantly increase the speed of operation. The cost of RAM memory has significantly decreased over the last year so that buying a system with a large RAM does not significantly increase the cost of image analyser; and this small extra cost is far outweighed by the time benefit accrued. Similarly, the speed of the CPU is constantly rising, with the new PentiumTM processors running at speeds in excess of 300 MHz, but bear in mind that a computer with a large RAM and slow CPU may in fact run faster than a computer with a fast CPU and small RAM especially when dealing with images. To conserve disk space there are compression programs that will reduce the size of the image by eliminating blank pixels; but compression always results in a loss of data so these programs should be used with care. 330

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Histomorphometry

Figure 5, PC-based automatic image analyser, A modern image analyser equipped with a colour camera and microscope adapter. No graphics tablet is shown here although one can he fitted. The computer is placed under the bench to save desk space. Photo courtesy of Imaging Associates.

4.2.1 Signal processing (erosion and dilatation) The image is recorded by a video camera which converts the image into a parallel array of linear scans with the video signal reflecting the light intensity of the image line by line. By s a m p l i n g the intensity of the video signal at discreet points along each line, the scan is chopped into lines of points. The resulting points arc called picture points or pixels which carry two pieces of information: • spatial position • l i g h t i n t e n s i t y or grey level The continuous band of light i n t e n s i t y from black to while is divided i n t o a n u m b e r of grey levels, normally 0 ( b l a c k ) to 255 ( w h i t e ) . The computer can display on the viewing screen the position of each pixel of a specified range of grey levels corresponding to the component of interest in the image, which can also be superimposed on the actual image of the sample. This discriminated image is called a binary image, also called the segmented image, as each pixel is e i t h e r black (non-diserimmated) or while (discriminated). The size of each pixel can be designated by using a graticule or line of known dimensions

A. J. Reynolds and thus it is a simple matter of totalling the number of pixels in each feature to give area measurements or ratioing the number of pixels of interest against the total number of pixels to give volume fraction. Modern automated systems allow a wide range of parameters to be measured or calculated. The fact that discrimination of image features depends on their grey levels leads to two problems. The first is that the specimen must be evenly illuminated, which is not always easy to achieve with a light microscope or on a micrograph illuminated by an external source; however most image analysers have correction procedures to deal with this. The second problem is that the grey level is not always an unambiguous criterion of a certain feature, i.e. other features not of interest may have the same grey level. This problem can be addressed in two ways. First some image analysis systems are fitted with a digitizing tablet from which the operator can either remove pixels, add in features, or separate touching features. This operator intervention is an important feature as it allows the use of very powerful instruments on images that would otherwise not be suitable. Secondly is by the use of techniques known as erosion and dilatation or a combination of these two. To perform either operation it is necessary to define the structuring element, e.g. a square of 3 X 3 pixels known as a kernel, which is stepped across each pixel in the binary image in turn. The kernel does not have to be a square array of pixels; octagonal, hexagonal, and other polygonal structures may be used. The 3 x 3 pixel kernel is the smallest in general use but the number of pixels in the kernel can be increased to 5 X 5 or larger. A diagrammatic representation of erosion and dilatation is shown in Figure 6. In erosion, if all of the pixels in the kernel fall on an image element then the central pixel of the image is retained. If, on the other hand, one or more pixels in the kernel falls onto background when the central pixel is on the image the central pixel is set to white. This has the effect of removing one layer of pixels from around each object in the binary image, to enlarge holes, and separate touching objects. Dilatation is the reverse of erosion; the kernel is again stepped across each pixel in the binary image. The centre point of the kernel will sometimes fall onto background or sometimes onto an image pixel. When the centre point falls onto an image pixel all of the pixels in the kernel that do not overlap an image pixel are added to the image. This has the effect of adding to the edges of the image and filling in holes. Erosion is a useful tool for removing noise from an image as the image can be eroded until all of the pixels contributing to the noise portion on the binary image are removed. If the image is then dilatated by the same amount the relevant parts of the image can be 'grown' back to their original sizes. A cautionary note—if shape is an important factor and numerous erosion cycles have been performed, the regrown image elements will not retain their original shape. Erosion followed by a dilatation is called opening and is very useful for cleaning-up the edges of features. Conversely dilatation followed by erosion is called closing and is useful for filling-in cracks and holes in the edges of features. 332

Figure 6. A diagram to illustrate the principles of erosion and dilatation, together with their sequential use (opening and closing). Two basic shapes are shown, that on the left representing an object with protrusions whilst that on the right has fissures. The structuring element used to erode and dilate is a kernel of 3 x 3 pixels. Note that when opening is carried out on the image with protrusions these are removed, leaving the basic shape whilst the converse operation closing has to be performed on the fissured image to achieve the same effect. From ref. 8 with permission.

A. J. Reynolds Modern image analysers have functions called ultimate erosion and dilatation. Ultimate erosion works like normal erosion except that the regions in the image remain intact but small regions may be eroded to a single pixel. Ultimate dilatation will grow the regions back to their original size but will place a single layer of pixels around each object so that objects will not touch. 4.2.2 Image enhancement Not all images presented for analysis may be suitable for measurement, this may be due to insufficient contrast in the image or other factors such as touching features or noise. Image analysis can often be simplified if the brightness and contrast can be altered, touching features can be separated, and noise reduced. Image analysers have facilities for enhancing images by independently altering the brightness and contrast values for each pixel in the image. Touching objects can be separated by using erosion and dilatation, or by the use of a digitizing tablet, as discussed in the previous section. By far the most significant addition to the field of image analysis has been the use of filters for removing noise and enhancing the image. All of the filters used are mathematical algorithms that operate on each pixel in the image but take into account the values of neighbouring pixels. A good account of how filters operate can be found in the book by Glasbey and Horgan (14). 4.2.3 Colour imaging The dramatic drop in prices of colour cameras and the increase in memory of personal computers has made image analysis of colour images feasible for most laboratories. Most colour cameras operate on the RGB system which takes separate images of the red, blue, and green portions of the image. This RGB signal is used to display the image in its original colours and also to segment features of interest by their colour. This is a satisfactory situation for most applications but has one drawback. In RGB, it is the ratios of the values of the three channels that determines the colour and also the brightness of the image which makes it difficult to use the colour information for analysis. An alternative method is to use hue, saturation, and intensity (HSI) where the shade of colour is represented by hue, the brightness by intensity, and saturation indicates the mixture of pure colour with white light (15). The advantage of this system is that colour becomes independent of brightness which, when examining slides for morphometry, means that the image analyser can be setup to measure the saturation (which is an indication of amount of staining) and the variations in brightness (intensity) can largely be ignored. 4.2.4 Use of macros With the advent of fast personal computers it is now possible to have very sophisticated image processing engines running in real time. The advantage of this is that the images can be seen on the computer screen and all of the operands on the image can be displayed in a gallery of 'thumbnail' images so 334

8: Histomorphometry that the operator can see w h a t the effect of a certain c o m m a n d has on the image. However the computers used by these systems have to be last w i t h large RAM capacity (see Section 4.2). The image analyser used as an example tor illustration is a Kontron KS300 supplied by Imaging Associates of Thame. Other m a n u f a c t u r e r s of image analysis equipment have their own u n i q u e way of p r e s e n t i n g data and image processing. One of the most u s e f u l functions of modern image analysers is the a b i l i t y to write and play macros. A macro is a string or sequence of commands that operate on an image and can i n c l u d e all of the operations needed to process t h a t image. They can be saved on disk and recalled w h e n needed which cuts down considerably the l i m e t a k e n for repetitious measurements on different images. Unless one is f a m i l i a r with the macro, or programming language of an image analyser the easiest way to write a macro is to process the first image w h e n the macro recorder is activated. Evcry key stroke or function w i l l be l i s t e d by the recorder as a list of commands. If the macro is stored on disc it can be recalled and replayed lor subsequent images. Macros can also be i n t e r a c t i v e if pause commands arc included in the macro list. When the macro reaches a pause command it will slop and wail for the user to i n p u t the appropriate command b e f o r e resuming. The results of r u n n i n g a macro that loads ami processes the image of the k i d n e y carcinoma section t h a t has been used t h r o u g h o u t this text can be seen in Figure 7 and 8. Figure 7 is the original image captured from the microscope

Figure 7. Image of a section of renal cell carcinoma. The nuclei are stained purple. The image shown is the processed image where the brightness and contrast have been stretched using the image analyser. The image was printed using a colour inkjet printer at a resollution of 300 dots per inch (dpi). Bar 10 um.

Figure 8. Final processed image of renal cell carcinoma with the detected nuclei shown in colour. The coloured nuclei have been overlaid as a graphics plain onto the original micrograph (Figure 7). The image was printed using a colour inkjet printer at a resolution of 300 dpi. Bar 10 um.

using a colour camera and frame grabber. The contrast of the image has been stretched to improve the delineation of the nuclei ( p u r p l e ) from the backu r o u n d cytoplasm (light blue). Figure 8 is the processed image where a technique called segmentation was used to detect the areas of m a x i m u m density ( n u c l e i ) . This was done by s e l e c t i n g the pixels in the image w h i c h c o n t a i n e d the density range of i n t e r e s t , in t h i s ease the m a x i m u m and m i n i m u m density v a l u e s of the n u c l e i , from a graphical representation of the t o t a l d e n s i t y range in the image; this created a b i n a r y image in which the areas of i n t e r e s t ( n u c l e i ) were represented as white against a black background. The binary image was then overlayed onto the original image as a graphics plane in w h i c h the segmented p i x e l s have been coloured red to make t h e m stand out. The image analyser was calibrated w i t h an image of a g r a t i c u l e that had been acquired u n d e r i d e n t i c a l conditions to t h a t of the o r i g i n a l image and the area of the pixels selected by the segmentation procedure measured. The results of the a n a l y s i s were a u t o m a t i c a l l y stored in a database f o r m a l . At the end of the macro, the contents of the database were pasted to the clipboard and t h e n to a proprietory spreadsheet package. The spreadsheet was used to c a l c u l a t e the v o l u m e of each nuclei. the average volume, and the standard deviation. "The value obtained was 265.7 = 414.2 um3 which agrees closely w i t h the figure of 245.0 + 307.7 um3 o b t a i n e d by the i n t e r c e p t method described in example 1 of t h i s chapter. Careful examination of the images in Figure 7 and Figure 8 shows t h a t the image analyser has overcslimated the size of some nuclei 336

8: Histomorphometry and underestimated that of others. This is because the image analyser is not capable of such fine discrimination of density changes as is possible with the human eye. Whilst this at first seems to be a disadvantage, the benefits of speed and the fact that they do not tire, compensate for the slight lack of precision. The image analysers remove the possibility of subjectivity that can be found with human operators. For analysing comparable images all one has to do is replay the appropriate macro which does the rest of the analysis. Whilst at first, image processing seems to be complicated, a few hours practice and an understanding of the terminology used in the manipulation of images to achieve the desired result can pay dividends in time and reproducibility of analysis.

5. Histomorphometry The study of the numbers, size, and shape factors of tissue components has become an increasingly popular technique in the diagnosis and prognosis of disease. A brief computerized survey of the literature between 1993 and 1996 revealed several hundred papers where morphometry was one of the main investigative tools. Simple apparatus (e.g. a ruler with a transparent overlay graticule) can be used for the measurement of mean nuclear volume and volume fraction as discussed in Section 3. Automated image analysers whether a digitizing pad attached to a computer or a complete modern image analyser only speed up the analysis for these simple measurements. Table 3 shows the results of analysis by line graticule (Figure 1), square graticule (Figure 2), and semi-automatic image analyser (Figure 4) of volume fraction measurements from the same micrograph of renal cell carcinoma, and the time taken for the measurement. The three values of volume fraction agree well but the image analyser has the advantage in speed. Measuring large numbers of micrographs or fields of view manually is tiring and can lead to errors whereas large amounts of data can be accumulated with the minimum of fatigue by automated analysis. The use of a graphics tablet for the measurement of nuclear shape factors has been described which helps to distinguish between malignant renal carcinoma and benign oncocytoma (16). Fully automated systems really come into their own when more complex measurements such as shape factors, perimeter,

Table 3. Comparison of counting techniques

Line graticule Square graticule Image analyser

Volume fraction (%)

Time taken (min)

10.27 10.74 10.37

10 3 0.5 337

A. J. Reynolds and centre of gravity are required as the computer performs the necessary mathematical algorithms. Nuclear shape factors when combined with the determination of DNA content has improved the ability to predict disease progression in many cancers (17-19). Measurement and numbers of nucleolar organizer regions (NORs) have been adopted as a measure of cell proliferation. These regions in the nucleolus are stained with a silver stain (see Protocol 3) and image analysers are used to count and size, or calculate volume fraction, of the silver stained regions (20-22). Immunoperoxidase reactions (see Protocol 4) are extensively used for the location and counting of specific cell proteins. These specific proteins are unique to certain cells, thus it is a useful way of highlighting these cells for automated image analysis. Rather than count the individual cells that show a positive reaction, it is probably better to determine the volume fraction of the tissue stained. The reason for this is that counting cells by eye is subjective and large variations can be found amongst people counting the same slide. Image analysers have difficulty in separating touching objects (15) unless operations on the binary image (such as opening and closing) are performed. Thus it is faster and more meaningful to estimate the percentage tissue stained. So far this text has been exclusively about the analysis of stained tissue sections but this does not mean that morphometry cannot be applied to other fields of biological science. In fact, automated image analysis is used for many other applications where quantitative analysis of size or shape is required, e.g. the routine screening of cytology samples, blood vessel morphometry (23), sperm head shape in veterinary medicine (24, 25), bacterial shape and colony size in microbiology (26, 27), and trabecula bone patterns for the measurement of bone density (28). This list is by no means exhaustive and the techniques described can be applied to almost any image with sufficient contrast to distinguish the features of interest from the background.

Acknowledgements I am grateful for the help given by Mr Martin Caswell and Mrs Emma Nagle of the Histopathology Department of Wycombe General Hospital for the provision of slides and access to references. I am also indebted to Mr Leslie Stump and Mr Sandy Monteith of Imaging Associates for supplying the Kontron KS300 image analyser which was of great help in writing the section on automatic image analysers.

Instrumentation and sources of supply It is very difficult to give a comprehensive list of suppliers of image analysis equipment in such a rapidly changing field and I apologize to those that I have left out. The addresses of those companies listed below can be found in the appendix. 338

8: Histomorphometry Calibration equipment

Graticules Ltd. Agar Scientific. Image analysers

Imaging Associates Ltd. Foster Findlay Ltd. Leica Cambridge Ltd. Synoptics Ltd. Data Cell Ltd. Media Cybernetics. Optimas Corporation. Data Translation.

References 1. Hamilton, P. W. and Allen, D. C. (1995). J. Pathol., 175, 369. 2. Weibel, E. R. (1979). Stereological methods, Vol. 1, p. 1. Practical methods for biological morphometry. Academic Press. 3. Williams, M. A. (1979). Quantitative methods in biology, pratical methods in electron microscopy (ed. A. M. Glauert). Elsevier, North Holland. 4. Underwood, E. E. (1964). Stereologia, 3, 5. 5. Smith, R. and Crocker, J. (1998). Histopathology, 12, 113. 6. Bancroft, J. D. and Stevens, A. (1990). In Theory and practice of histological technique. Churchill Livingstone. 7. Ploton, D., Menager, M., Jeannesson, P., Himber, G., Pigeon, F., and Adnett, I. J. (1986). Histochem. J., 18, 5. 8. Bradbury, S. J. (1989). In Light microscopy in biology (ed. A. J. Lacey). Oxford University Press. 9. Artacho-Perula, E., Roldan-Villalobos, R., and Martinez-Cuevas, J. F. (1994). J. Clin. Pathol., 47, 324. 10. Frolov, Y. S. and Maling, D. H. (1969). Cartographic J., 6, 21. 11. Hilliard, J. E. and Cahn, J. W. (1961). Trans. Am. Inst. Met. Eng., 221, 344. 12. Williams, M. A. (1969). In Advances in optical and electron microscopy (ed. R. Barev and V. E. Coslett), p. 219. Academic Press, London and New York. 13. Halley, A. D. (1964). Q. J. Microsc. Sci., 110, 295. 14. Glasbey, C. A. and Horgan, G. W. (1995). Image analysis for the biological sciences, statistics in practice (ed. V. Barnett). John Wiley and Sons. 15. Poston, R. (1996). Image processing, December, p. 4. 16. Castren, J. P., Kuopio, T., Nurmi, M. J., and Collan, Y. U. (1995). J. Urol., 154, 1302. 17. Linder, S., Lindholm, I., Falkmer, U. S., Blasjo, M., Sundelin, P., and Vourosen, A. (1995). Int. J. Pancreatol., 18, 241. 18. Yoshii, Y., Saito, A., and Nose, T. (1995). /. Neuro-oncol., 26, 1. 19. Ruizcerda, J. L., Hernandez, M., Gomis, F., Vera, C. D., Kimler, B. F., O'Connor, J. E., et al. (1996). J. Urol., 155, 459. 339

A. J. Reynolds 20. Freeman, J., Kellock, D. B., Yu, C. G. W., Crocker, J., Levison, D. A., and Hall, P. A. (1993). J. Clin. Pathol, 46, 446. 21. Derenzini, M., Trere, D., Oliveri, F., David, E., Colombatto, P., Bonino, F., et al. (1993). J. Clin. Pathol., 46, 727. 22. Kossard, S. and Wilkinson, B. (1995). J. Cutaneous Pathol., 22, 132. 23. Tipoe, G. L. and White, F. H. (1995). Histol. Histopathol., 10, 589. 24. Gravance, C. G., Lewis, K. M., and Casey, P. J. (1995). Theriogenology, 44, 989. 25. Gravance, C. G., Lin, I. K. M., Davis, R. O., Hughes, J. P., and Casey, P. J. (1996). J. Reprod. Fertil., 108, 41. 26. Schafer, C. (1992). Image News, 2,16. 27. Wilkinson, M. H. F. and Meijer, B. C. (1995). Compter methods and programs in biomedicine, 47, 35. 28. Chappard, E., Legrand, E., Basle, M. F., and Audran, M. (1997). Microsc. Anal., 62,23.

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9 Near-field optical microscopy NIEK F. VAN HULST

1. Introduction 1.1 Optical microscopy This is a chapter on an emerging field in optical microscopy: near-field optics, i.e. the optics of subwavelength-sized structures. It may seem surprising to find any aspect unexplored within the mature and well-established domain of optical microscopy. Indeed optical microscopy has reached an outstandingly high level of perfection through centuries and established itself as the main tool in biological studies through its flexibility, applicability, and multitude of optical contrast methods. Yet, a natural limitation had always to be faced: the diffraction limit. It is fundamentally impossible to focus light to an area smaller than the size of the wavelength. Consequently the spatial resolution in lens-based optical microscopy is limited to about half the wavelength. This barrier, already recognized in the previous century by Abbe, has been a stimulus to develop alternative techniques like electron and ion microscopy. Indeed these alternatives have pushed the resolution to the atomic level (in transmission electron microscopy), however generally with the loss of the 'optical' advantages, such as: non-invasiveness, non-destructiveness, operation in native environment (in vivo), high resolution spectroscopic contrast, polarization contrast, and time resolution. The optical approach towards subwavelength resolution naturally involved phase-contrast, however the near-field domain beyond the diffraction limit, dealing with non-propagating waves, has remained until recently beyond the scope of classical opticians. Nowadays in the fields of biology and microelectronics nanometre-sized structures are gaining increasing significance, making the study of the near-field optical properties of 'nano-structures' highly appropriate and stimulating both near-field optical instrumentation and theory. Experimental near-field optics has taken off in 1982, directly following the development of scanning tunnelling microscopy in 1981 (1). Though among the first in the expanding variety of scanning probe methods, near-field optics has been suffering the problem of probe fabrication for almost a decade. Only gradually, with improving efficiency and versatility near-field optics has started to show its latent

Niek F. van Hulst promises of 'optical contrast' at nanometre dimensions (2). The steadily growing number of near-field optical biological applications is indicative of its future importance to light microscopy in biology (3).

1.2 Probe microscopy In the last decade, scanning probe microscopy (1, 4) has emerged as a useful tool in biological research. Especially, atomic force microscopy (AFM) has demonstrated nanometre scale resolution on biological samples at a variety of conditions (5). The introduction of 'tapping mode' AFM (6) and specifically the operation in liquids (7, 8) has enabled the imaging of living biological material, almost unperturbed in near-physiological conditions (9). Indeed activity of cells (9, 10) and enzymes (11) has been followed with high spatial and temporal resolution (12). Lately the intrinsic chemical specificity of AFM, as contained in the very nature of the force, is successfully being exploited for (bio)chemical mapping. Thus 'adhesion mode' AFM has been developed (13), where the difference in chemical bond strength between probe and sample is used as a contrast mechanism. By additional coating of the force sensor with a defined molecular monolayer the interaction can be made specific, leading to 'molecular' force microscopy (14). However, despite the outstanding vertical and lateral sensitivity and emerging chemical specificity of force sensing, optical detection has remained essential to biological investigation, due to its convenience, non-invasiveness, and the extensive variety of contrast mechanisms associated with light. In particular, the chemical specificity contained in spectroscopic information is vital for the understanding of many biological processes. Thus combination of probe microscopy with an optical contrast mechanism has the potential to combine the best of both. This is the domain of near-field optical microscopy (15). The feasibility of near-field optics has been explored experimentally immediately following the start of probe microscopy, even before AFM, by Pohl et al. (16). Yet among the probe techniques nearfield optics has long been considered as an academic peculiarity, especially compared to the enormous impact of AFM. Only in recent years through decisive technological achievements, such as the development of adiabatic fibre pulling and shear force feedback in 1991 by Betzig et al. (17), near-field scanning optical microscopy (NSOM) has gained recognition as a microscopic technique offering optical contrast beyond the diffraction limit with rapidly expanding applications in material science, surface chemistry, and indeed biology (2).

1.3 Breaking the diffraction limit The minimum spot size of a standard or confocal optical microscope is limited by diffraction to approximately 0.5 \/NA, i.e. ~ 300 nm in the optical regime (18). Using a shorter wavelength, such as ultraviolet light, another factor of two might be gained. However, both the lens materials and the biological material 342

9: Near-field optical microscopy are getting opaque towards far UV wavelengths, rendering both lens fabrication and application problematic. The detection of phase yields a vertical resolution down to the nanometre scale, but the lateral resolution is still limited by diffraction. Currently the detection of lateral phase information, in socalled 4rr eon focal microscopy, is an active field of research in an effort to achieve full nanometric resolution in three-dimensions using lens-based optical microscopy (19). In contrast to far-field optics based on propagating waves, near-field optics is restrieted to suhwavelength dimensions, i.e. the optics of non-propagating waves. As a consequence near-field optics does not (cannot) apply lenses, but is based on an antenna (a nanometre-sized dielectric or metallic structure) interacting on subwavelength scale with the local field of a sample to he examined. The local interaction generally modulates the far-field radiation, which is subsequently collected using conventional optics. The interaction volume is determined by the size of the antenna, basically independent of the applied wavelength. The concept of near-field optical microscopy, with a resolution beyond the diffraction limit is schematically illustrated in Figure 1 for the case of an aperture-type antenna. To the left is sketched the diffraction limited finite beam waist (> \/2) of a tightly-focused beam. In the middle the far-field spot size is geometrically confinement to a size a « \ using metallic screening with an aperture. For practical purposes of operation, the actual configuration of an aperture-type near-field optical probe is based on a tapered optical fibre with a metallic coaling for screening off the far-field contribution, as shown on the right side of Figure 1.

Figure 1. Breaking the diffraction limit Schematic illustration of (left) the diffraction limited finite beam waist (> \/2) in a tightly-focused beam, (middle) the geometric confinement to a size a « \ using metallic screening, and (right) the actual configuration of an aperture-type near-field optical probe, based on a tapered fibre with Al coating. 343

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1.4 Scope of this chapter This chapter is focused on near-field optical instrumentation and applications in biology. First an overview of several near-field optical microscopic arrangements will be presented with a discussion on specific advantages and disadvantages as to resolution, efficiency, versatility, and applicability. Next near-field transmission, absorption, polarization, and fluorescence microscopy of various biological surfaces is shown with a lateral optical resolution typically below 100 nm, depending on the arrangement used, but mainly using aperture-type probes: (a) Single fluorophores and proteins. (b) Monolayers and J-aggregates, visualizing the orientation of the molecular dipole and polymer backbone. (c) A virus. (d) Fluorescence of cytoskeletal actin. (e) Chromosomes and fluorescence in situ hybridization (FISH), where the localized fluorescence allows the identification of specific DNA sequences. The presented images are all beyond the diffraction limit and generally accompanied by the simultaneously acquired topographic force image, enabling direct comparison of the optical contrast with the sample topography on nanometre scale. It will be argued that the unique combination of high resolution, specific optical contrast, and ambient operation opens many yet unexplored directions in biological studies.

2. Instrumentation 2.1 Probes and distance regulation The heart of any near-field optical microscope is the near-field optical probe: a nanometre-sized antenna that interacts with the sample and transmits the modulated response to the observer, while scanning over the sample surface at nanometre distance. The probe determines the contrast mechanism, the spatial resolution, and the sensitivity. As such the importance of the probe is comparable to that of the objective in far-field optical microscopy. Many types of probes have been fabricated and applied in various configurations. Figure 2 gives a Figure 2. Types of near-field optical microscopes. Schematic overview of the various experimental configurations for near-field optical microscopy. Top: the antenna (or 'apertureless') type. A nanometric-sized particle, acting as a source or detector (left), is scanned in close proximity over a sample surface. In practise the particle is a passive metallic tip (right), excited by far-field illumination, where the local interaction with the sample surface is detected as a modulation in the scattered far-field. Middle: the aperture-type near-field scanning optical microscope (a-NSOM). A subwavelength-sized aperture is scanned in close proximity to the sample surface. The modulation of the aperture 344

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transmission caused by optical interaction with the sample is detected in the far-field, either by illuminating (transmission mode) or collecting through the aperture (collection mode). Bottom: the photon scanning tunnelling microscope (PSTM). An optical field, evanescent perpendicular to the surface and propagating along the sample surface, is created by dark-field illumination at an angle beyond the critical angle for total internal reflection. The evanescent wave is locally frustrated by a dielectric probe (a tapered fibre or a micro-fabricated lever structure) and converted into a propagating wave. The fraction coupled into the probe is detected in the far-field. 345

Niek F. van Hulst schematic overview of the main experimental configurations encountered in practical near-field optical microscopy, divided into the antenna (or 'apertureless')-type, the aperture-type, and the photon tunnelling configuration.

2.2 Antenna-type or 'apertureless' near-field scanning optical microscopy Ideally the near-field optical probe is of molecular dimensions, serving as a source or detector, which is scanned in close proximity over the sample surface (Figure 2, top). The probe has to be manipulated in a scan pattern over the surface and addressed to collect the near-field interaction information. As a result the probe will be rather a macroscopic tip ending in a nanometresized apex supporting the source or detector. Fabrication of an optical source or detector of nanometre-size dimensions is a challenge in itself. Although efforts based on luminescent particles, polaritons, or submicron-sized photodiodes have been undertaken, none of these has been fully developed. In practice the antenna is simply a passive metallic tip, excited in far-field illumination by a tightly-focused spot, and the local interaction with the sample surface is subsequently detected as a modulation in the scattered far-field. Extreme sensitivity is required to observe the weakly scattered light from the nanometre-sized tip in the presence of light scattered from the sample. Zenhausern et al. (20) have developed a sensitive interferometric detection scheme, while vibrating probe and sample, to discriminate the tip-sample interaction from the background. Images with resolvable features a small as a few nanometres were obtained with contrast based on dipole-dipole coupling. Similarly Koglin et al. (21) obtained 6 nm resolution, in absorption contrast, by using small gold grains on a sharp glass support. The nanometric lateral resolution, far beyond the diffraction limit (even approaching atomic resolution), is illustrative for the potential of probe methods. However these configurations are far from routine, only operational on rather specific samples, and require strong illumination conditions. Biological applications based on metallic tip interactions are yet to be demonstrated.

2.3 Aperture-type near-field scanning optical microscopy The most widely accepted and applied configuration is the aperture-type near-field scanning optical microscope (a-NSOM, Figure 2, middle): a subwavelength-sized aperture is scanned over the sample surface at a distance of a few nanometres. A small fraction of the light passes the aperture and is detected in the far-field, either by illuminating (transmission mode) or collecting through the aperture (collection mode). The close proximity of the aperture to the sample surface, with its spatially varying refractive index, opacity, and anisotropy, results in a modulation of the aperture transmission depending on the aperture position. Additionally the sample topography affects the aperture transmission. Thus images are obtained containing index, absorption, 346

9: Near-field optical microscopy polarization, and topographic contrast. Moreover detection of non-resonant light, such as fluorescence, gives valuable spectroscopic information. The practical feasibility of near-field optical microscopy was first demonstrated by Pohl et al. (16) in 1982 with an optical super-resolution of 20 nm using an aperture at the apex of a sharpened quartz rod. Further development has for long been hampered by fabrication problems of efficient aperture probes. Since 1990 considerable progress has been made with the advent of efficient and reproducible fibre probes, as introduced by Betzig et al. (17, 22). This near-field aperture probe is fabricated by adiabatic tapering of an optical fibre using a special purpose fibre puller (Sutter Instrum. P2000) and subsequent directional coating with ~ 100 nm of aluminium. Thus a 50-100 nm diameter aperture is created, with surrounding aluminium for screening of the far-field. The probe has a brightness of 1-10 nW when several mW laser light is coupled into the fibre, i.e. an efficiency of 10-6 to 10-5, where the input power is limited by the thermal damage threshold of the aluminium coating. The electric field at the probe aperture is highly non-homogeneous due to the metal coating and the polarization of the light: e.g. the electric field is zero at the position where the polarization is parallel to the metal coating. Bouwkamp (23) derived theoretical expressions for the electromagnetic field distribution behind a small aperture (radius a « \) in a perfectly conducting metallic screen. Figure 3 shows the electric field intensity (|E|2) profile at several distances (\/500, \/100, \/20, and X/4) behind a small aperture with a = X/20, with incident light polarized in the horizontal x direction. At short distances (< \/100) the size of the distribution is confined to the dimension of the aperture. At larger distances (> X/4) the field decays rapidly and the confinement diffuses to dimensions rapidly exceeding the size of the aperture. Consequently high resolution imaging can only be expected while operating in close proximity at a distance of only a few nanometres. Using the relatively efficient aperture probes first application of near-field optical microscopy to biological and chemical samples has been explored by Betzig et al. (24) and Moers et al (25). Moreover the pulled fibre probe enabled fluorescence detection of a single molecule, an important achievement in the field of molecular physics, first demonstrated by Betzig and Chichester (26) and immediately followed by single molecular spectroscopy (27) and single molecular fluorescence lifetime detection (28). For biological applications it is important to design the near-field microscope such that standard object glasses or coverslips can be accommodated and the sample can be viewed with conventional high magnification optics for localization of a specific area of interest. Often the near-field optical microscope is based on an inverted microscope with sufficient mechanical stability on the nanometre scale (e.g. the Zeiss Axiovert). The inverted configuration leaves sufficient space for the mounting of a near-field optical probe in the immediate vicinity of the sample surface. The commercial sample table has to be replaced by a combined mechanical and piezo-electric scan table with 347

Figure 3. Electric field intensity profiles IEI2, in x, y, and z direction, in several planes (z = \/500, \/100, \/20, and \/4) behind a small aperture (radius a \/20) in a perfectly conducting metallic screen, as calculated according to the theory of Bouwkamp (23), with incident light polarized in x direction. The grey scale is normalized for optimum contrast in each plot. Close to the aperture the field is concentrated at the rim due to the abrupt dielectric/conductor transition. For increasing distance the field decays rapidly and the confinement diffuses to dimensions beyond the aperture.

sufficient range for coarse and fine positioning, A high NA objective (0.75 NA dry or 1,4 NA immersion) is used for efficient collection, over a large angular range, of the light transmitted by the aperture in interaction with the sample. For fluorescence detection a dichrotc mirror and long-pass filter are used, which block the excitation light, identical to far-field methods. In near-field operation the probe source is confocally imaged onto a point detector, mounted at one of the microscope exit ports. For high light levels, > 1 fW, a photomultiplier tube in combination with a pin-hole (~ 100 um) in the image 348

9: Near-field optical microscopy plane is generally sufficient. For low light levels, < 106 photo-counts/sec, a 100 (Jim area photon counting avalanche photodiode (EG&G, Electro-Optics) is preferentially used because of the high quantum efficiency (~ 59% at \ = 600 nm) and low dark count level (< 10 photo-counts/sec).

2.4 Photon scanning tunnelling microscopy (PSTM) An alternative arrangement is photon scanning tunnelling microscopy (PSTM, Figure 2, bottom), based on frustration of total internal reflection with a sharp uncoated dielectric probe and operated in transmission mode. The optical field is created by dark-field illumination at the inner side of a supporting glass substrate at an angle beyond the critical angle for total internal reflection. The resulting field is propagating along the sample surface and non-propagating (evanescent) perpendicular to the surface. The presence of a sample at the glass-air interface modulates this field and generates new near- and far-field components. The perpendicular evanescent wave is locally frustrated by a dielectric probe and converted into a propagating wave. The fraction coupled into the probe by 'optical tunnelling' is detected in the farfield. The probe can be an uncoated tapered fibre or a micro-fabricated structure (29). Micro-fabricated silicon nitride (SiN) probes are commercially available (Park Scientific Instruments) for conventional AFM applications and are suitable high-index probes with 20-50 nm apex and transparency into the UV. Moreover due to the integrated cantilever the probe can be scanned in close contact with a sample surface with feedback regulation on the force interaction. The combined PSTM/AFM yields simultaneously a topographic and a near-field optical image (30). The operation with uncoated dielectric probes makes PSTM experimentally easier than 'aperture' NSOM, occasionally yielding very high lateral resolution, down to 20 nm (29). However PSTM responds strongly to farfield scattering, which limits the method to samples with topography much smaller than the wavelength. PSTM images of biological objects are generally dominated by far-field scattering, making it hard to find suitable biological applications.

2.5 Distance regulation: shear force microscopy The use of a fibre probe in close proximity to the sample surface, typically a few nanometres, requires a highly sensitive distance sensing and regulating mechanism to avoid the fibre from crashing into the surface. Fortunately with the development of scanning probe microscopy this problem has been solved through the use of piezo-electric manipulators and sensing of probe-sample interaction based on electron tunnelling or atomic forces. In the case of biological applications the sample will generally not be electrically conductive, which eliminates the possibility of electron tunnelling. Force sensing has been 349

Niek F. van Hulst utilized in the many biological AFM applications. Consequently the method of choice is to combine near-field optical and force sensing. Force microscopy based on tapered fibres was demonstrated in 1992 by Toledo-Crow et al. (31) and Betzig et al. (32). They attached the fibre probe to a piezo-electric element, which oscillates the fibre in the lateral direction (parallel to the sample surface) with an amplitude of about 20-30 nm at its resonance frequency (typically > 10 kHz). The oscillation amplitude is measured with a sensitivity of ~ 1 nm using optical detection, either by interferometry (31) or by projecting the far-field diffraction pattern on a split detector, where the difference signal is a measure for the fibre amplitude (32). The oscillation amplitude decreases on approaching the sample surface due to 'shear' forces between probe and sample. The distance between the aperture probe and the sample surface can be adjusted between 1-15 nm, with subnanometre accuracy, by a feedback system based on this shear force detection. Simultaneously a topographic 'shear' force image of the surface is obtained, similar to regular AFM operation. The optical detection of shear forces has the disadvantage that additional stray light is brought into the vicinity of the aperture and that accurate alignment of the detection system with respect to the probe is necessary. An alternative method is to use the piezo-electric material itself to generate a voltage proportional to the amplitude of the oscillation. Based on this idea, the use of quartz crystalline tuning forks was developed by Karrai et al. (33). In this case, the end of the fibre is attached to one arm of a tuning fork and the fork is oscillated at resonance (usually 32 kHz). Again on approaching the sample surface, a decrease of the oscillation amplitude of the tuning fork is observed, which is subsequently used for distance regulation. The tuning fork configuration can be realized very compactly and has a vertical sensitivity of ~ 0.1 nm (34).

2.6 Conclusion Aperture-type NSOM based on metal coated adiabatically tapered fibres, combined with shear force feedback and operated in transmission mode, has proven to be the most powerful NSOM arrangement, because of its true localization of the optical interaction, its various optical contrast possibilities (fluorescence, polarization, etc.), and its sensitivity down to the single molecular level. The system has been commercially available from TopoMetrix Corp. (35) since 1993, and their 'Lumina' microscope is illustrated in Figure 4. In 1996 Nanonics (36) has launched an alternative configuration, utilizing a bent fibre probe. Both systems are based on a commercially available inverted microscope, enabling all contrast modes of conventional optical microscopy, such as phase, DIG, reflection, and fluorescence. A sample area of choice can be examined in further detail with combined near-field optical and force microscopy. 350

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Figure 4. The 'Lumma' NSOM of TopoMetrix Corp. The system is based on a research grade commercial inverted optical microscope, enabling all 'conventional' contrast modes (phase, DIG, reflection, fluorescence) and selection of a sample area for examination on nanometre scale by combined near-field optical and force microscopy. To this end the sample stage has been modified to include a 100 x 100 um2 piezo-electric scanner. The adaptation stage at the right side contains optical and electronic components for probe microscopy and supports the probe head. The probe head (shown tilted at ~ 45") can be rotated away from the sample to facilitate sample exchange and far-field inspection. The probe head allows exchange of aperture-type fibre probes and cantilever-based force sensors for near-field optical, shear force, and normal force microscopy. Reproduced by courtesy of Topometrix (35). a ••••• the base of the inverted microscope; b = bright-field illumination for conventional imaging; c = CCD camera; d = Ar+ laser source to be coupled in near-field fibre probe; e = sample stage with triangular-shaped largearea' piezo-electric scanner around objective; f = probe head, uplifted to facilitate access to sample area; g = position of fibre probe and/or force sensor; h — adaptation stage, supporting probe head; i - microscope support with vibration damping.

Two protocols are given illustrating the more or less standardized steps in fabricating the fibre probe (Protocol 1), and setting up the aperture-type NSOM to examine a specimen (Protocol 2), It should be stressed that safety procedures to protect the eyes from damage should always be followed when using laser light.

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Protocol 1. Fabrication of an aperture-type near-field optical fibre probe Equipment • Optical fibre, preferably standard 125 um • Optical fibre/pipette puller, e.g. type P2000 diameter and single mode for the waveof Sutter Instrum. Co. (37) length to be applied . Thin film aluminium coaler with film thick• Spectroscopic grade acetone and ethanol ness monitor (any type suffices, though the . Fibre cleaver (York Technology Ltd., e.g. sample holder should be modified to carry type FK11) optical fibre which can be rotated) • High NA (> 0.7) optical microscope

Method 1. Cut a piece of fibre of sufficient length for tapering one end and cleaving the other end (> 50 cm). Remove the plastic jacket at one end by mechanical stripping. Remove residual plastic using optical grade tissue rinsed in acetone. Wipe again using ethanol. A clean glass end is obtained by a final rinse in ethanol. 2. The next step, the tapering, is very critical. By local heating and controlled pulling the fibre will be tapered to end in a sharp apex. The sequence of events during execution of a pulling cycle is as follows: (a) The heat turns on. (b) The glass heats up and draws apart until a critical velocity, at which point the heat is switched off. (c) After a delay a hard pull is performed to form the final tapered fibre ends. The taper angle close to the apex should be as large as possible to avoid optical loss in the final end with subwavelength dimensions. The Sutter P2000 fibre puller features CO2 laser heating (~ 10 W on 0 1 mm) and allows reproducible setting of critical parameters such as heating power, heating profile, pull strength, velocity, and delay between heating and pulling. Alternatively hot wire heating can be applied, which requires a tightly coiled wire around the fibre to achieve sufficient heating for fibre melt. 3. Check the taper shape using a high resolution optical microscope. An angle larger than 10° should be obtained while values approaching 30° are preferred. Iterate the pulling cycle to optimize the conditions. 4. The apex should be sharp, typically smaller than 50 nm, preferably with a flat end to facilitate the aperture formation. On optical inspection a sharp tip will only show diffraction fringes. Any irregularity in the diffraction pattern surrounding the final end is an indication of rough structure on a scale > 0.5 um. Regular diffraction fringes indicate a tip size smaller than the diffraction limit, but give little information on the 352

9: Near-field optical microscopy actual size. Only electron or ion microscopy will reveal the tip size and structure. These elaborate inspection techniques are not performed routinely, however, as they require tip coating and enhance the risk of tip damage. 5. In order to define an aperture and confine the light transmission to the dimension of this aperture the sides of the tapered fibre are then coated with metal. Aluminium, with an optical penetration depth of 6 nm, is most suitable for this purpose, where a thickness of 100-200 nm is sufficient for screening the light. The fibre is coated at 10-7 to 10-6 T with high purity (> 99.99%) Al. The free end of the fibre should be coiled to fit in the coater. Evaporation coating at 70°-90° angle to the fibre axis is performed while rotating (~ 60 r.p.m.) the fibre. Thus the apex end is kept free and the aperture formed. The Al coating should be free of grains. In this respect fast coating (10-20 nm/sec) of hot Al (preferably e-beam heated) from a source close to the fibre ( — 1 5 cm) is important. The film thickness is ideally measured using a quartz thickness monitor, while a shutter is important to set the coating time. 6. Finally the fabricated aperture fibre probe is checked by coupling light into the cleaved end of the fibre. Inspect the emitted light using a high resolution optical microscope to check for possible pin-holes in the coating and irregular patterns in the emitted light. A good aperture probe displays a single light spot surrounded by diffraction circles: the Airy pattern as determined by the inspection objective. A O ~ 100 nm aperture should transmit ~ 10 nW with a 10 mW laser coupled into the fibre. Additionally polarization characteristics can be examined. Electron or ion microscopy will reveal the aperture size, but again, however, there is the risk of losing the probe on inspection. Often the probe characteristics are deduced from the performance in near-field optical imaging.

The operation of an aperture-type NSOM with shear force feedback involves: (a) The approach of the fibre probe towards the sample surface until stable shear force feedback is established, very much as in conventional atomic force microscopy based on cantilevered probes. Touching the sample surface in NSOM, however, generally destroys the fibre probe, and replacement with a new good aperture probe requires skilled experience. (b) Optical alignment of the aperture with respect to the collection optics and detector(s) is rather comparable to alignment procedures used in confocal microscopy. (c) Positioning the sample area of choice within the scan range of the piezoelectric scanners. 353

Protocol 2. Fibre approach and optical alignment Equipment • The presence of an aperture-type NSOM with shear force feedback is assumed

Method 1. Mount a fibre probe in the scanning probe head of the microscope. Position a sample or a bare substrate (for alignment) on the sample stage. Locate position and focus of the fibre and the sample area utilizing the far-field imaging high NA objective. It is helpful in practice to illuminate the fibre from the side, to produce a shadow that is easy to locate, and to couple light into the fibre, which makes the aperture light up. 2. Using the coarse approach of the scanning probe head, manipulate the fibre probe to the heart of the field of view, using lateral displacement, and to within 10 um from the sample surface. 3. Close the shear force sensing feedback loop with a high feedback gain, such that the vertical piezo element is at the limit of its range, corresponding to minimized tip-sample distance. Adjust the initial set-point of the feedback loop at a value corresponding to weak shear force interaction, i.e. long working distance. Using manual or motorized fine adjustment, bring the tip close to the sample surface slowly enough for the feedback loop to respond promptly when the interaction comes within the range of the feedback interaction. 4. Optimize the shear force feedback loop by adjusting gain, set-point, mechanical fine adjustment, and optional frequency filters. This procedure depends on the specific electronics, the scanning speed, the scan area, and on the topography of the sample. Generally some initial scanning while optimizing is required. 5. The light emitted by the aperture should be imaged confocally onto the detector, which may be a pin-hole followed by a photomultiplier tube or a small area avalanche photodiode. Optimize the final focusing and the lateral alignment of the aperture relative to detector. Filters or polarization optics in the detection path allow fluorescence contrast, polarization contrast, etc. 6. Through the whole procedure it should be kept in mind that the probe is operated at a distance of some nanometres from the sample surface. Although best avoided altogether, manual adjustment of the system, while in shear force feedback, can be carried out, but with great caution. In mechanical shifting to a new sample area the probe should be retracted (out of feedback range) to a safe distance. The optical alignment is not affected by shifting the sample. 354

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3. Applications 3.1 Single fluorophores and proteins At the fundamental level of high resolution light microscopy is the observation of individual molecules. The exact determination of the molecular location and its dipolar orientation is of interest because many dynamic biological processes, such as protein and molecular conformational changes, can be studied by tracking the position and particular orientation of the molecules. Moreover, the single molecular response provides a sensitive tool to study the local environment of a single molecule at biologically relevant conditions. For instance, the exactness of fluorescence energy transfer experiments between a donor and acceptor pair depends critically on their relative distance and molecular dipole orientation. In addition, if molecules are selectively excited according to their particular orientation, it can be exploited as a tool to trigger or inhibit specific biological reactions. To date, light microscopy of single molecules at ambient conditions with high spatial and temporal resolution is readily achievable by ultrasensitive fluorescence detection in a confocal arrangement (38). In 1993 Betzig and Chichester (26) were the first to image single molecules using the near-field method. Molecules could be localized within a few nanometres and their orientation in three-dimensions could be determined using polarization contrast. This achievement was directly followed by single molecular spectroscopy (27) and single molecular fluorescence lifetime detection (28). Here a typical example of single molecular photodynamics as observed in near-field fluorescence by Garcia-Parajo et al. (39) is presented. A sample consisting of carbocyanine (DiI-C18) fluorophores embedded in a thin polymethyl-methacrylate (PMMA) layer was prepared by spin coating a 5 X 10-8 M concentration of DiI (Molecular Probes, D-282) molecules in methanol, added to a 0.5% weight PMMA in chloroform, onto a freshly cleaned coverslip, resulting in a 5-10 nm layer with a surface coverage of typically a few dye molecules per square micrometre. A time sequence of seven fluorescence aNSOM images displaying single Di molecules dispersed in PMMA is presented in Figure 5. An area of 1.5 X 1.5 um2 is scanned with 10 min interval. A maximum fluorescence signal of ~ 5000 counts/sec is detected. Single molecules with a full width half-maximum (FWHM) of approx. 100 nm are easily discriminated. The observed 100 nm spatial resolution is limited by the aperture of the probe used. In order to observe single molecular rotational movement the near-field scanning optical microscope is extended with two polarization detection channels, allowing fluorescence to be detected simultaneously at 0° and 90° polarization. In time, changes in the fluorescence ratio between both polarization channels are observed from frame to frame for the molecules marked (a) and (b). The ratio signal is a direct measure for the orientation of the molecular dipole moment. Additionally the signal 355

Figure 5. Time sequence of near-field fluorescence images (1 to 7) displaying single carbocyanine molecules (a and b) dispersed in PMMA. An area of 1.5 x 1.5 um2 (200 x 200 pixels, 10 msec/pixel) is scanned with 10 min interval, while detecting single molecular fluorescence simultaneously at 0° and 90° polarization. Slow rotation of the molecules (a) and (b) can be observed in the changing fluorescence ratio between both polarization channels, until the discrete molecular photodissociation in image 5 and 7, respectively. Reproduced from ref. 39 with permission.

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Figure 6. Near-field fluorescence image (600 x 600 nm2) of single allophycocyan (ARC) trimers dispersed on a glass coverslip. The FWHM of the individual molecules is 100 nm, which is limited by the aperture of the probe used. The extrapolated signal for positioning the probe above a peak is 5000-10000 counts/sec, which is sufficient to carry out spectroscopic measurements on a single protein basis. Reproduced from ref. 40 with permission.

fluctuations are indicative of variations in the local molecular photophysics. A unique sign of single molecular detection is the discrete molecular photodissociation of molecules (a) and (b) in image 5 and 7, respectively (39). Dunn et al. (40) have pioneered the near-field approach to image individual photosynthetic proteins. Allophycocyan (APC) was chosen as one of the phycobiliproteins, contained in light harvesting protein complexes, that facilitate the collection and funnelling of light energy towards reaction centres. APC has a trimeric structure, where each of the monomer units contains two open chain totrapyrrole chromophores, covalentty bound to the protein structure, with strong absorption and emission bands in the visible regime. A sample of cross-linked APC trimers was prepared by spin coating a 5 X 10 8 M concentration in 0.1 M of phosphate buffer on a cleaned coverslip. Figure 6 shows a 600 x 600 nm2 fluorescence a-NSOM image of single APC trimers. Distinct features with a width of approx, 100 m are clearly discernible, again limited by the diameter of the tip. The actual individual trimers are 11 nm in diameter. Slight variations are observed in the emission intensities probably caused by the random orientation of the chromophorc dipoles. No bleaching 357

F. van Hulst was observed, reflecting the photoslability of the APC trimer. The extrapolated signal for positioning the probe above a peak is 5000-10000 counts/sec, which is sufficient to carry out spectroscopic measurements on a single protein basis. This achievement demonstrates the feasibility of studying energy transfer dynamics on single molecule pairs (40).

3.2 Monolayers and aggregates A Langmuir-Blodgett film is a highly organized and oriented mono-molecular film. Generally these films serve as model systems for molecular organization in (bio)chemical membranes. For near-field optical microscopy, LangmuirBlodgett films are practical test samples because of their homogeneous surface with nearly no topography and their well-defined molecular orientation. Monolayers of diethylene glycol diamine pentacosa-diynoic amide (DPDA) (25), and 10, 12-pentacosa-diynoic-acid (PCA) (30), prepared by LangmuirBlodgett technique, have been investigated. After UV polymerization the layers were transferred to a glass substrate (41). Depending on the lateral pressure during polymerization and the transfer procedure, several uniform polymerized domains were formed, with a wide range of lateral dimensions, but all with the molecular thickness of 6 nm. The monolaycrs display strong absorption around \ = 500 nm and fluorescence at X = 550-600 nm, where absorption and emission dipole moment are along the highly oriented polycarbon backbone. A combined PSTM/AFM scan of a 1 X 1 um2 area of a PCA film is shown in Figure 7. The force image (Figure 7a) displays the monolayer topography of

Figure 7 PSTM/AFM image (scan area 1 x 1 um2:) of a 10, 12-pentacosa-diynoic-acid (PCA) Langmuir-Blodgett monolayer on a glass substrate. Reproduced from ref. 30 with permission, (a) Force image displaying the topography of monolayer patches, 6 nm thick and dimensions of a few 100 nm. (b) Simultaneously obtained PSTM image displaying absorption of the excitation at \ = 514 nm. Edge steepness of the optical contrast is 30 nm. 358

9: Near-field optical microscopy 6 nm height, as contained in the z-piezo signal in force feedback mode. The corresponding photon tunnelling image (Figure 7b) displays the fraction of the incident p-polarized light of the Ar+ line at 514 nm which is directed to the detector by the local coupling of a dielectric SiN probe (Figure 2, bottom). Monolayer domains with an area smaller than a square wavelength are clearly visible in the photon tunnelling image. Clearly the lateral resolution is beyond the diffraction limit, with an edge steepness of 30 nm. The PSTM signal on the domains is 10% below the signal detected on the surrounding glass, which is in agreement with the measured absorption of a PCA monolayer for green light by far-field methods. Consequently, the PSTM contrast is mainly caused by absorption for this sample (30). The influence of the topography on the optical coupling is limited for this specific thin sample, however for thicker biological samples the effect of topography and far-field scattering becomes dominant (29). An a-NSOM image of a 4 X 4 (um2 area of a DPDA film is presented in Figure 8. In the shear force image (Figure 8a) several domains and the underlying glass substrate are visible. Figures 8b and 8c show the corresponding near-field fluorescence images, where the incident linear polarization is directed perpendicular between the two images. Peak value of the fluorescence intensity is about 105 photons/sec. The absorption and emission dipole moments of the polydiacetylenes are oriented parallel to the polymer backbone, where the orientation is uniform over each domain due to the crystallinity of the Langmuir-Blodgett film. The near-field fluorescence images clearly demonstrate the high anisotropy of the polymerized diacetylenic films with ~ 100 nm lateral resolution: domains which are fluorescent for a given polarization direction in one image are dark for the perpendicular polarization direction in the other image. The force image displays the topography of the monolayer domains with a lateral resolution of ~ 30 nm, showing some surface roughness and a few non-fluorescent structures. Comparison of these images clearly demonstrates the advantage of near-field optics in combination with force microscopy: the near-field optical images allow determination of the polycarbon backbone orientation and polymerization efficiency, additional to the topography in the simultaneously recorded force image, both with high lateral resolution (25). Thin films of J-aggregates are also interesting model systems for study of molecular orientation and ordering at nanometre scale. The strong coupling of the dipoles in J-aggregates over many nanometres causes large spectral shifts and narrowing of bands with peculiar exciton dynamics, which makes them specifically interesting subjects for near-field optical microscopy. The group of Barbara (42) has performed extensive studies of J-aggregates by near-field fluorescence microscopy. Interest was focused on J-aggregates of pseudoisocyanine (PIC) grown in thin poly(vinyl sulfate) (PVS) films on quartz substrates, which display a fibrous structure along several hundreds of microns with about 150 nm width and 10 nm thickness. Absorption and 359

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Figure 8. A 4 x 4 um2 scan of a Langmuir-Blodgett monolayer: UV polymerized diethylene glycol diamine pentacosa-diynoic arnide, DPDA. Reproduced from ref. 25 with permission, (a) Shear force image, showing the topography of 8 nm monolayer patches on the glass substrate. (b) and (c) Near-field fluorescence images with mutually perpendicular directions of linearly polarized excitation, showing the high anisotropy of the Langmuir-Blodgett film and indicating the orientation of the polymer backbone.

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b)

Position um) Figure 9. Near-field fluorescence images (5 x 5 um2) of PIC J-aggregates in thin PVS films excited with 570 nm light from a a-NSOM probe. Reproduced from ref. 42 with permission, (a) and (b) Excitation light from the probe polarized vertical and horizontal (on the page) respectively, with all polarization directions of fluorescence detected. The strong polarization dependence of direct excitation to the lowest energy excitonic state of the aggregate is clearly observed, (c) Line scans across several of the aggregate fibres of (a) and (b).

emission of these PIC aggregates peaks sharply at 573 nm, the so-called J-band corresponding to transitions to the lower edge of the excitonic hand. Fluorescence a-NSOM images for linearly polarized excitation at \ = 570 nm and detection at \ > 590 nm are presented in Figure 9. Both polarization perpendicular and parallel to the scan direction are shown in Figure 9a and 9b. respectively, while all polarization directions of fluorescence arc detected. A strong polarization dependence of direct excitation to the lowest energy excitonic state of the aggregate is clearly observed in the preferentially vertically and horizontally aligned fluorescent fibres in Figures 9a and 9b, respectively. 361

Niek F. van Hulst The line scans across several of the aggregate fibres in Figure 9c shows extinction ratios up to 9:1. The local transition dipole is along the long axes of the aggregates, both for absorption and emission. In contrast to this, fluorescence excitation images recorded at lower wavelength (488 and 514 nm), i.e. higher energy excitonic transitions, show little dependence on the excitation polarization with rather a slight preference for polarization perpendicular to the long axis of the aggregates. From these combined spectroscopic and polarization data a herringbone ordering of monomers within the aggregate could be concluded. Moreover, the high spatial resolution sets an upper limit of ~ 50 nm to the scale of excitonic energy migration (42). Clearly the unique combination of spectroscopic, polarization, and topographic data, all on nanometre scale, reveals details inaccessible to far-field light microscopy.

3.3 Virus Viruses are generally not accessible by conventional optical microscopy and in consequence an interesting object of study for the near-field alternative, to demonstrate the high resolution potential in biological applications. A wellknown candidate is the tobacco mosaic virus (TMV), a rod-shaped virus about 300 nm long and 18 nm in diameter, with a central hole of 4 nm diameter enclosing a single-stranded RNA molecule. The TMV protein subunits pack to form a helix with a well-known structure. Indeed TMV was one of the first objects used to test the a-NSOM of Topometrix Corp. (43). TMV particles were attached to amino-silanated atomically flat mica by glutaraldehyde, and imaged after rinsing and drying. Individual TMV particles could be resolved by this technique with separations of less than 30 nm. Interestingly the TMV particles displayed higher optical transmission than the supporting mica substrate, probably caused by the difference in refractive index, but more likely induced by the distance regulation on the TMV topography. Detailed nearfield optical imaging of isolated viruses is an area yet to be explored.

3.4 Cellular surface and cytoskeleton Cellular structures have been the object of study for conventional optical microscopy ever since the earliest days of microscopy. For scanning probe microscopy cells are rather large objects, where generally only the surface can be studied. AFM studies have yielded however a wealth of information on cell membrane structure (pores (10), antigens (44), and proteins), mechanical cellular properties (membrane hardness, visco-elasticity) (9), and the temporal behaviour of living cells (interaction, growth, formation of pseudopodia) (9). Occasionally, subsurface structure is obtained by applying high forces (> 10 nN) in contact force microscopy (10) such that the cytoskeleton shows up while the membrane is locally pressed down. The spectrum of structural information can be extended using specific labelling techniques (immunogold (44), enzymes (45), or fluorescence), as they have been developed for electron 362

9: Near-field optical microscopy microscopy and (confocal) fluorescence microscopy. Clearly fluorescent labels offer interesting potential for near-field fluorescence detection with its single molecular sensitivity. As such fluorescence NSOM forms a bridge technique between AFM and confocal fluorescence microscopy, because in addition to the surface topography specific cell surface and skeletal constituents can be localized with subdiffraction resolution through their fluorescence. In 1993 Betzig et al. (24) showed the first application of near-field fluorescence microscopy to labelled cytoskeletal actin in fixed mouse fibroblast cells. Hereto Swiss mouse 3T3 fibroblast cells were fixed in formaldehyde and air dried on a glass coverslip, after specific fluorescent staining of filamentous actin with rhodamine-phalloidin. Figure 10 shows images of a cellular protrusion: a flat thin lamellipodium of the 3T3 cell. Conventional and scanning confocal fluorescence images, both with X 100, NA 1.3 objective, are presented in Figures 10A and 10D, clearly showing the effect of reduced detection volume in confocal detection. Figures 10B and 10E are 10 X 10 um2 magnifications of the lamellopodium area to facilitate comparison with the shear force (Figure 10C) and near-field fluorescence images (Figure 10F) of the same region. Clearly, the force image is distinctly different from the near-field fluorescence image although they were acquired simultaneously. The force image shows mainly surface topography with about 50 nm lateral resolution, whereas the near-field fluorescence image shows the cytoskeleton organization in fine detail with a lateral resolution of about 100 nm, well beyond the diffraction limited resolution of the confocal fluorescence image. The nearfield fluorescence contrast is similar to that seen in confocal fluorescence images, easy to interpret, though with improved resolution, and almost bleach-free. Yet, it should be noted that the near-field fluorescence mainly originates from the cellular volume within about the first 100 nm below the cell surface, while the confocal section as generally obtained is fluorescence of the plane of focus.

3.5 Chromosomes and fluorescence in situ hybridization Drosophila polytene interphase chromosomes are a classical object of study with phase-contrast optical microscopy, because of their relatively large size compared to human chromosomes and their characteristic natural band pattern of alternating dark bands and lighter interbands. Both bands and interbands contain fibre-like structures, where bands are composed of densely coiled fibres, while interbands display more dispersed fibrils with a higher content of DNA and protein. The fibre width is typically 10-30 nm, which is associated to the first winding of the DNA double helix around histones forming a 11 nm thick nucleosome fibre and the subsequent coiling of the nucleosomes into a 30 nm thick solenoid fibre. Polytene chromosome substructure has been extensively studied by electron microscopy and more recently also by AFM (46, 47). The band structure is revealed as strong topography in 363

9: Near-field optical microscopy Figure 10. Six images of a stained single lamellipodium from a Swiss mouse 3T3 fibroblast cell. Reproduced from ref. 24 with permission. (A) Conventional fluorescence image with a x 100, 1.3 NA objective. (B) Magnified image of the boxed area in (A). (C) Shear force topographic image of the same area. (D) Confocal fluorescence image with the same x 100, 1.3 NA objective. (E) Magnified confocal image from (D). (F) Near-field fluorescence image. The NSOM image combines resolution superior to confocal microscopy with contrast that provides more detail than in force microscopy.

AFM data, as is clearly illustrated in Figure 11. Figure 11a displays a large area AFM scan of a Drosophila polytene chromosome after fixing, squashing, and air drying on an object glass, as it was imaged at TopoMetrix Corp. (35). The correspondence with the well-known optical picture is striking, where in the AFM image the bands are high and interbands lower. Figure 11b shows a more detailed scan of an interband region, revealing 10-30 nm fibres with a variable extent of coiling, especially when extending into the band area. Finally Figure 11 shows the results obtained with the TopoMetrix Aurora a-NSOM obtained on the same chromosome sample, with the corresponding shear force and near-field 'bright-light transmission' image in Figures 11c and lid, respectively (35). The high correlation between the force and near-field image are an indication for topographic-induced contrast in the near-field optical images. Yet, also differences are observed, e.g. the stretched fibres appear bright in transmission, while the condensed coils appear dark. Again a better defined high resolution contrast is expected by monitoring near-field fluorescence.

Obviously, human chromosome structure has been investigated with all the microscopic techniques available. In addition to the pure morphological examination of chromosomes, specific information on the DNA sequence can be correlated to the chromosome structure by the use of labelling techniques, specifically in situ hybridization as developed almost 30 years ago (48). Especially optical detection of in situ hybridized DNA, on the basis of fluorescence labels or enzyme generated dyes, has promoted fluorescence in situ hybridization (FISH) to one of the major cytogenetic detection methodologies for human genetics (49). FISH enables direct visualization of topological or positional information of gene sequences in a fluorescence microscope, allowing rapid localization of genomic DNA fragments in morphologically preserved inter- and metaphase chromosomes. A resolution better than 106 base pairs can be obtained using (pro)metaphase chromosomes (50), while 1 kb is feasible on stretched DNA. The resolution of the fluorescence labels is fundamentally limited to ~ 300 nm by diffraction in conventional fluorescence microscopy, yet localization of the numerous closely linked genes requires mapping at a higher resolution. Using electron microscopy immunogold DNA probes can be imaged with nanometre resolution. Putman et al. (45) pioneered the potential of atomic force microscopy (AFM) in the detection of morphological in situ hybridization labels, and were able to discriminate morphological labels 365

Niek F, van Hulsl

Figure 11. Four images of a Drosophila polytene chromosome after fixing, squashing, and air drying. Reproduced by courtesy of Topometrix (35). (a) Large area (150 x 150 um2) AFM scan revealing the band structure as high bands and low interbands, with remarkable correspondence to the well-known optical picture, (b) A more detailed AFM scan (12 x 12 um3) of an interbank region, revealing 10-30 nm fibres with a variable extend of coiling, (c) Shear force topographic surface plot (7.5 x 7.5 um2). (d) Corresponding nearfield 'bright-light transmission' image.

of 75-100 nm diameter, after enhancement by an enzymatic reaction, however without observing single copy DNA targets. Despite the higher resolution of electron and force microscopy they lack the multiplicity of fluorescence detection. In fact, the widespread use of FISH is mainly due to its unparalleled specificity afforded by the potential of multicolour labelling in one preparation (50). Near-field fluorescence microscopy has the potential to combine the best of both: optical resolution beyond the diffraction limit and multiplicity through multicolour fluorescence detection with sensitivity down to the single molecular level. Moers et al. (51) have explored near-field two-colour fluorescence detection of FISH labels on human metaphase chromosomes, combined with the chromosomal morphology as obtained by shear force detection. Some results are presented below. 366

9: Near-field optical microscopy Figure 12 shows a scan of a human metaphase chromosome 1, with specific labelling of the telomeric region (top of the image). The metaphase chromosome spread was prepared on a microscope coverslipv according to routine procedures. The repetitive DNA probe pl-79 was used, which is specific for the telomere region (1p36) of the short arm of chromosome 1, with an insert size of 0.9 kb. The probe pl-79 was labelled with digoxigenin and detected by cyanine (Cy3, an orange fluorescent dye) (51). The pixel size is 35 nm. The scan speed is 40 msec/pixel. In the shear force image (Figure 12a) the high spatial frequency filtered piezo feedback signal is displayed, revealing the well-known metaphase chromosomal structure with well-separated chromatids, details as small as 40 nm and height up to ~ 150 nm. The corresponding nearfield fluorescence image (Figure 12b) displays the green fluorescence at X > 570 nm, using BG39 and KV550 Schott filters, with excitation by the 521 nm Kr+ line. The pl-79 probes are visualized as distinct substructure in the telomere region with at least five probes in each chromatid (maximum 700 counts/pixel). Some bright fluorescent material, not associated with the chromosome, is detected on the glass substrate in the right side of Figure 12b. In addition to the bright signals at the telomere several isolated weak spots (—50 counts/pixel) are detected on the chromatids in the centromeric region, as presented in a magnified view in Figure 12c. These labels are most probably due to non-specific hybridization of pl-79. Figure 12d shows a 1 um line trace through two weak Cy3 labels as indicated in Figure 12c. Both fluorescence spots have a width of 80 nm, while separated at a distance of only 110 nm. The line trace clearly indicates the superior optical resolution provided by NSOM, including the ability to localize the fluorescence maximum with an extreme accuracy of a few nanometres. The signal level of the fluorophores in Figure 12d corresponds to ~ 1000 counts/sec, which is estimated to originate from less than ten Cy3 molecules. Evidently the single molecular sensitivity is retained also when imaging 'large scale' biological structures, such as chromosomes.

4. Conclusions Near-field scanning optical microscopy is a true optical microscopic technique allowing fluorescence, absorption, and polarization contrast with the additional advantage of nanometre lateral resolution, unlimited by diffraction. Especially fluorescence NSOM gives a clear high resolution contrast and induces virtually no bleaching as opposed to confocal fluorescence microscopy. Bright-field NSOM in transmission generally yields a complicated contrast caused by a mixture of dielectric and topographic contributions. Shear force feedback is essential for reliable operation of aperture NSOM based on fibres, especially while scanning over soft surfaces of cells and chromosomes. The force feedback gives a topographic map of the sample surface simultaneously with the near-field optical image. Also photon tunnelling microscopy in 367

Figure 12. Human metaphase chromosome 1, with specific labelling of the telomeric region (top). Scan area 7 x 7 um2, with 35 nm pixel size and 40 msec/pixel. Reproduced from ref. 51 with permission, (at Shear force image, high pass filtered in horizontal direction, showing chromosomal structure with details as small as 40 nm and height up to - 150 nm. (b) Corresponding near-field fluorescence image, displaying green Cy3 fluorescence with at least five probes in the telomere region of each chromatid. (c) Magnified view of the centromeric region of the near-field fluorescence image, showing several isolated weak spots. (d) A 1 (um line trace through two weak Cy3 labels as indicated in (c). Both fluorescence spots have a width (FWHM) of 80 nm, while separated at a distance of only 110 nm, which clearly illustrates the superior lateral optical resolution provided by NSOM. 368

9: Near-field optical microscopy combination with force microscopy allows routine scanning with a high optical lateral resolution, however interference effects can be dominant on surfaces which display extensive scattering. As such, in contrast to a-NSOM, the application potential of PSTM to biological surfaces is rather limited. Several biological applications of near-field optical microscopy, in combination with force microscopy have been presented. It is shown that aperture NSOM with fluorescence detection gives (bio)chemical specificity and orientational information, in addition to the simultaneously acquired topographical image. On the microbiological scale the technique has large potential for DNA sequencing and molecular organization on membranes. At the cellular level it allows study of the role of the cytoskeleton in cellular mobility in cell growth, cell migration, formation of protrusions, etc.

5. Future outlook Near-field optics is gradually becoming mature and moving from the developmental stage towards applications. The first commercial instruments have appeared on the market, yet the operation requires a rather specialized and experienced microscopist. The aperture probe brightness of ~ 10 nW is sufficient for single molecular detection, yet is still rather limited for spectroscopic applications, such as fluorescence and Raman, where photon noise will be a fundamental limit in the speed of imaging. The lateral resolution is 20-100 nm, almost an order of magnitude beyond the confocal microscopic resolution, yet still rather poor compared to resolution achieved with force microscopy. Ongoing research on probe fabrication through optimized fibre pulling and etching or micro-machining may result in future probes with higher efficiency, versatility, and resolution. All biological applications presented have been obtained on dry samples. Imaging at physiological conditions obviously requires operation in liquid. In principle near-field optical probes can be made to function in liquid, just like force sensors, however reliable imaging on soft material is rather delicate and yet to be developed for the optical alternative. Finally the monitoring of biological processes requires the development of high frequency scanning and high efficiency probes for sufficiently fast image acquisition. For further information on recent advances in the development and application of near-field optical microscopy the reader is referred to the Proceedings of the international conferences on near-field optics (52, 53).

Acknowledgements Images were generously made available by: Paul West, Gary Williams, and Wouter Rensen of TopoMetrix Corp., Santa Clara, California; Paul Barbara, 369

NiekF. van Hulst Daniel Higgins, and Chuck Tomlinson of University of Minnesota, Minneapolis; Sunney Xie of Pacific Northwest Labs, Richland, Washington; Eric Betzig of NSOM enterprises. The author thanks Ton Ruiter, Marco Moers, Joost-Anne Veerman, Maria Garcia-Parajo, Wouter Rensen, Kees van der Werf, Frans Segerink, Eric Schipper, Ine Segers, and Bart de Grooth, all of the Applied Optics group at University of Twente, The Netherlands, for numerous data, their assistance, and suggestions. This research is supported by the European Human Capital & Mobility network on Near-field Optics and Nanotechnology and the Dutch Foundation for Fundamental Research (FOM).

References 1. Binnig, G., Rohrer, H., Gerber, Ch., and Weibel, E. (1982). Phys. Rev. Lett., 49, 57. 2. Paesler, M. A. and Moyer, P. J. (1996). In Near-field optics, theory, Instrumentation and applications. Wiley, New York. 3. van Hulst, N. F. and Moers, M. H. P. (1996). IEEE Eng. Med. Biol., 51. 4. Binnig, G., Quate, C. F., and Gerber, C. (1986). Phys. Rev. Lett., 56, 930. 5. Hoh, J. H. and Hansma, P. K. (1992). Trends Cell Biol., 2, 208. 6. Zhong, Q., Inniss, D., Kjoller, K., and Elings, V. (1993). Surf. Sci. Lett., 290, L688. 7. Putman, C. A. J., van der Werf, K. O., de Grooth, B. G., van Hulst, N. F., and Greve, J. (1994). Appl. Phys. Lett., 64, 2454. 8. Hansma, P. K., Cleveland, J. P., Radmacher, M., Walters, D. A., Hilner, P. E., et al. (1994). Appl. Phys. Lett, 64, 1738. 9. Putman, C. A. J., van der Werf, K. O., de Grooth, B. G., van Hulst, N. F., and Greve, J. (1994). Biophys. J., 67, 1749. 10. Henderson, E., Haydon, P. G., and Sakaguchi, D. S. (1992). Science, 257, 1944. 11. Radmacher, M., Fritz, M., Hansma, H. G., and Hansma, P. K. (1994). Science, 265, 1577. 12. Guthold, M., Bezanilla, M., Erie, D. A., Jenkins, B., Hansma, H. G., and Bustamante, C. (1994). Proc. Natl. Acad. Sci. USA, 91, 12927. 13. van der Werf, K. O., Putman, C. A. J., de Grooth, B. G., and Greve, J. (1994). Appl. Phys. Lett., 65, 1195. 14. Florin, E.-L., Moy, V. T., and Gaub, H. E. (1994). Science, 264, 415. 15. Pohl, D. W. and Novotny, L. (1994). /. Vac. Sci. Technol B, 12, 1441. 16. Pohl, D. W., Denk, W., and Lanz, M. (1984). Appl. Phys. Lett., 44, 651. 17. Betzig, E., Trautman, J. K., Harris, T. D., Weiner, J. S., and Kostelak, R. L. (1991). Science, 251, 1468. 18. Abbe, E. J. (1882). R. Microsc. Soc., 2, 300 and 460. 19. Hell, S. W. and Stelzer, E. H. K. (1992). J. Opt. Soc. Am., A9, 2157. 20. Zenhausern, F., Martin, Y., and Wickramasinghe, H. K. (1995). Science, 269, 1083. 21. Koglin, J., Fischer, U. C., and Fuchs, H. (1996). J. Biomed. Opt., 1, 75. 22. Betzig, E. and Trautman, J. K. (1992). Science, 257, 189. 23. Bouwkamp, C. J. (1950). Philips Res. Rep., 5, 321. 370

9: Near-field optical microscopy 24. Betzig, E., Chichester, R. J., Lanni, F., and Taylor, D. L. (1993). Bioimaging, 1, 129. 25. Moers, M. H. P., Gaub, H. E., and van Hulst, N. F. (1994). Langmuir, 10, 2774. 26. Betzig, E. and Chichester, R. J. (1993). Science, 262, 1422. 27. Trautman, J. K., Macklin, J. J., Brus, L. E., and Betzig, E. (1994). Nature, 369, 40. 28. Xie, X. S. and Dunn, R. C. (1994). Science, 265, 361. 29. van Hulst, N. F, Moers, M. H. P., and Bolger, B. (1993). J. Microsc., 171, 95. 30. Moers, M. H. P., Tack, R. G., van Hulst, N. F., and Bolger, B. (1994). J. Appl. Phys., 75, 1254. 31. Toledo-Crow, R., Yang, P. C., Chen, Y., and Vaez-Iravani, M. (1992). Appl. Phys. Lett., 60, 2957. 32. Betzig, E., Finn, P.L., and Weiner, J. S. (1992). Appl. Phys. Lett., 60, 2484. 33. Karrai, K. and Grober, R. D. (1995). Appl. Phys. Lett., 66, 1842. 34. Ruiter, A. G. T., Veerman, J.-A., van der Werf, K. O., and van Hulst, N. F. (1997). Appl. Phys. Lett., 71, 28. 35. Topometrix Corp., 5403 Betsy Ross Drive, Santa Clara, California 95054-1162, USA. 36. Nanonics, Manhat Technology Park, Malcha 91487 Jerusalem, Israel. 37. Sutler Instrument Company, 40 Leveroni Court, Novato, California 94949, USA. 38. Xie, X. S. (1996). Ace. Chem. Res., 29, 598, and references therein. 39. Garcia-Parajo, M. F., Veerman, J.-A., Ruiter, A. G. T., and van Hulst, N. F. (1998). Ultramicroscopy, 71, 311. 40. Dunn, R. C., Allen, E. V., Joyce, S. A., Anderson, G. A., and Xie, X. S. (1995). Ultramicroscopy, 57, 113. 41. Tillmann, R. W., Radmacher, M., Gaub, H. E., Kenney, P., and Ribi, H. O. (1993). J. Phys. Chem., 97, 2928. 42. Higgins, D., Reid, P. J., and Barbara, P. F. (1996). J. Phys. Chem., 100, 1174. 43. Pylkki, R. J., Moyer, P. J., and West, P. E. (1994). Jpn. J. Appl. Phys., 33, 3785. 44. Neagu, C. R., van der Werf, K. O., Putman, C. A. J., Kraan, Y. M., van Hulst, N. F., et al. (1994). J. Struct. Biol., 112, 32. 45. Putman, C. A. J., de Grooth, B. G., Wiegant, J., van der Werf, K. O., van Hulst, N. F, et al. (1993). Cytometry, 14, 356. 46. Puppels, G. J., Putman, C. A. J., de Grooth, B. G., and Greve, J. (1992). Proc. SPIE, 1922, 145. 47. Mosher, C., Jondle, D., Ambrosio, L., Vesenka, J., and Henderson, E. (1994). Scanning Microsc., 8, 491. 48. Gall, J. G. and Pardue, M. L. (1969). Proc. Natl. Acad. Sci. USA, 63, 378. 49. Rudkin, G. T. and Stollar, B. D. (1977). Nature, 265, 472. 50. Wiegant, J., Wiesmeijer, C. C., Hoovers, J. M. N., Schuuring, E., d'Azzo, A., et al. (1993). Cytogenet. Cell Genet., 63, 73. 51. Moers, M. H. P., Kalle, W. H. J., Raap, A. K., de Grooth, B. G., van Hulst, N. F., et al. (1996). J. Microsc., 182, 40. 52. Proc. 3rd Int. Conf. on Near-field Optics, Brno, Czech Rep., May 1995 (ed. M. Paesler and N. F. van Hulst), Ultramicroscopy, Vol. 61, Dec. 1995. 53. Proc. 4th Int. Conf. on Near-field Optics, Jerusalem, Israel, Feb. 1997 (ed. N. F. van Hulst and A. Lewis), Ultramicroscopy, Vol. 71, March 1998.

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10 Introduction of materials into living cells 1. Particle bombardment as a means of DNA transfer into plant cells CHRISTIAN SCHOPKE and CLAUDE M. FAUQUET

1.1 Introduction Since its inception a decade ago, particle bombardment, also known as the 'Biolistic process' (1), has developed into a method of considerable importance in many areas of biology (see Table 1). It has been especially successful in the genetic transformation of plants, where the presence of a cell wall poses a significant obstacle for the introduction of large molecules, including DNA. The purpose of the first part of this chapter is to illustrate the use of particle bombardment for the introduction of DNA into plant cells and the utilization of reporter genes to follow the fate of genetically transformed cells, using cassava (Manihot esculenta) as an example. The reader who is interested in more detailed information on particle gun types and on different applications of particle bombardment is referred to several recently published reviews (2-4).

Table 1. Examples for the application of particle bombardment Target

Species

Mitochondria Saccharomyces cerevisiae Chloroplasts Tobacco Cyanobacteria" Synechococcus Fungi Paxillus involutes Plants More than 20 important crops Plants Cucurbitaceae Mammalian cells Various and tissues, live animals

Result

Reference

Stable transformation 5 Stable transformation 6 Stable transformation 7 Stable transformation 8 Stable transformation 9 Virus infection 10 Transient expression, stable 11 transformation, immunization

aInthis case bacterial magnetic particles were used for bombardment.

Christian Schopke, Claude M. Fauquet, and H. F. Paterson

1.2 Practical considerations The following discussion will focus on a particle gun type that has been used routinely by many laboratories, the particle delivery system PDS-1000/He (Bio-Rad) (see Figures 1 and 2). In this device pressurized helium is used to accelerate a macrocarrier onto which DNA coated microparticles have been applied. The accelerated macrocarrier then hits a stopping screen, permitting only the particles to continue their flight. The momentum imparted to the particles is such that they can penetrate plant cell walls (12). In a small proportion of cells that receive the particles (in the range of 100-2000 cells/cm2 of bombarded surface area), transient expression of the introduced gene can be observed. The number of cells that eventually incorporate the introduced DNA into chromosomal DNA is in the range of 0.1-1% of transiently expressing cells, 1.2.1 Parameters that affect the success of particle bombardment Numerous factors can have an influence on the outcome of a bombardment experiment (4, 13). If a new type of tissue is to be used for particle bombardment, a good strategy is to start with the basic settings given by the manufacturer of the PDS-1000/He (similar to Protocol 3, step 4) and to optimize

Figure 1. The particle delivery system PDS-1000/He. 374

10: Introduction of materials into living cells Before

After

Figure 2. Schematic representation of the PDS-1000/He system before and after activation (not to scale). A, B, and C are the adjustable distances that influence the velocity with which the microcarriers hit the target cells. (A) Rupture disk-macrocarrier distance. (B) Macrocarrier travel distance . (C) Target distance. The arrows indicate the direction of the helium flow (from ref. 12, modified with permission from Bio-Rad).

parameters, including bombardment pressure, distance between the barrel and tissue sample, number of bombardments per sample, and particle size. It is very important that the tissue is in a favourable physiological state to withstand the stress related to bombardment. In many cases culture of the tissue before and after bombardment on a medium containing elevated concentrations of osmotically active substances results in higher rates of transient gene expression. For a thorough discussion of factors that influence the efficiency of particle bombardment the reader is referred to refs 14 and 15. 1.2.2 Particle coating The basic protocol for coating microparticles with DNA consists of mixing a sterile suspension of tungsten or gold particles in water (see Protocol 1) with DNA, CaCl2, and spermidine. Numerous variations of this procedure have been employed, e.g. with differences in total amounts of DNA and particles used, the ratio of DNA to particles, the addition of buffers to the particle suspension, and the extent of the use of sonication and/or vortexing to mix the suspension. In the case of the PDS-1000/He, in the final step of the coating procedure the particles are resuspended in 100% ethanol. Aliquots of this suspension then are transferred to macrocarriers. After drying they are ready to be used for bombardment. At the International Laboratory for Tropical Agricultural Biotechnology (ILTAB) we routinely use with good results a protocol based on that developed by Sivamani et al. (16) (see Protocol 2). 375

Christian Schopke, Claude M. Fauquet, and H. F. Paterson Protocol 1. Sterilization of particles Equipment and reagents • Eppendorf microcentrifuge 5415C (Fisher Scientific Co.) • Microcentrifuge tubes, 1.7 ml, polypropylene, siliconized" (Sigma, T-3406)

• Gold particles (Bio-Rad or Alfa Chemicals) or tungsten particlesb (Bio-Rad)

Method 1. Weigh 30 mg particles into a microcentrifuge tube. 2. Add 500 u| 100% ethanol and vortex for 3 min at maximum speed. 3. Centrifuge for 10 sec at 10 000 r.p.m. 4. Remove supernatant and add 500 ul 100% ethanol. 5. Vortex for 2 min at maximum speed. 6. Repeat steps 3-5. 7. Centrifuge for 10 sec at 10000 r.p.m. 8. Discard supernatant and add 500 ul sterile distilled water. 9. Vortex for 2 min at maximum speed. 10. Centrifuge for 30 sec at 10 000 r.p.m. 11. Discard supernatant and add 500 u| sterile distilled water. The particles are now ready for coating.c a The use of siliconized tubes is recommended because the particles tend to stick to the surface of non-siliconized tubes during the following manipulations. bIn many publications on particle bombardment the source for tungsten particles is given as GTE (now Osram Sylvania, Danvers, Massachusetts). c Gold particles should be prepared the day they are to be used because they agglomerate irreversibly over time in aqueous suspension (4). Tungsten particles should not be stored for longer than one to two weeks as the oxidation of the surface of the particles may negatively affect the DNA binding capacity (4).

Protocol 2. Coating of particles with DNA Equipment and reagents • See Protocol 1 • Macrocarriers (Bio-Rad) . 2.5 M CaCI2

• 0.1 M spermidine, free base (Sigma, S 4139) • DNA at a concentration of 1 ug/ul in water

Method 1. Resuspend a sterile suspension of gold or tungsten particles (see Protocol 1, step 11) and transfer an aliquot of 50 uJ containing 3 mg particles to a siliconized microcentrifuge tube. While vortexing the 376

10: Introduction of materials into living cells

2. 3. 4. 5.

6. 7. 8.

open tube on a low setting, add 5 ul DNA solution (1 ug/uj). Vortex continuously through steps 2-3. Add 20 ul 0.1 M spermidine. Add 50 ul 2.5 M CaCI2, drop by drop. Let the particles settle for 10 min at room temperature. In the meantime sterilize the macrocarriers and the macrocarrier holders by leaving them for 10 min in 100% ethanol. Remove macrocarriers and macrocarrier holders from the ethanol, let them dry, and insert the macrocarrier in the holders. Store them in sterile Petri dishes until they are used. We routinely prepare ten holders at a time for use up to 1 h after preparation. Remove the supernatant from the settled suspension (step 4) and add 50 (uJcold 100% ethanol. Vortex at a low speed to resuspend the pellet. Distribute samples (usually in the range of 5-10 ul) onto the centre of the macrocarriers and let them dry in a desiccator. Leave them in a desiccator until they are used. While pipetting the suspension, it is important to maintain the pipette in a vertical position to ensure that the suspension spreads evenly on the surface of the macrocarrier.

1.2.3 The choice of a target tissue If the purpose of particle bombardment is transient expression of the introduced gene, for example to evaluate the expression of transcriptional promoters (17), any tissue that remains viable over the course of the experiment can be used as a target. However, if the production of stably transformed tissue is required, transformed cells must be able to undergo continuous cell division. While many plant cell types can resume mitotic activity and produce unorganized callus under appropriate conditions, the capacity to regenerate into whole plants usually is restricted to only a few cell types. The ideal target for the production of transgenic plants would be a leaf with an epidermis that contains a high percentage of single cells capable of regeneration. The flat shape would ensure that the surface of the tissue is at right angles to the beam of accelerated particles, and the target cells would be in the cell layer that is hit by particles with the highest efficiency. In the case of cassava, regeneration can be achieved via organized embryogenie structures (embryo clumps) derived from young leaves or via friable embryogenic tissue (maintained either as callus or as suspension culture) derived from somatic embryos. Both tissue types have been used as targets for particle bombardment for the purpose of obtaining transgenic cassava plants (18-20) (see Protocols 3-5). However, they differ in important aspects with regard to particle bombardment and to regeneration patterns. Embryo clumps are composed of tightly packed embryos at the globular to torpedo stage, i.e. 377

Christian Schopke, Claude M. Fauquet, and H. F. Paterson they have an epidermis and, depending on the developmental stage, a visible shoot pole. They can be induced either to grow into plants or, alternatively, to form secondary embryos (21). Secondary embryogenesis proceeds through a cleavage-like process in which many cells in layers below the epidermis are involved (22). On the other hand, tissue derived from embryogenic suspensions is composed mainly of small globular structures, 100-1000 u.m in diameter, which grow freely suspended in liquid medium. Secondary embryogenesis is induced in single cells on the surface of the globular structures (23). When this type of tissue is spread on a flat support such as filter paper, it comes close to the ideal target mentioned above: a layer of tissue with a high percentage of the cells on the surface capable of regeneration. Protocol 3. Particle bombardment of embryo clumps of cassava Equipment • Biolistic particle delivery system PDS-1000/ He (Bio-Rad)

• Micropipetters • Macrocarriers (Bio-Rad)

Method 1. Establish cultures of embryo clumps of cassava as described in ref. 21. 2. Three to four weeks after their last subculture, these clumps consist of more or less globular embryos in their centre, torpedo-shaped embryos at their margins, and often some non-embryogenic callus. Remove this callus and arrange pieces of embryo clumps consisting mainly of globular embryos in a circle of 2 cm diameter (about 30 clumps with a diameter of 3-6 mm) in the centre of a Petri dish with agar medium. It is important to choose clumps in which the embryos are arranged more or less horizontally in order to be accessible for the particles during bombardment. 3. Prepare DNA coated particles (gold, 1 um diameter) according to Protocol 2. 4. Choose the following bombardment parameters: (a) Distance macrocarrier/stopping screen: 6 mm (upper position of screen). (b) Distance rupture disc assembly/macrocarrier cover (gap): 1/4" (63.5 mm). (c) Suspension volume per bombardment: 7.5 ul. (d) Rupture disks: 650 psi. (e) Level of sample holder: third from below. 5. Place the Petri dish with the embryo clumps in the centre of the sample holder. Bombard the sample under a partial vacuum of 10-6.5 kPa (vacuum of 27-28 in Hg). 378

10: Introduction of materials into living cells Protocol 4. Preparation of tissue derived from embryogenic suspensions of cassava for particle bombardment Equipment • Polypropylene grids, autoclavable, opening 210 um (Spectrum Medical Industries, Inc.): the grids can either be purchased as circles (5 cm diameter, order no. 145 771) or as 30 cm2 sheets (order no. 146 428)

• 15 ml graduated polystyrene centrifuge tubes (Falcon, 2099)

Method 1. Establish an embryogenic suspension of cassava as described in ref. 23. Take a sample of 10 ml with a wide-mouth glass pipette or with a plastic pipette that has been cut with a hot scalpel blade to obtain a wide opening and transfer it to a 15 ml graduated centrifuge tube. 2. Leave the suspension undisturbed for 20 min and read the settled cell volume (SCV) of the tissue. Take an aliquot of 1 ml tissue and initiate a new suspension. Subculture every second day in fresh medium. 3. After 12-14 days, sieve the suspension to obtain the fraction of embryogenic cell clusters with a diameter between 250-500 um. 4. Place the grids in Petri dishes on top of a dry filter paper. Transfer aliquots of 200 ul SCV in a volume of 1 ml culture medium with a wide-mouth pipette onto the grids in such a way that the liquid forms a drop kept in place by surface tension. 5. Prepare the amount of dishes needed for one bombardment session. Including the time for particle coating and tissue preparation, approx. 40 samples can be bombarded in a period of 8 h. If each sample is bombarded twice, the amount of dishes is reduced correspondingly.

Protocol 5. Particle bombardment of tissue derived from embryogenic suspensions of cassavaa Equipment and reagents • Biolistic particle delivery system PDS1000/Helium (Bio-Rad) • 1100 psi pressure disks (Bio-Rad)

• DNA coated particles (see Protocol 7) .P|anttissue (see Protocol 2

Method 1. Use the bombardment parameters as described in Protocol 3, step 4. Use 1100 psi rupture disks and a volume of 5 uJ suspension of coated particles per bombardment.

379

Christian Schopke, Claude M. Fauquet, and H. F. Paterson Protocol 5.

Continued

2. Immediately before bombardment, bring the droplet with tissue on the grid in contact with the underlying filter paper. This can be done either by pressing down the grid with a forceps or by adding a drop of medium between the grid and the filter paper. As a result, the liquid is absorbed and the tissue remains as an evenly spread layer on the grid. If the grid has changed its position during the manipulations, move it back to the centre of the dish. 3. Place the Petri dish with the tissue on the sample holder inserted at the third level from below in the particle gun. Bombard the sample under a partial vacuum of 10-6.5 kPa absolute pressure (vacuum of 27-28 in Hg). 4. After the bombardment, transfer the grid with tissue to a Petri dish or any other suitable vessel with liquid culture medium. At the end of the bombardment session transfer the tissue to a culture flask. 5. If the sample is to be bombarded a second time, add enough medium to the dish so that the tissue is just covered. This ensures that it does not dry out during the preparations for the next bombardment. 6. For a second bombardment, transfer the grid to a Petri dish with a filter paper to absorb excess medium. Continue with steps 3 and 4. a Adapted from ref. 19.

1.3 Identification of cells transformed with reporter genes Several genes have been routinely employed in the context of particle bombardment with the purpose of visually identifying transgenic plant cells or tissues. The expression of these genes can be detected either directly through their coloured product or indirectly through a colour reaction catalysed by the gene product (see Table 2).

Table 2. Examples of reporter genes used in conjunction with particle bombardment Gene(s)

Origin

Gene product

Result of gene expression

Reference

uidA. luc Rand C1

E. coli Firefly Maize

Blue stain Light emission Expressing cells red

24 20 25

gfp

Jellyfish

B-glucuronidase (GUS) Luciferase (LUC) Enzymes involved in anthocyanin synthesis Green fluorescent protein (GFP)

Fluorescence

26

380

10: Introduction of materials into living cells 1.3.1 The B-glucuronidase gene (uidA) uidA is a gene isolated from Escherichia coll and encodes the enzyme (3glucuronidase (GUS), which can be detected by a colour reaction (Protocol 6). This enzyme catalyses the cleavage of B-glucuronides into glucuronic acid and the corresponding aglycone. The substrate used most frequently for histological purposes is 5-bromo-4-chloro-3-indolyl-p-D-glucuronide cyclohexyl-ammonium salt (X-gluc). The reaction with B-glucuronidase results in glucuronic acid and the colourless, water soluble 5-bromo-4-chloroindoxyl, which upon oxidative dimerization forms an insoluble blue stain. The localization of the blue stain depends on the diffusion of the water soluble monomer, 5-bromo-4-chloroindoxyl, and on the velocity of the dimerization reaction. The longer this reaction takes, the farther away from its place of origin the monomer can diffuse. This means that, depending on the assay conditions, the blue stain can be formed in cells adjacent to the cell that received the uidA gene and thus reduce the resolution of the assay. The histological GUS assay has been widely used for experiments to optimize microbombardment parameters. In the context of particle bombardment, the GUS assay can be used to determine the number of cells transiently expressing the uidA gene, and to optimize each experimental system. When embryogenie tissue of cassava was bombarded with uidA and analysed by standard GUS assay conditions (24), it was found that two to three days after bombardment large areas of tissue were stained light blue (27). With increasing concentrations in the assay buffer of equimolar mixtures of the oxidation catalysts, ferri- and ferrocyanide, the blue stain became more intense and restricted to smaller areas, and at a concentration of 6.4 mM each most of the stain was confined to single cells (Figure 3A, 3B, 3D). This, together with a clearing method that does not affect the blue stain (see Protocol 6, steps 4-6), increased the contrast between stained and non-stained cells sufficiently to permit computer-aided image analysis for the quantification of blue cells (27).

Protocol 6. Histological GUS assay of cassava tissues Equipment and reagents • • • •

Incubator (37°C) Slide warmer Desiccator and vacuum pump Assay buffer: 0.08 M sodium phosphate pH 7, 0.8 mM 5-bromo-4-chloro-3-indolyl-B-Dglucuronide cyclohexyl-ammonium salt (Xgluc; Biosynth), 0.16% Triton X-100 (Sigma), and 20% (v/v) methanol • Clearing solution (optional): 160 g chloral hydrate, 50 ml water pH 7

• Assay buffer A: for tissue from embryogenie callus or suspensions and embryos add 6.4 mM potassium ferrocyanide and potassium ferricyanide • Assay buffer B: for leaves, shoots, and roots reduce the concentrations to 0.64 mM potassium ferrocyanide and potassium ferricyanide • Phenolic glycerol gelatin (Sigma)

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Continued

A. Tissue from embryogenic callus or suspensions and embryos 1. Incubate tissue in assay buffer A for 2 h at 37°C. 2. Wash the tissue several times with water. 3. Observe the tissue immediately or store in 70% ethanol. 4. If a good contrast between blue cells and the surrounding tissue is desired, transfer the tissue to clearing solution. After three days, remove the clearing solution by washing the tissue several times in 50% glycerol (v/v). After the last wash blot the tissue on filter paper to remove excess liquid. 5. Transfer the tissue to a microscope slide and place it on a slide warmer heated to 40-45°C. Cover the tissue drop by drop with liquefied phenolic glycerol gelatin. 6. Carefully place a coverglass on the liquid glycerol gelatin and store the slide horizontally at room temperature. Samples prepared in this way have been kept for more than a year without noticeable change in stain intensity. B. Tissue from plants (leaves, shoots, roots) 1. Incubate the tissue in assay buffer B for 16 h at 37°C. Vacuum infiltrate the tissue to facilitate the penetration. 2. Proceed as described in part A, steps 3 and 4. If the tissue is green, the chlorophyll can be removed after washing the tissue with water by leaving it in 70% ethanol until it becomes white. The extraction of chlorophyll can be shortened by transfer to 100% ethanol, but in this case shrinkage of cells can occur.

GUS expression in cassava transformed with the uidA gene driven by the 35S promoter from cauliflower mosaic virus (CaMV) as detected by the histological GUS assay (see Protocol 6A) is highest in embryogenic and embryo tissues (Figure 3A-F). Most of the GUS positive cells are stained dark blue. When cassava tissues from regenerated, transgenic plants are subjected to the same GUS assay conditions, macroscopically they appear light blue (Figure 4A). However, microscopic examination reveals that some cell types stain with an intensity comparable to embryogenic and embryo cells. Without further analysis one might interpret GUS expression patterns as shown in Figure 4 as cell type-specific expression of guard cells (Figure 4B) or laticifers (Figure 4D). However, by using an assay buffer containing 1/10 of the concentration of ferri- and ferrocyanide (0.64 mM each), and by extending the time for assay from 2 h to 16 h, all cells stain blue to dark blue (Figure 4A and 4C). The conclusion from these findings is that the outcome of a histological GUS assay 382

10: Introduction of materials into living ceils

Figure 3. GUS expression in cassava cells and tissues after bombardment with a plasmid containing the uidA gene under the control of the CaMV 35S promoter. (A-C) Bombardment of embryo clumps. (A) Embryo clump, three days after bombardment. (B) Close-up of (A); single GUS-expressing cell. Several gold particles are visible. (C) Chimeric embryo six weeks after bombardment, (D-F) Bombardment of tissue derived from embryogenic suspensions. (D) Two days after bombardment. (E) Ten days after bombardment. (F) Transgenic embryos, ten months after bombardment. Bars in (A) 3 mm; (8} 5 um; (C) 2 mm; (D) 1 cm; (E) 100 um; (F) 2 mm. 383

Figure 4. (A-D) GUS expression in a cassava plant transformed with the uidA gene. (A) Longitudinal sections through the shoot tip. (B) and [C} Leaf epidermis, (D) Close-up of the section on the left shown in (A), depicting a lacticifer. The intensity of the stain during the GUS assay depends on the assay conditions. Tissues in (A (section on the left), B, and D) were subjected to a 2 h assay with 6.4 mM ferri- and ferrocyanide, tissues in (A (section on the right) and C) to a 16 h assay with 0.64 mM ferri- and ferrocyanide in the assay buffer. (E-G) Light emission in cassava tissues transformed with the luc gene. (E) Light-emitting embryogenic tissue of cassava three months after bombardment with the luc gene. Light emission was measured for 1 min, using an intensified VIM camera an Argus-50 image processor (Hamamatsu Photonics). (F) Regenerated transformed plantlet. (G) The same plantlet as shown in (F); light emission measured for 5 min. (H-l) Transient expression of the gfp gene in microbombarded rice tissue. (H) Tissue derived from embryogenic suspension cultures three days after bombardment. The photograph was taken with a UV light source and an FITC filter set. (I) Close-up of (H); single GFP-expressing cell. Bars in (A) 5 mm; (B, C, D) 100 (um; IE) 2 cm; (F, G) 1 cm; (H) 1 mm; (I) 30 um. Figures E-G were kindly provided by C.J.J.M. Raemakers; Figures H and I by A. de Kochko. Figure G with permission from Kluwer Academic Publishers. 384

10: Introduction of materials into living cells depends very much on the assay conditions, and that different incubation times and oxidation catalyst concentrations should be tested before interpreting the assay results. This is even more important when cell-specific promoters are used to control GUS expression. 1.3.2 Other reporter genes A major disadvantage of the histological GUS assay described above is that it is usually lethal to plant tissues and therefore cannot be used for the visual selection of transgenic tissue with the purpose of further cultivation and regeneration. Other reporter genes that have been used in conjunction with particle bombardment of plant tissues and whose expression can be detected in vivo are the genes for firefly luciferase (luc), genes involved in anthocyanin biosynthesis (R and Cl), and green fluorescent protein (gfp) (see Table 2). Firefly luciferase (LUC) is an enzyme that catalyses a reaction that results in the production of visible light. Apart from being helpful for visualizing transformed cells it also can be used for their selection. It is known that the antibiotics used for the selection of transgenic tissues after transformation with antibiotic resistance genes can have negative effects on the regeneration capacity of certain transformed tissues. LUC therefore offers a selection strategy that avoids this problem. The detection of LUC requires only the substrate, luciferin, as well as ATP, Mg2+ (both present in the plant tissue), and O2. Living plant that tissues express LUC genes after exposure to luciferin produce light at a wavelength of 560 nm which can be recorded by low light level detectors (e.g. CCD cameras), and quantified by image processing systems. Raemakers et al. (20) used this approach to select transgenic tissue of cassava. In Figure 4E-G examples for the detection of LUC activity are shown. Tissue derived from embryogenic suspensions of cassava was bombarded with a plasmid containing the firefly luciferase coding sequence controlled by the cauliflower mosaic virus (CaMV) 35S promoter. Repeated selection and subculture of light-emitting tissue resulted in an enrichment of transformed tissue (Figure 4E), from which eventually plantlets were regenerated (Figure 4F and 4G). Despite the advantage of a marker whose expression can be followed in living tissue, the use of the firefly luciferase gene to identify transformed tissue is not as widespread as the use of the GUS gene. The reason is the relatively high cost for the detection equipment. Recently it was shown that a luciferase coded by a gene isolated from another organism, the marine soft coral Renilla reniformis, produces a stronger light emission, and is more stable than the firefly luciferase (28). This luciferase has the potential of being detectable with less costly equipment and might become a useful marker gene in the future. Genes encoding transcriptional activators that control anthocyanin synthesis in maize have been used to identify transiently or stably transformed cells by their red coloration (25). In this case neither a substrate nor special equipment are needed to identify transformed cells. In addition, the red stain is 385

Christian Schopke, Claude M. Fauquet, and H. F. Paterson restricted to the cell expressing the introduced gene(s), i.e. there is no problem of diffusion to neighbouring cells as is the case with cells expressing GUS. Limitations that restrict the use of these genes as reporter genes are the fact that their expression can depend on the presence or absence of other genes in the anthocyanin biosynthesis pathway; as a result they may be useful only in certain plants (e.g. maize). In addition, activation of the pathway can interfere with developmental processes: constitutive expression of the R and Cl genes in sugar cane plantlets did not allow survival of anthocyanin pigmented plants taller than 3 cm (29). Another marker gene that can be detected in vivo is gfp, which encodes the green fluorescent protein (GFP) from the jellyfish Aequorea victoria. This protein fluoresces green when excited by ultraviolet (395 nm) or blue light (490 nm). The green fluorescence can be observed with a UV microscope equipped with a FITC filter set or with a hand-held ultraviolet light. Variants of this protein with different absorption and emission spectra have been developed by changing the gene coding sequence (26). In transformation experiments with the native gfp, low expression levels of GFP were observed in plant cells due to cryptic splice sites and poor codon usage. However, recent work with codon optimized versions of the gene has shown that expression levels can be achieved that are sufficient for GFP detection in planta (30). At ILTAB preliminary experiments have been performed using a mutated version of GFP (mutation S65T) (26) in conjunction with microbombardment of embryogenic tissue of rice (31). Figure 4H and 4I show transient expression of GFP, two days after bombardment.

2. Microinjection as a preparative technique for microscopical analysis H. F. PATERSON

2.1 Advantages of microinjection by glass capillary needle Although there are now numerous, widely-used methods available for introducing exogenous substances into mammalian cells in culture (see Chapter 6), no technique has proved itself so well-suited or adaptable for subsequent microscopic analysis as direct microinjection by glass needle. Not only are there few constraints on the molecular size or nature of the substances which can be introduced in this fashion, but also the intracellular site (nucleus or cytoplasm) and the quantity of the injected material can be controlled and recorded on a cell-by-cell basis. Adherent cells growing on a tissue culture substratum are microinjected in situ with minimal disruption or damage, allowing unhindered observation of rapidly occurring responses to the in386

10: Introduction of materials into living cells jected material. Additionally, the ability to microinject discrete groups of cells in adjacent areas of the culture dish with experimental and control substances allows subsequent comparison of their effects within the same microscopical field. Microinjection is applicable to all mammalian cells which grow (or can be induced to grow) as an adherent monolayer culture, and success is far less influenced by cell type than most other methods. Where substances are microinjected directly into the cytoplasm, efficiency of delivery may approach 100%, and intracellular concentrations > 5 mg/ml of the injected substance are obtainable if required. Efficiency of expression of microinjected DNA is more variable, since successful introduction of plasmid into the cell nucleus does not guarantee gene expression. As with all methods of DNA delivery, good gene expression following microinjection is dependent on the choice of a suitable vector.

2.2 Equipment required for microinjection of adherent mammalian cells Microinjection may be performed with relatively simple equipment. Basic requirements are for a microscope through which the cells and the injection procedure may be observed, together with some form of micromanipulator which allows the operator fine control over the movement of the injection needle. Access to a needle pulling device is also necessary, enabling consistentlyshaped needles to be produced from capillary tubes. Finally, a source of variable air or hydraulic pressure is needed to force the injection solution from the needle into the cell. 2.2.1 Microscopes A compound microscope forms the basis of any microinjection system. This should be equipped with at least three objective lenses, enabling the injection needle to be gradually manoeuvred to the centre of the field by progressive increases in magnification. A typical system employs X 5, X 20, and X 40 objectives, in conjunction with X 10 eyepieces. Phase-contrast is generally found to be the most useful optical system for rendering the cells and their internal structures visible during the injection process, being relatively inexpensive, straightforward to use, and robust in operation. Alternatively, DIC or Nomarski optics provide unrivalled perception of depth and detail within the specimen, but suffer the disadvantage of being unsuitable for use with plastic tissue culture vessels. Inverted microscopes are the most easily adapted for microinjection usage, since their design is ideally suited to working with live cells in culture dishes. When fitted with long working distance condensers, such microscopes permit unrestricted access of the injection needle into the culture dish. Conventional upright microscopes may be used (Figure 5a), but have several shortcomings 387

Christian Schopke, Claude M. Fauquet, and H. F. Paterson

Figure 5, Equipment required for microinjection. (a) Basic manual system employing an upright microscope, Leitz mechanical micromanipulator (m) with integral hanging joystick (j), and 50 ml syringe as pressure source (s). Note 45° angle of needle holder (n). (b) Zeiss semi-automatic microinjection workstation employing an inverted microscope with incubator jacket, and Eppendorf 5170 stage-mounted electrically operated micromanipulator (m) with remote upright joystick (j).

by comparison with inverted types. Not only is there little room between the culture dish and the objective lens for positioning the injection needle, but the meniscus formed around the needle as it passes through the surface of the culture medium, beneath the objective, and close to the optical axis, may cause degradation of the image quality. A further problem with the majority of upright microscopes is that focusing is achieved by moving the height of the stage relative to the stationary objective turret. This design feature renders the microscope incompatible with separately mounted micromanipulators, since any attempt at refocusing during microinjection will cause the cells to move in the vertical axis relative to the needle. An optional refinement for the microscope is the provision of a regulated environment for the cells during lengthy microinjection sessions. This normally takes the form of a heated stage (incorporating a suitable recess to accept the culture dish) surrounded by a transparent incubator jacket equipped with adjustable temperature and CO2 control, as in Figure 5b. Since a sizeable hole must be left open in the incubator jacket for access to the cells by the micromanipulator, the control system should be of high capacity and capable of constantly monitoring and adjusting the internal atmosphere. 388

10: Introduction of materials into living cells 2.2.2 Micromanipulators Several designs of micromanipulator are commercially available, employing mechanical, electrical, or hydraulic principles of operation. In most models a hand-operated joystick controls movement of the needle in the horizontal plane (x and y axes), reduced proportionately by a factor of up to 200-fold (usually adjustable). Typically, one centimetre of movement at the joystick will cause the injection needle tip to traverse a distance roughly equivalent to the diameter of a large adherent cell. Vertical (z axis) movement of the injection needle is achieved in many systems by twisting the joystick around its own axis, although in some designs rotation of a separate wheel is required. Sophisticated models may incorporate a feature which combines vertical and lateral movements in order to effect an axial, spearing motion of the microinjection needle. Coarse controls are also provided to facilitate comparatively large, rapid movements during initial setting up and changing of needles. These will often include a simple mechanism for swinging the needle into and out of the culture medium over the high side of the dish. Detailed discussion of different micromanipulator designs is beyond the scope of this section, but the following points are worth considering. Mechanical micromanipulators have the advantage of robust simplicity, but the directness of the linkage renders them prone to the transmission of incidental knocks and vibrations from the operator's hand through to the needle. With free-standing mechanical units such as the Leitz (Figure 5a), it is also imperative to use a fixed-stage microscope. Hydraulically operated and electrically driven systems allow flexible linkage, so that the needle holder and its drive unit can be mounted directly on the microscope stage while the joystick controls are separately sited on the base plate or table top. This arrangement keeps the transmission of unwanted vibration to a minimum. A further advantage of electrically driven systems is their potential for computerized control, permitting automatic or semi-automatic microinjection of cells selected by the operator (Figure 5b). 2.2.3 Needle pullers A mechanized puller is required to produce consistently-shaped needles from glass microcapillaries. Several devices are commercially available, although a simple machine can be constructed in the laboratory workshop. With the most common design a straight-walled capillary tube, encircled at its mid-point by a small electrical heating coil, is held under tension at both ends (Figure 6a). When the heating coil is made red hot, the softened length of glass in the adjacent middle section of the capillary is drawn out and eventually pulled apart into two identical sharp points. The shape of the twin needles thus formed can be controlled by altering the temperature of the heating coil and by adjusting the tension force on the ends of the capillary. Most modern machines allow a two-stage pull to be applied to the capillary, thereby giving 389

Christian Schopke, Claude M. Fauquet, and H. F. Paterson

Figure 6. Needles for microinjection. (a) Central section of a Narishige PE-2 needle puller, showing close-up of capillary held under tension (c), and heating:coil (h). (b) Comparison of capillary tubing used to make microinjection needles (c), finished needle loaded with injection solution (n), and tip of Eppendorf microloader (ml). Note the internal filament (arrowed) in capillary and needle. Bar = 1 mm.

greater control over the dimension and taper of the needle tips. Initial gentle stretching of the heated region to a predetermined length, in order to narrow the capillary diameter, triggers the application of a stronger secondary pull which forms the final taper.

2.2.4 Pressure control unit Means of applying sufficient pressure to force the injection solution through the minute hole in the tip of the microcapillary needle must be provided. The needle is firmly gripped by an airtight rubber seal in the needle holder, which is connected to a suitable source of air pressure via a length of small-bore, flexible, pressure tubing. For simple, non-automatetE microinjection, a large hand-held syringe provides a surprisingly capable and continuously variable pressure source (see Figure 5a). More advanced systems employ a source of compressed air connected to a series of valves, which allow pre-set pressures to be delivered to the needle. The Eppendorf System 5242 incorporates three adjustable control valves for use with automated or semi-automated systems of microinjection. The first is designed to deliver a low, continuous pressure which counteracts the tendency of the needle to draw up liquid from the culture medium by capillary attraction. The second valve is operated either by 390

10: Introduction of materials into living cells foot switch (semi-automated systems) or computer (fully automated systems) to deliver the short pulse of higher pressure required to inject the cells. The third valve can be set to pass full pressure from source, and is intended primarily as a means of clearing blockages in the needle.

2.3 Preparation of materials for microinjection 2.3.1 Cell cultures Cells for microinjection may be seeded either onto plastic dishes or glass coverslips. Standard tissue culture Petri dishes of 50-60 mm diameter are convenient for most purposes, and adaptors to hold these dishes snugly on the microscope stage are available from microscope manufacturers. Smaller dishes (30 mm diameter) may cause difficulty of access for the injection needle. Glass coverslips must be cleaned of grease and powdered glass before use by washing in absolute alcohol. After sterilization by flaming, they are placed in Petri dishes and seeded with cells in the normal way. Only firmly adherent cells are suitable for microinjection. Dishes precoated with poly-lysine, collagen, fibronectin, or laminin such as the 'BioCoat' range from Collaborative Biomedical Products will often improve the adhesion of loosely attached cell types. 2.3.3 Capillaries and needles Mechanical pulling devices constitute the only reliable way to create reproducibly-shaped microinjection needles from glass capillaries. Optimal settings of such machines must be determined by trial and error. Capillary tubes suitable for making needles contain a fine glass filament running along the entire length of the inner wall (arrowed in Figure 6b) which has the important property of drawing the injection solution down to the very tip of the needle without airlocks. Needles with a taper such as shown in Figure 6b are slender enough in use to produce minimal interference with the optics of the injection microscope, sufficiently rigid to be moved through the culture medium without flexing, and consistently open at the tip. The high temperature involved in pulling the capillaries renders the needles sterile and ready for use. Protected from dust, they may be stored for several weeks until required. Needles are loaded with injection sample immediately before use by means of very long, fine, micropipette tips (e.g. Eppendorf 'microloaders'), sufficiently narrow in diameter to be inserted into the shaft of the needle (see Figure 6b). 2.3.2 Marking the area of cells to be injected Several million adherent cells can be accommodated on the surface of a culture dish, and even a small coverslip will support many thousands. Therefore it is important that the position of microinjected cells on the culture surface should be marked so that they can be easily relocated for analysis. When plastic Petri dishes are used, this can be done after cells have been plated. A region 391

Christian Schopke, Claude M. Fauquet, and H. F. Paterson

Figure 7. Microinjection techniques, (a) Injection into the nucleus of an epithelial cell, showing the phase-bright halo around the tip of the needle (arrowed). Bar = 30 um. (b) A single field of mouse fibroblasts divided into four quadrants by scoring a cross in the surface of the culture dish. The cells in each quadrant have been microinjected with different recornbinant ras proteins, allowing direct comparison of their effect on morphology. Bar = 200 um.

of the dish should be selected where the cells are evenly distributed and free of clumps or other anomalies. With the aid of a low power inverted microscope, two lines forming a cross are scored on the upper (tissue culture) surface of the dish using a sterile scalpel blade, thereby providing four adjacent quadrants of cells for injection (Figure 7b). More elaborate box or grid patterns may be fashioned according to the demands and complexity of the experiment. This technique has the advantage of ensuring that the marking lines are in the same focal plane as the cells, permitting both to be resolved simultaneously under high magnification where depth of field is very limited. The ridge of plastic thrown upwards by the scalpel blade also constitutes a useful physical barrier to cell movement between adjacent marked areas. Marking coverslips by hand is less precise, since a diamond-point scriber or similar instrument must be used to scratch suitable markings on the glass surface prior to seeding the cells. More satisfactory are the pre-sterilized 'Cellocate' coverslips from Eppendorf which are embossed in the centre with a labelled grid to allow a wide choice of well-defined, easily relocated areas of cells for microinjection.

2.4 Microinjection technique A step by step procedure for microinjecting cells is given in Protocol 7, modified from the technique of Graessmann and Graessmann (32). The description is confined to manual injection which can be used with the most simple 392

10: Introduction of materials into living cells equipment. Those intending to employ automated systems should follow the guide-lines provided by the manufacturer. In this basic form of microinjection the tip of the loaded needle is maintained at an angle of 45 degrees to the surface of the culture dish, and is lowered vertically into the cell with a slicing movement similar to cutting a cake. The needle contents, which are kept under sufficient air pressure to maintain a continuous stream from the tip, will begin to flow into the cell. This is evidenced by swelling of the injected compartment, and movement of intracellular structures (Figure 7d). Experience will guide the operator in judging when to raise the needle out of the cell in order to complete the injection process. Under phase-contrast optics any cells which have been killed by over-injection become evident within a few minutes, exhibiting dark shrunken nuclei and coarsely speckled cytoplasm. For most cells, injection into the cytoplasm is better tolerated than into the nucleus. No fixed figures can be given concerning injection pressure or duration, as this is dependent on the aperture of the needle tip, the viscosity of the injection solution, and the type of cell. However, for good viability the pressure should be high enough to keep injection duration to less than half a second. Many cell types will survive injection of up to 10% of their own volume, although smaller quantities are desirable to avoid unwanted stress. Graessmann and Graessmann (32) have determined mean injection volume to be 1-2 X 10~u ml/cell. Up to 500 cells per hour may be microinjected using this technique. Protocol 7. Technique for manual microinjection of adherent mammalian cells in culture Equipment and reagents • Compound microscope equipped with x 5, x 20, and x 40 objectives and phase-contrast optics • Micromanipulator, with 1.2 mm gauge needle holder . Pressure source: 50 ml syringe, or compressed air with pressure control unit • Adjustable micropipette and 'microloader' tips (Eppendorf) mm

• Needles pulled from glass capillaries with internal filament, 100 mm long x 1.2 mm external diameter x 0.69 mm internal diameter (GC120F-10, Clarke Electromedical Instruments) • Injection sample, prepared in Dulbecco's phosphate-buffered saline (PBSa) • Adherent cells cultured in marked 50 or 60 diameter Petri dishes

Method 1. Place the prepared Petri dish, containing normal culture medium, on the microscope stage. Locate the marked area of cells to be injected, and bring into focus with the x 5 phase-contrast objective. 2. Draw 2-3 ul of injection sample solution into a microloader, and insert carefully into the shaft of a microinjection needle from the back-end. Dispense 0.5-1 ul of the sample into the tapered end of the needle, ignoring any bubbles which may form. 393

Christian Schopke, Claude M. Fauquet, and H. F. Paterson Protocol 7. Continued 3. Insert the rear half of the loaded needle into the needle holder, and tighten the securing ring to ensure a firm, airtight seal. Clamp the assembly into the micromanipulator, ensuring that the needle points downwards at an angle of 45° to the surface of the culture dish (see Figure 5a). 4. Use the coarse controls of the micromanipulator to swing the needle over the side of the dish, and guide it towards the central, illuminated area. 5. Lower the needle into the culture medium. Using the X 5, x 20, and x 40 objectives sequentially, bring the needle tip to the exact centre of the microscope field, resting in partial focus above the cell monolayer. Reset the joystick to the centre of its travel. 6. Deliver a burst of high pressure to the needle in order to initiate the flow of injection solution. Adjust the pressure to give light continuous flow (~ 100-400 hPa on a pressure regulator, or light finger pressure on a 50 ml syringe). 7. Select a group of cells for practice, just outside the marked experimental area, and position the needle directly above a test cell by means of the joystick. Lower the tip vertically until the presence of a small, phasebright halo indicates contact with the plasma membrane (Figure 7a). Lower the tip further until the cytoplasm or nucleus is penetrated, hesitate until a slight swelling reveals that injection has occurred, then swiftly withdraw the needle up to the resting position. 8. Examine the cell for signs of over-injection (see text), and adjust the needle pressure accordingly. Perform further test injections, on adjacent cells, until satisfied with viability. 9. Return to the marked area, and microinject as many cells as required for the experiment.

2.5 Microscopical analysis of microinjected cells Analysis of microinjection experiments is almost invariably performed microscopically, since the number of injected cells rarely constitutes sufficient material for biochemical techniques. In most cases groups of microinjected cells are examined either by conventional or time lapse photomicroscopy for changes in behaviour and morphology, or fixed and analysed using standard immunofluorescence protocols with antibodies against injection-derived or associated endogenous proteins. Where antibodies are not available against the injected material, it is often helpful to incorporate a suitable inert marker into the injection solution, such as 1 mg/ml purified immunoglobulin, which is retained within the cell and can be stained to identify injected cells during 394

10: Introduction of materials into living cells

Figure 8. Analysis by fluorescence microscopy in living cells of the intracellular localization of raf-GFP fusion protein, 16 h after microinjection of DNA vector. (a) Live cells expressing raf-GFP alone exhibit cytosolic localization of the protein, (b) Co-expression with oncogenic ras (not visible) causes translocation of raf-GFP to the plasma membrane. x 40 water immersion objective with standard FITC fluorescence filter set. Bar = 20 um.

subsequent analysis. An important technique for analysis is that of 'epitopc tagging', where the DNA coding sequence of a gene is modified such that the protein expressed contains an additional small polypeptide 'tag' corresponding to the binding site of a pre-existing, high affinity antibody. Immunostaining is performed against the tag, obviating the need to raise and characterize antibodies specific for the original protein. An additional advantage is that the localization of exogenous proteins containing experimental mutations can be analysed in isolation from the endogenous wild-type pool (33). A recent development in protein tagging has been the discovery of green fluorescent protein (GFP), an intrinsically fluorescent polypeptide, variants of which can be detected with fluorescence filter sets routinely used for fluorescein (FITC) immunostaining. When the DNA coding sequence for GFP is suitably incorporated into a gene of interest, and introduced into cells via an appropriate expression vector, the resultant fused protein can be detected in vivo by fluoreseenee imaging as shown in Figure 8. Using CCD video fluorescence microscopy, the intracellular localization and movements of GFP fusion proteins expressed from microinjected plasmids can be observed and recorded in living cells over extended periods of time (34), 395

Christian Schopke, Claude M. Fauquet, and H. F. Paterson

Acknowledgements The authors (C. S. and C. M. F.) wish to thank Dr R. N. Beachy (The Scripps Research Institute, La Jolla, USA) for his critical review of their manuscript.

References 1. Sanford, J.C., Klein, T.M., Wolf, E.D., and Allen, N. (1987). Part. Sci. Technol, 5, 27. 2. Ning-Sun Yang, N.-S. and Christou, P. (ed.) (1994). Particle bombardment technology for gene transfer. Oxford University Press, New York. 3. Gray, D.J. and Finer, J.J. (ed.) (1993). Plant cell tissue organ culture, 33, 221. (Special section on particle bombardment.) 4. Sanford, J.C., Smith, F.D., and Russel, J.A. (1993). In Methods in enzymology (ed. R. Wu), Vol. 217, p. 483. Academic Press, London. 5. Belcher, S.M., Perlman, P.S., and Butow, RA. (1994). In Particle bombardment technology for gene transfer (ed. N.-S.Yang and P. Christou), p. 101. Oxford University Press, New York. 6. Svab, Z. and Maliga, P. (1993). Proc. Natl. Acad. Sci. USA, 90, 913. 7. Takeyama, H., Yamazawa, A., Nakamura, C., and Matsunaga, T. (1995). Biotechnol. Techn., 9, 335. 8. Bills, S.N., Richter, D.L., and Podila, G.K. (1995). Mycol. Res., 99, 557. 9. Christou, P. (1996). Trends Plant Sci., 1, 423. 10. Gal-On, A., Meiri, E., Huet, H., Hua, W.J., Raccah, B., and Gaba, V. (1995). J. Gen. Virol.,76, 3223. 11. Yang, N.-S. and Ziegelhoffer, P.R. (1994). In Particle bombardment technology for gene transfer (ed. N.-S. Yang and P. Christou), p. 117. Oxford University Press, New York. 12. Reiser, W. (1992). Optimisation of biolistic transformation using the helium-driven PDS-1000-He system. Bio-Rad Bulletin 1688, Bio-Rad Life Science Group. 13. Sanford, J.C., DeVit, M.J., Wolf, E.D., Russel, J.A., Smith, F.D., Harpending, P.R., et al. (1991). Technique, 3, 3. 14. Southgate, E.M., Davey, M.R., Power, J.B., and Marchant, R. (1995). Biotechnol. Adv., 13, 631. 15. Birch, R.G. and Bower, R. (1994). In Particle bombardment technology for gene transfer (ed. N.-S. Yang and P. Christou), p. 3. Oxford University Press, New York. 16. Sivamani, E., Shen, P., Opalka, N., Beachy, R.N., and Fauquet, C.M. (1996). Plant Cell Rep., 15, 322. 17. Russell, D.A. and Fromm, M.E. (1995). In Gene transfer to plants (ed. I. Potrykus and G. Spangenberg), p. 118. Springer-Verlag, Berlin, New York. 18. Schopke, C., Chavarriaga, P., Mathews, H., Li, G.-G., Fauquet, C., and Beachy, R.N. (1993). In Vitro Cell. Dev. Biol., 29A, 64A. 19. Schopke, C., Taylor, N., Carcamo, R., Konan, N.K., Marmey, P., Henshaw, G.G., et al. (1996). Nature Biotechnol., 14, 731. 20. Raemakers, C.J.J.M., Sofiari, E., Taylor, N., Henshaw, G., Jacobsen, E., and Visser, R.G.F. (1996). Mol. Breeding, 2, 339. 396

10: Introduction of materials into living cells 21. Szabados, L., Hoyos, R., and Roca, W.M. (1987). Plant Cell Rep., 6, 248. 22. Raemakers, C.J.J.M., Jacobsen, E., and Visser, R.G.F. (1995). In The cassava biotechnology network; proceedings of the second international scientific meeting, Bogor, Indonesia, 22-26 August 1994 (ed. W.M. Roca and A.M. Thro), Vol. I, p. 336. CIAT, Cali, Colombia. 23. Taylor, N.J., Edwards, M., Kiernan, R.J., Davey, C., Blakesley, D., and Henshaw, G.G. (1996). Nature Biotechnol, 14, 726. 24. Jefferson, R.A. (1987). Plant Mol. Biol. Rep., 5, 387. 25. Bowen, B. (1992). In GUS protocols: using the GUS gene as a reporter of gene expression (ed. S.R. Gallagher), p. 163. Academic Press, San Diego. 26. Cubitt, A.B., Heim, R., Adams, S.R., Boyd, A.E., Gross, L.A., and Tsien, R.Y. (1995). Trends Biochem. Sci, 20, 448. 27. Schopke, C., Taylor, N.J., Carcamo, R., Beachy, R.N., and Fauquet, C. (1997). Plant Cell Rep., 16, 526. 28. Mayerhofer, R., Langridge, W.H.R., Cormier, M.J., and Szalay, A.A. (1995). Plant J.,7, 1031. 29. Bower, R., Elliott, A.R., Potier, B.A.M., and Birch, R.G. (1996). Mol. Breeding, 2, 239. 30. Leffel, S.M., Mabon, S.A., and Stewart, C.N. (1997). Biotechniques, 23, 912. 31. Zhang, S., Chen, L., Qu, R., Marmey, P., Beachy, R.N., and Fauquet, C.M. (1996). Plant Cell Rep., 15, 465. 32. Graessmann, M. and Graessmann, A. (1983). In Methods in enzymology (ed. R. Wu, L. Grossman, and K. Moldave), Vol. 101, p. 482. Academic Press, London. 33. Paterson, H., Adamson, P., and Robertson, D. (1995). In Methods in enzymology (ed. W. Balch, C. Der, and A.Hall), Vol. 256, p. 162. Academic Press, London. 34. Kaether, G. and Gerds, H.-H. (1995). FEBS Lett., 369, 267.

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11 Surface fluorescence microscopy with evanescent illumination D.AXELROD

1. Fluorescence at surfaces The interaction of molecules with surfaces is central to numerous phenomena in biology: e.g. binding to and triggering of cells by hormones, neurotransmitters, and antigens; the deposition of proteins upon foreign surfaces leading to thrombogenesis; electron transport in the mitochondrial membrane; cell adhesion to surfaces; concentration of reactants at surfaces and consequent enhancement of their reaction rates with other surface bound molecules; the dynamical arrangement of submembrane cytoskeletal structures involved in cell shape, motility, and mechanoelastic properties; and molecular events preceding cellular secretion from intracellular granules. In most of these examples, certain functionally relevant molecules coexist in both a surface-associated and non-associated state. If such molecules are detected by a conventional fluorescence technique (such as epi-illumination in a microscope), the fluorescence from surface-associated molecules may be dwarfed by the fluorescence from non-associated molecules in the adjacent detection volume. As an optical technique designed to overcome this problem, total internal reflection fluorescence (TIRF) allows selective excitation of just those fluorescent molecules in close (~ 100 nm) proximity to the surface. One note of caution: TIRF generally involves the use of a laser (with green or blue colour) for fluorescence excitation. Appropriate care should be taken to protect the eyes against exposure to direct or reflected laser light by the use of attenuating goggles when possible.

1.1 TIRF for biochemical samples TIRF can be used quantitatively on non-microscopic or featureless samples to measure concentrations of fluorophores as a function of distance from the substrate, or to measure binding/unbinding equilibria and kinetic rates at a biological surface. This type of study can be done with a custom TIRF configuration in a commercial spectrofluorimeter, but it is just as convenient and

D. AxeIrod

Figure 1. Three views of a mouse BC3H1 smooth muscle cell micro injected with rhodamine dextran: (a) TIR, using configuration E (Section 3.3); (b) epi-illumination; and {c} phasecontrast. The objective was a Zeias x 40 water immersion, NA - 0.75 achromat, used on a Leitz Diavert microscope. The images were recorded digitally on a 576 l. 384 pixel slow scan cooled CCD camera ( S t a r , Photometries Inc., Tucson, AZ), The image was then displayed on a VGA monitor screen using custom FORTRAN-based software and photographed onto standard 35 mm film. 400

11: Surface fluorescence microscopy with evanescent illumination much more light-efficient to use a microscope adapted for TIRF. Recently, single molecule fluorescence in biochemical systems has been detected by reducing extraneous fluorescence from out-of-focus planes, notably by TIRF.

1.2 TIRF for biological samples As applied to biological cell cultures observed through a microscope, TIRF allows selective visualization of cell/substrate contact regions, even in samples in which fluorescence elsewhere would otherwise obscure the fluorescent pattern in contact regions. TIRF can be used qualitatively to observe the position, extent, composition, and motion of these contact regions. Figure 1 shows an example of TIRF on a intact living cells in culture, compared with standard epifluorescence and phase-contrast. Since the cells are exposed to excitation light only at their cell/substrate contact regions but not through their bulk, they tend to survive longer under observation. This makes TIRF time lapse movies quite feasible, even over periods of days. TIRF is easy to set up on a conventional upright or inverted microscope with a laser light source or, in a special configuration, with a conventional arc source. TIRF is completely compatible with standard epifluorescence, bright-field, dark-field, or phase-contrast illumination; these methods of illumination can be switched back and forth readily. Since the illumination system can be 'chopped' between TIRF and standard through the lens 'epi' illumination, fluorescence changes at the submembrane and deeper in the cytoplasm can be simultaneously recorded and compared.

2. Theory of TIRF TIRF is conceptually simple. An excitation light beam travelling in a solid (e.g. a glass coverslip or tissue culture plastic) is incident at a high angle 0 upon the solid/liquid surface to which the cells adhere. That angle 6, measured from the normal, must be large enough for the beam to totally internally reflect rather than refract through the interface, a condition that occurs above some 'critical angle'. TIR generates a very thin (generally less than 200 nm) electromagnetic field in the liquid with the same frequency as the incident light, exponentially decaying in intensity with distance from the surface. This field is capable of exciting fluorophores near the surface while avoiding excitation of a possibly much larger number of fluorophores farther out in the liquid.

2.1 Single interfaces: intensity and polarization 2.1.1 Infinite incident plane waves When a light beam propagating through a transparent medium 3 of high index of refraction (e.g. glass) encounters a planar interface with medium 1 of lower index of refraction (e.g. water), it undergoes total internal reflection for 401

D. Axelrod incidence angles (measured from the normal to the interface) greater than the 'critical angle'. The critical angle 0C for TIR is given by: where n1 and n3 are the refractive indices of the liquid and the solid respectively, and n - n1/n 3 . Ratio n must be less than unity for TIR to occur. (A refractive index n2 will refer to an optional intermediate layer to be discussed in Section 2.2.) For incidence angle 0 < 0C, most of the light propagates through the interface with a refraction angle (also measured from the normal) given by Snell's Law. (Some of the incident light internally reflects back into the solid.) For 0 > 0C, all of the light reflects back into the solid. However, even with TIR, some of the incident energy penetrates through the interface and propagates parallel to the surface in the plane of incidence. The field in the liquid, called the 'evanescent field' (or 'wave'), is capable of exciting fluorescent molecules that might be present near the surface. For an infinitely wide beam (i.e. a beam width many times the wavelength of the light, which is a good approximation for unfocused or weakly focused light), the intensity of the evanescent wave (measured in units of energy/area/ sec) exponentially decays with perpendicular distance z from the interface:

l0 is the wavelength of the incident light in vacuum. Depth d is independent of the polarization of the incident light and decreases with increasing 0. Except for 0 —> 0C (where d -» °°), d is in the order of l0 or smaller. A more complete mathematical description of the electric and magnetic fields of the evanescent wave produced by infinite incident plane waves can be found in ref. 1. A physical picture of refraction at an interface shows TIR to be part of a continuum, rather than a sudden new phenomenon appearing at 6 = 0C. For small 0, the refracted light waves in the liquid are sinusoidal, with a certain characteristic period noted as one moves normally away from the surface. As 0 approaches 0C, that period becomes longer as the refracted rays propagate increasingly parallel to the surface. At exactly 0 = 0C, that period is infinite, since the wavefronts of the refracted light are themselves normal to the surface. This situation corresponds to d = °°. As 0 increases beyond 0C , the period becomes mathematically imaginary; physically, this corresponds to the exponential decay of Equation 2. The polarization of the electric field of the evanescent wave depends on the incident light polarization, which can be either 's' (polarized normal to the plane of incidence formed by the incident and reflected rays) or 'p' (polarized in 402

11: Surface fluorescence microscopy with evanescent illumination

Figure 2. Polarization snapshot of the electric field for p-polarized (in-plane arrows) and s-polarized (dots) light at a TIR surface. 8 refers to the non-zero phase shift between the incident light at the TIR surface and the evanescent wave.

the plane of incidence). For s-polarized incident light, the evanescent electric field vector direction remains purely normal to the plane of incidence. For ppolarized incident light, the evanescent electric field vector direction remains in the plane of incidence, but it 'cartwheels' along the surface with a non-zero longitudinal component (see Figure 2). This feature distinguishes evanescent light from freely propagating subcritical refracted light, which has no longitudinal component. The longitudinal component approaches zero as the incidence angle is reduced from the supercritical range back toward the critical angle. Regardless of polarization, the spatial period of the evanescent electric field is lo/(n3 sin 0) as it propagates along the surface. Unlike the case of freely propagating light, the evanescent spatial period is not at all affected by the medium 1 in which it resides. It is determined only by the spacing of the incident light wavefronts in medium 3 as they intersect the interface. The product of the evanescent electric field E with its complex-conjugate E* is proportional to the probability rate of energy absorption by a fluorophore in the evanescent wave. This product is the 'intensity' I in Equation 2. Given corresponding incident intensities I'p,s, the evanescent intensities Ip>s at z = 0 are:

Intensities I p,S (0) are plotted versus 0 in Figure 3, assuming the incident intensities in the glass I'p>s are set equal to unity. The evanescent intensity approaches zero as 0 -» 90°. On the other hand, for supercritical angles within ten degrees of 6C, the evanescent intensity is as great or greater than the incident light intensity. The plots can be extended without breaks to the 403

D. Axelrod

Figure 3. Intensity EE" (proportional to the probability of excitation of a fluorophore) versus incidence angle 6 for transmitted light in the low refractive index medium 1 at z = 0 at a TIR interface. The incident intensity in medium 3 is assumed to be unity. At angles 0 > 6C, the transmitted light is evanescent; at angles 0 < 0C, it is propagating. Both s- and p-polarizations are shown. Refractive indices n3 = 1.46 (fused silica) and n1 = 1.33 are assumed here, corresponding to 0C = 65.7°.

subcritical angle range, where the intensity is that of the freely propagating refracted light in medium 1. One might at first expect the subcritical intensity to be slightly less than the incident intensity (accounting for some reflection at the interface) but certainly not more as shown. The discrepancy arises because the intensity in Figure 3 refers to EE* alone rather than to the actual energy flux of the light, which involves a product of EE* with the refractive index of the medium in which the light propagates. 2.1.2 Finite width incident beams For a finite width beam, the evanescent wave can be pictured as the beam's partial emergence from the solid into the liquid, travel for some finite distance along the surface, and then re-entrance into the solid. The distance of propagation along the surface is measurable for a finite width beam and is called the Goos-Hanchen shift. The Goos-Hanchen shift ranges from a fraction of a wavelength at 0 = 90° to infinite at 0 = 6C , which of course corresponds to the refracted beam skimming along the interface. A finite beam can be expressed as an integral of infinite plane waves approaching at a range of incidence angles. In general, the intensity profile of the finite beam evanescent field can be calculated from the mathematical form for the evanescent wave at each infinite plane wave incidence angle, integrated over all the constituent incident plane wave angles. 404

11: Surface fluorescence microscopy with evanescent illumination

Figure 4, Intensity profile of the TIR region illuminated by a Gaussian profile beam from a CW argon laser, as visualized upon a Dil coated surface (see Protocol 1). Configuration A (see Section 3.3) was used here with a Leitz X 10, NA = 0.25 air achromat objective on a Leitz Diavert microscope, (a) Beam slightly defocused at the TIR surface, (b) Beam focused at the TIR surface. The incidence angle is about 2° greater than the critical angle.

For a TIR Gaussian laser beam focused with a typically narrow angle of convergence, the experimentally observed evanescent illumination is approximately an elliptical Gaussian profile, and the polarization and penetration depth are approximately equal to those of a single infinite plane wave. However, if the angle of convergence is greater and the mean angle is within a few degrees of the critical angle, the evanescent field tends to become a long thin stripe (see Figure 4). A more complete mathematical description of the evanescent wave produced by a focused TIR laser beam can be found in ref. 2.

2.2 Intermediate dielectric layers In actual experiments in biophysics, the interface may not be a simple interface between two media, but rather a stratified multilayer system. One example is the case of a biological membrane or lipid bilayer interposed between glass and an aqueous medium. Another example is a thin metal film coating, which can be used to quench fluorescence within the first — 10 nm of the surface. We discuss here the TIR evanescent wave in a three layer system in which incident light travels from medium 3 (refractive index n 3 ) through the intermediate layer (n2) toward medium 1 (n1). 405

D. Axelrod Qualitatively, several features can be noted: (a) Insertion of an intermediate layer never thwarts TIR, regardless of the intermediate layer's refractive index n2. The only question is whether TIR takes place at the n3:n2 interface or the n2:n1 interface. Since the intermediate layer is likely to be very thin (no deeper than several tens of nanometres) in many applications, precisely which interface supports TIR is not important for qualitative studies. (b) Regardless of n2 and the thickness of the intermediate layer, the evanescent wave's profile in medium 1 will be exponentially decaying with a characteristic decay distance given by Equation 3. However, the overall distance of penetration of the field measured from the surface of medium 3 is affected by the intermediate layer. (c) Irregularities in the intermediate layer can cause scattering of incident light which then propagates in all directions in medium 1. Experimentally, scattering appears not be a problem on samples even as inhomogeneous as biological cells. Direct viewing of incident light scattered by a cell surface lying between the glass substrate and an aqueous medium confirms that scattering is many orders of magnitude dimmer than the incident or evanescent intensity, and will thereby excite a correspondingly dim contribution to the fluorescence.

2.3 Intermediate metal film A particularly interesting kind of intermediate layer is a metal film. Theory (1) shows that such a film will reduce the s-polarized evanescent intensity to nearly zero at all incidence angles. But the p-polarized behaviour is quite different. At a certain sharply defined angle of incidence 6P ('the surface plasmon angle'), the p-polarized evanescent intensity becomes an order of magnitude brighter than the incident light at the peak. This strongly peaked effect is due to a resonant excitation of electron oscillations at the metal/water interface. For an aluminium film at a glass/water interface, 6p is greater than the critical angle 0C for TIR. The intensity enhancement is rather remarkable since a 20 nm thick metal film is almost opaque to the eye. There are some potentially useful experimental consequences of TIR excitation through a thin metal film coated on glass: (a) The metal film will almost totally quench fluorescence within the first 10 nm of the surface, and the quenching effect is virtually gone at a distance of 100 nm. Therefore, TIR with a metal film coated glass can be used to selectively excite fluorophores in the 10-200 nm distance range. (b) A light beam incident upon a 20 nm aluminium film from the glass side at a glass/aluminum film/water interface evidently does not have to be collimated to produce TIR. Those rays that are incident at the surface plasmon angle will create a strong evanescent wave; those rays that are 406

11: Surface fluorescence microscopy with evanescent illumination too low or high in incidence angle will create a negligible field in the water. This phenomenon may ease the practical requirement for a collimated incident beam in TIR and make it easier to set up TIR with a conventional arc light source. (c) Thirdly, the metal film leads to a highly polarized evanescent wave (provided I'p = 0), regardless of the purity of the incident polarization.

3. Optical configurations A wide range of optical arrangements for TIRF have been employed. In general, an inverted microscope is more convenient because it provides more room to add TIR optics above rather than below the stage. However, some upright microscope configurations are still very workable. Most configurations use an added prism to direct the light toward the TIR interface, but it is also possible to use the microscope objective itself for this purpose. This section gives examples of these arrangements, partly as a guide to simple systems that work and partly as a basis for creative variations. For concreteness in the descriptions, we assume that the sample consists of fluorescence labelled cells in culture adhered to a glass coverslip. In all cases, the critical angle must be considered in the design. The index of refraction of the standard glass coverslip upon which cells are grown is about n1 = 1.52. The index of refraction of the intact cell interior can be as high as n3 = 1.38. Therefore, to obtain TIR at this interface, the angle of incidence must be larger than the critical angle of 65°. If the cells are not intact (e.g. permeabilized, haemolysed, or fixed) so that the lower refractive index is that of aqueous buffer (n1 = 1.33) instead of cytoplasm, then the critical incidence angle is 61°. Alignment of the optics for TIR is not difficult but it is not immediately obvious either. Immediately after the sections for prism-based TIR and for prismless TIR, protocol boxes are provided to help in the alignment procedure.

3.1 Inverted microscope TIR with prism on top Figure 5 shows several schematic drawings designated A-D for setting up TIR in an inverted microscope with a prism over the sample, all using a laser (usually blue or green output) as a light source. In all these configurations, the buffer-filled sample chamber consists of a lower bare glass coverslip, a spacer ring (often made of 60 um thick Teflon), and the cell coverslip inverted so the cells face down. The upper surface of the cell coverslip is put in optical contact with the prism lowered from above by a layer of immersion oil or glycerol. The lateral prism of the prism is fixed but the sample can be translated while still maintaining optical contact. The lower coverslip can be oversized and the Teflon spacer can be cut with gaps so that solutions can be changed by capillary action with entrance and exit ports. Alternatively, commercial solution 407

D. Axelrod

Figure 5, Four configurations (A-D) for an inverted microscope with a TIR prism above the sample. The sample is a coverslip sandwich with an intervening space; all the vertical distances are exaggerated for clarity here. In configuration B, the dotted lines refer to the option of truncating the prism in order to clear the optical path for a condenser system. In configuration D, the option of producing interference fringes with intersecting beams is depicted with a dashed line.

changing chambers can be used, such as the Dvorak-Stotler (Nicholson Precision Scientific, Gailhersburg, MD), the Sykes-Moore (Bellco Glass Co. Vineland, NJ), or rectangular cross-section microcapillary tubes (Wilmad Glass, Buena. NJ). The configurations in this set share some advantages: (a) (b) (c) (d)

Inexpensive optics. Ample room for set-up, Prism may be mounted on condenser holder for case in raising and lowering, Ease in checking alignment.

However, these configurations share some common drawbacks: (a) Sample not easily accessible from above. (b) Shortest working distance objectives may not reach focus across the buffer layer, (c) Image quality with highest aperture objectives somewhat reduced byviewing through the buffer layer. 408

11: Surface fluorescence microscopy with evanescent illumination The four configurations A-D each have their own advantages and disadvantages. 3.1.1 Configuration A The rectangular solid prism allows transmitted light from the microscope condenser above to reach the sample undistorted, so that phase-contrast and dark-field can be viewed simultaneously. Such a rectangular solid prism must be custom cut and polished, but can be substituted with a commercial polarizing cube. The size of the TIR illuminated region and the incidence angle are easily varied by adjusting the position and angle of the incoming laser beam and the focusing lens. 3.1.2 Configuration B The triangular prism is commercially available inexpensively and in numerous sizes, generally with 45-45-90 right angle and 60-60-60 equilateral shapes. The maximum incidence angle is obtained by introducing the beam from the horizontal direction. For standard glass (n = 1.52), the maximum incidence angle is 73° for the right angle prism and 79° for the equilateral prism. Phasecontrast and dark-field are not compatible with this configuration because the upper surface of the triangular prism is not flat. However, custom truncation and polishing of the top of the prism (shown as an option in the figure) provides compatibility with phase-contrast and dark-field. 3.1.3 Configuration C The 60° trapezoidal prism is the most convenient and reproducible system of all when mounted on the microscope condenser mount. The incoming beam is vertical so the TIR spot shifts laterally very little when the prism is raised and relowered during changes of sample. Phase-contrast and dark-field are compatible with this flat-topped prism. The incidence angle is fixed at 60°. As discussed above, this angle is not sufficient to produce TIR if the prism is made from standard n3 = 1.52 glass. Therefore, the prism must be made of higher than standard refractive index material (e.g. flint glass at n = 1.64). The beam will then refract away from the normal to 0 = 69° in passing from the prism into the coverslip, thereby exceeding the critical angle at the coverslip/water or coverslip/cell interface. Ideally, a trapezoid with walls between 45 and 60 degrees would be best, but that might have to be fabricated as a more expensive special order. Neither 45 or 60 degree trapezoids are commercially available either, but they can be cheaply manufactured by truncating and polishing the apex of a commercially available triangular prism (available from Rolyn Optics, Covina, CA in equilateral flint glass). 3.1.4 Configuration D The parabolic mirror and hemispherical prism here are positioned so that the beam traverses a radius of the prism toward a TIR spot at the focus of the 409

D. Axelrod parabola. In this manner, a lateral shift of the vertical incoming beam will always focus at the same spot. Substantial changes of incident angle thereby can be accessed quite conveniently. In addition, interference fringes in the TIR evanescent field can be created by splitting the incoming beam into two beams, each reflecting at different azimuthal positions in the parabola but recombining at the same parabola focus. The spacing of the fringes can be adjusted by varying the relative azimuthal positions of the two beams. Despite all this versatility, this set-up is rather difficult to align and it can be rather sensitive to vibrations.

3.2 Inverted microscope TIR with prism below TIR can be set up to be compatible with simultaneous microinjection or microelectrophysiology. To provide for easy and continuous access to a liquidcontaining sample from above, a prism TIR set-up should have the prism deployed below stage level. Unfortunately, that is also where the objective resides, so the geometry is rather tight. Configuration E (shown in Figure 6) accomplishes the goal by taking advantage of multiple TIR in the coverslip. This transfers the excitation light as a waveguide from far off-axis to the centre of the field of view. In the simplest form, only a small commercially available triangular prism need be placed in optical contact (via oil) with the bottom of the cell-containing coverslip. The sample can be translated while the prism remains laterally fixed. The problem is that this may leave a smear of oil on the lower side of the coverslip which destroys the first internal reflection. A very feasible alternative is to use an additional intervening coverslip fixed to the prism with optical glue. The sliding motion occurs between the intervening coverslip

Figure 6. Configuration E for an inverted microscope with a TIR prism below the sample. The lower coverslip (shown darker) is fixed by optical glue to the prism. That coverslip is optional, intended to avoid smearing immersion oil.

410

11: Surface fluorescence microscopy with evanescent illumination and the cell coverslip, lubricated and optically contacted by a thin layer of immersion oil. The disadvantage of this set-up is that oil or glycerol immersion objectives cannot be used because that might destroy the TIR of the final internal reflection before the spot under view. However, water (or air) immersion objectives work well with this configuration.

3.3 Upright microscope TIR with prism below Possibly the most convenient TIR set-up of all is configuration F, shown in Figure 7. The same kind of trapezoidal prism as discussed in Section 3.2 can be used here, mounted on the microscope's condenser holder. The laser beam is introduced in the same port in the microscope base as intended for the transmitted light illuminator (which should be removed), thereby utilizing the

Figure 7. Configuration F for an upright microscope with a trapezoidal prism below the sample. It is shown here with a special sealed sample chamber for long-term viewing of cells in a plastic tissue culture dish.

411

D. Axelrod microscope's own in-base optics to direct the beam vertically upward. An extra lens just above the microscope base may be useful to position and focus the TIR spot. As discussed earlier, flexibility in incidence angle is sacrificed for convenience; however, a set of various angled trapezoids will allow one to employ various discrete incidence angles. This system gives particularly high quality images if a water immersion objective is employed and submerged directly into the buffer solution an uncovered cell chamber. This system is also easily used with cells adhering directly on tissue culture plastic dishes rather than on coverslips; the plastic/cell interface is then the site of TIR. If the objective has a long enough working distance, reasonable accessibility to micropipettes is possible. Protocol 1. Alignment of prism TIR Equipment • Laser with blue or green output, at least 1 W total power . Fluorescence microscope equipped with epi-illumination dichroic mirror and barrier filter appropriate for the laser colour, but with the excitation filter removed

• Prism as selected for the desired configuration • Assorted optical mounts • Focusing lens of approx. +50 to 150 mm focal length • Safety goggles

Method 1. Mount the prism on the condenser mount carrier if possible. This need not be done in a precision fashion, but only accurate enough so a usable area at the sample-contacting surface of the prism lies directly in the optical axis of the microscope objective. The mounting may require some custom machining of plexiglass or brass plates and use of a glue (e.g. Duco cement) that can be easily cracked off and reglued if necessary. If a standard condenser will be used for simultaneous phase-contrast or dark-field and the condenser mount cannot hold two separate carriers, then the prism must be mounted on a separate holder with the capability of vertical motion. If the microscope focuses by moving the stage up and down, then this separate holder must be fixed to the microscope stage itself. Otherwise, the prism holder can be fixed directly to the optical table. 2. Depending on the configuration, a system of mirrors with adjustable angle mounts fixed to the table must be used to direct the beam toward the prism. One of these mirrors (or a system of shutters) should be movable and placed near the microscope so switching between standard epi-illumination and TIR is possible without interrupting viewing. Mounts for a focusing lens should also be prepared. 3. Place a uniform fluorescent sample coverslip in the same kind of sample holder to be used for cell experiments. A convenient and 412

11: Surface fluorescence microscopy with evanescent illumination durable uniform film is made from 3,3' dioctadecylindocarbocyanine (also known as 'Dil', available from Molecular Probes, Eugene, OR). If Dil is to be used, dissolve it at about 0.5 mg/ml in ethanol, and place a single droplet of the solution on a glass coverslip. Then, before it dries, rinse off the coverslip with distilled water. A monolayer of fluorophore will remain adhered to the glass; the monolayer is fluorescent and stable either in air or in water. Filter sets appropriate for either fluorescein or rhodamine will work with Dil. 4. With the uniform sample on the stage, focus on the fluorescent surface with transmitted (usually tungsten) illumination. Usually, dust and defects can be seen well enough to assay the focus. However, a deliberately scribed scratch in the fluorescent surface can be made to aid this focusing process. Fluorescent epi-illumination can also be used to find the focus because only at the focal position are laser interference fringes seen sharply. 5. Place a small droplet of immersion oil on the non-Dil surface of the sample coverslip or directly on the prism (depending on which one faces upward in the chosen configuration) and carefully translate the prism vertically so it touches and spreads the oil but does not squeeze it so tightly that lateral sliding motion is inhibited. Too much oil will bead up around the edges of the prism and possibly interfere with the illumination path. 6. By naked eye (perhaps with safety goggles to attenuate errant reflections) and without any focusing lens in place, adjust the unfocused ('raw') collimated laser beam position with the mirrors so that TIR occurs directly in line with the objective's optical axis. This can usually be seen by observing the scattering of the laser light as it traverses through the prism, oil, and TIR surface. 7. Insert the focusing lens so that the focus is roughly at the TIR area under observation. Again by naked eye, adjust its lateral position with translators on the focusing lens mount (not with the mirrors controlling the raw laser beam) so that the TIR region occurs directly in line with the objective. To guide this adjustment, look for three closely aligned spots of scattered light, corresponding to where the focused beam first crosses the immersion oil layer, where it totally reflects off the sample surface, and where it exits by recrossing the oil. 8. The TIR region should now be positioned well enough to appear in view in the microscope when viewed as fluorescence with the standard filters in place. In general, the TIR region will appear as a yellow ellipse or streak. Make final adjustments with the focusing lens to centre this area. The TIR area can be distinguished from two out-of-focus blurs past either end of the ellipse or streak (arising from autofluorescence of the immersion oil) because the TIR spot contains sharply

413

D. Axelrod Protocol 1.

Continued

focused images of defects in the Dil coating. The focusing lens can be moved forward or backward along the laser optical path to achieve the desired size of the TIR area. 9. With the optics now correctly aligned for TIR, translate the prism vertically to remove the Dil sample, and replace it with the actual cell sample. Relower the prism to make optical contact. Although the TIR region will not be exactly in the same spot (because of irreproducibility in the prism height), it will be close enough to make final adjustments with the focusing lens while observing fluorescence from the cell sample.

3.4 Inverted microscope TIR without a prism By using an objective with a numerical aperture of 1.4 (the highest commercially available), supercritical angle incident light can be cast upon the sample by epi-illumination through the objective. The incident beam must be constrained to pass through the periphery of the objective's pupil and must emerge with only a narrow spread of angles; this can be accomplished by assuring that the incident beam is focused off-axis at the objective's back focal plane. It emerges into the immersion oil (n3 = 1.52) at a maximum angle 0 given by: For total internal reflection to take place at the sample surface, 6 must be greater than the critical angle 0C given by: From Equations 6 and 7, it is evident that the NA must be greater than n1, preferably by a substantial margin. This is no problem for an interface with water with n1 - 1.33 and a NA = 1.4 objective. But for viewing the inside of a cell at n1 — 1.38, this configuration is marginal at best for producing TIR. The advantages of prismless inverted TIR include the possibility of viewing a sample that is completely accessible from the top, and the compatibility (in fact necessity) for using the highest aperture, highest resolution, brightest objectives available. The arrangement is also easily compatible with intersecting beams to produce interference fringes: the high mechanical stability enables one to achieve interfringe spacings of ~ 0.3 (um without the blurring effects of small vibrations. Three possible arrangements for prismless TIR are shown in Figure 8: one as described in the Protocol 2; one for producing interference fringes at the TIR surface; and one which utilizes a conventional arc source rather than a laser beam. 414

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Figure 8. Three TIR configurations G, H, and I for an inverted microscope and a NA = 1.4 aperture objective but no prism. The microscope epi-illuminator system is drawn schematically here and connected with configuration H, but the same system is implied for the extensions of configurations G and I. In configuration G, two beams (split from the same laser) intersect at the field diaphragm, forming an interference pattern there and on the sample. In configuration I, an arc lamp rather than a laser is used. An opaque disk must be used here to block subcritical arc lamp light. The conical prism serves only to increase the light flux at supercritical angles so that less energy is lost at the opaque disk.

Protocol 2. Alignment of prismless TIR Equipment • Laser with blue or green output, at least 1 W total power • Inverted fluorescence microscope equipped with epi-illumination dichroic mirror and barrier filter appropriate for the laser colour, but with the excitation filter removed • Objective with numerical aperture of 1.4

• Assorted optical mounts • Plano-convex lens of short radius of curvature or, alternatively, a hemispherical or triangular prism • Converging lens of several centimetres focal length • Safety goggles

Method 1. Place a bare coverslip on the microscope stage (with immersion oil between the objective and the coverslip), and focus on its upper surface. 2. Remove all obstructions between the coverslip sample and the ceiling. Allow a collimated laser beam (the 'raw' beam) to enter the standard

D. Axelrod Protocol 2.

3.

4.

5.

6.

7.

8.

9.

10.

Continued

epi-illumination port field diaphragm along the optical axis. A large area of laser illumination will be seen on the ceiling, roughly straight up. Place the triangular or hemispherical prism or plano-convex lens (flat side down) on the coverslip over the objective, making optical contact with a layer of immersion oil. This prism or lens is not going to be used in actual experiments. It is used here to avert total internal reflection and thereby couple light out of the coverslip and onto the wall or ceiling of the room. Here again, safety goggles are advisable. Reposition the laser beam with mirrors positioned upbeam from the microscope so that the beam still enters the centre of the field diaphragm but now at a small angle to the optical axis. This angle should be continuously adjustable; slowly increase the angle. The ceiling laser illumination will 'set' to an ever-lower position on the wall until it just disappears. At this angle, it is just blocked by the internal aperture of the objective. Back off the entrance angle so that half the illuminated area is seen. Place the converging lens about 20 cm 'upbeam' from the field diaphragm and concentric with the incoming beam. The illuminated region on the wall will now be a different size, probably smaller. Move the converging lens longitudinally (along the axis of the laser beam) to minimize the illuminated region on the wall. This will occur where the converging lens focal point falls exactly at a plane outside the microscope equivalent to the objective's back focal plane. At this position, the beam is thereby also focused at the objective's actual back focal plane and emerges from the objective in a roughly collimated form. Fine-tune the lateral position of the converging lens and the raw beam mirrors so that the beam on the wall is just barely above the point at which it disappears. This ensures that the beam is propagating up along the inside periphery of the objective. Verify that TIR is achievable by moving to the sample to a clean new spot not under the prism. No light should emerge (except for some scattering) because of total internal reflection at the glass coverslip/ air interface. Replace the bare coverslip with an identical one coated with Dil (see Protocol 7). Place a droplet of water on the Dil coverslip directly above the objective, and again verify that no light emerges even at this glass/water interface. View the Dil fluorescence through the microscope; the illuminated region should be circular or elliptical and roughly centred. Fine-tune 416

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11.

12.

13.

14.

the mirrors and converging lens to centre the TIR fluorescence in the field of view. The size of the TIR fluorescence area on the sample is directly proportional to the laser beam size at the field diaphragm. To change this size, replace the converging lens with another one of different focal length, but always keep its focal point at the objective's equivalent back focal plane, which is at a fixed position upbeam from the microscope. Replace the Dil coverslip with the actual cell sample. When the cells are in-focus, the TIR optics should be perfectly aligned without need of further adjustment. If switching back and forth between TIR and epi-illumination is desirable, design the optics with movable mirrors (within easy reach) that can select angular or straight-on laser paths. If interference fringe TIR is desirable, use a beam splitter and arrange the second beam to enter the centre of the field diaphragm from an approach at the same angle but at different azimuthal position around the optical axis. A relative azimuthal angle of 180° will give the closest spaced fringes. It may be most convenient to use the same converging lens for both beams. Then the converging lens should be positioned on-axis but each beam should enter it off-axis so that each still arrives at the centre of the field diaphragm. Be sure that any difference in path length of the two beams from the beam splitter to their reintersection point is less than the coherence length of the laser (a few millimetres or centimetres); otherwise, no interference will occur.

3.5 Rapid chopping between TIR and epi-illumination Regardless of the method chosen to produce TIR in a microscope, it is often desirable to switch rapidly between illumination of the surface (by TIR) and deeper illumination of the bulk (by standard epifluorescence). For example, a transient process may involve simultaneous but somewhat different fluorescence changes in both the submembrane and the cytoplasm, and both must be followed on the same cell in response to some stimulus (3). Figure 9 shows a method using computer driven acousto-optic modulators by which the rapid chopping can be done.

4. General experimental suggestions Regardless of the optical configuration chosen, the following suggestions may be helpful. (a) The prism used to couple the light into the system and the (usually disposable) slide or coverslip in which TIR takes place need not be matched exactly in refractive index. 417

D. Axelrod

Figure 9. Optoelectronic system used for rapid chopping between TIR and EPI. The zero order diffraction direction of acousto-optic modulator 1 (AOM1) is the TIR path; the first order is the EPI path. AOM 2 and AOM3 serve only to increase the on/off contrast of the EPI and TIR beams, respectively, so that they cleanly alternate. The computer that controls the square wave pulses to the AOMs also acquires data from the single-channel photon counter so that the time course of the two illumination modes can be separated in software.

(b) The prism and slide may be optically coupled with glycerol, cyclohexanol, or microscope immersion oil, among other liquids. Immersion oil has a higher refractive index (thereby avoiding possible TIR at the prism/ coupling liquid interface at low incidence angles) but it tends to be more autofluorescent (even the 'extremely low' fluorescence types). (c) The prism and slide can both be made of ordinary optical glass for many applications, unless shorter penetration depths arising from higher refractive indices are desired. Optical glass does not transmit light below about 310 nm and also has a dim autoluminescence with a long (several hundred microsecond) decay time, which can be a problem in some photobleaching (FRAP) experiments (see Section 5d). The autoluminescence of high quality fused silica (often called 'quartz') is much lower. Tissue culture dish plastic (particularly convenient as a substrate in the upright microscope set-up) is also suitable, but tends to have a significant autofluorescence compared to ordinary glass. Different brands of tissue culture plastic have significantly different amounts of autofluorescence; Corning brand is one of the least fluorescent. More exotic high n3 materials such as 418

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(d) (e)

(f)

(g)

(h)

(i)

sapphire, titanium dioxide, and strontium titanate can yield exponential decay depths d as low as l0/20. The TIR surface need not be specially polished: the smoothness of a standard commercial microscope slide is adequate. Illumination of surface adsorbed proteins can lead to apparent photochemically-induced cross-linking. This effect is observed as a slow, continual, illumination-dependent increase in the observed fluorescence. It can be inhibited by deoxygenation (aided by the use of an O2-consuming enzyme/ substrate system such as protocatachuic deoxygenase/protocatachuic acid or a glucose/glucose oxidase system), or by 0.05 M cysteamine. Virtually any laser with a total visible output in the 0.5 W or greater range should be adequate. The most popular laser for cell biological work with a microscope appears to be a 3 W continuous wave argon laser. TIRF experiments often involve specially coated substrates. A glass surface can be chemically derivatized to yield special physi- or chemi-absorptive properties. Covalent attachment of certain specific chemicals are particularly useful in cell biology and biophysics, including: poly-L-lysine for enhanced adherence of cells; hydrocarbon chains for hydrophobicizing the surface in preparation for lipid monolayer adsorption; and antibodies, antigens, or lectins for producing specific reactivities. Derivatization generally involves pre-treatment of the glass by an organosilane. A planar phospholipid coating (possibly with incorporated proteins) on glass can be used as a model of a biological membrane. Methods for preparing such model membranes on planar surfaces suitable for TIR are reviewed in ref. 4. Aluminium coating (for surface fluorescence quenching; see Section 2.3) can be accomplished in a standard vacuum evaporator; the amount of deposition can be made reproducible by completely evaporating a premeasured constant amount of aluminium. After deposition, the upper surface of the aluminium film spontaneously oxidizes in air very rapidly. This aluminium oxide layer appears to have some similar chemical properties to the silicon dioxide of a glass surface; it can be derivatized by organosilanes in much the same manner. The TIRF spot should be focused to a width no larger than the field of view; the larger the spot, the more that spurious scattering and out-offocus fluorescence from the immersion oil layer between the prism and coverslip will increase the generally very low fluorescence background attainable by TIRF. A laser source is generally preferable to an arc lamp for TIRF, because collimation of arc lamp light entails a great loss of intensity. On the other hand, laser illumination suffers from unavoidable interference fringing on the sample. This can be minimized by very clean optics. But for critical 419

D. Axelrod applications, it may be advisable to perform rapid jiggling of the beam or to compute a normalization of sample digital images with the control digital image of a uniform concentration of fluorophores. (j) The incidence angle should exceed the critical angle by at least a couple of degrees. At incidence angles very near the critical angle, the cells cast a noticeable 'shadow' along the surface.

5, Applications of TIRF microscopy Applications of TIRF microscopy in cell biology include: (a) Localization of cell/substrate contact regions in cell culture. Quantitative determination of the absolute distance from the surface to a labelled cell membrane at a cell/substrate contact region can be based on the variation of the emitted fluorescence with 0. This effort is challenging because corrections have to be made for 6-dependent reflection and transmission through four stratified layers (glass, culture medium, membrane, and cytoplasm), all with different refractive indices. A variation of TIRF to identify cell/substrate contacts produces essentially a negative of the standard fluorescence view of labelled cells (5). The solution surrounding the cells is doped with a non-adsorbing and non-permeable fluorescent volume marker, fluorescein labelled dextran. Focal contacts then appear as dark areas and other areas appear brighter, depending on the depth of solution illuminated by the evanescent wave in the cell/substrate gap. (b) High contrast visualization of submembrane cytoskeletal structure on thick cells. Optical sectioning by TIRF is particularly useful in viewing submembrane cytoplasmic filaments on thick cells. Epi-illumination excites fluorescence from out-of-focus planes and leads to a diffuse fluorescence that obscures detail. Although TIRF cannot view deeply into the cell, it can display the submembrane filament structure with high contrast. (c) Time lapse fluorescence movies. Epifluorescence illumination appears to adversely affect cell viability, even with occasional exposure for making movies. TIRF seems particularly advantageous for long-term viewing of cells, since the evanescent wave minimizes exposure of the cells' organdies to excitation light. (d) Desorption kinetic rates of reversibly bound biomolecules at biological membrane surfaces. If the evanescent wave intensity is briefly flashed brightly, then some of the fluorophores associated with the surface will be photobleached. Subsequent exchange with unbleached dissolved fluorophores in equilibrium with the surface will lead to a recovery of fluorescence, excited by a continuous but much attenuated evanescent wave. The time course of this recovery is a measure of the desorption kinetic rate (6). (This technique is called TIR/FRAP in reference to fluorescence 420

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(e)

(f)

(g) (h)

(i)

(j)

recovery after photobleaching.) Kinetic rates of hormones and cytoskeletal proteins at model membranes, flattened biological membranes, and intact cell surfaces have been measured by this technique. Kinetics at the cytofacial surface of intact cells can be viewed employing TIR/FRAP on cells that have been microinjected with a labelled molecule of interest. Surface diffusion of reversibly bound biomolecules at biological membrane surfaces. TIR/FRAP can be used to measure surface diffusion coefficients along with on/off kinetics, if the evanescent wave intensity is variegated over a distance on the surface short compared to the characteristic distance covered by surface diffusion within the time available before desorption. One way of producing this variegation is simply by focusing; another way is by interfering two TIRF beams, as discussed with configuration D (Section 3.1.4); a third way is by placing a striped pattern Ronchi ruling in the incident beam at a position where it forms a real image on the sample. Orientational distributions of fluorescent molecules at a surface. The polarization properties of the evanescent wave can be used to excite selected orientations of fluorophores. Standard polarized epi-illumination cannot distinguish order from disorder in the polar direction (defined as the optical axis) because epi-illumination is polarized transverse to the optical axis. But microscope TIR illumination can be partially polarized in the optical axis direction (the z direction) and can thereby detect order in the polar direction. Note that fluorescence polarization detected through a microscope should be corrected for the slight depolarizing effect of high aperture objectives (7). Reduction of cell autofluorescence relative to fluorescence excited at cell/substrate contacts. Viewing a cell bound fluorescent ligand in the presence of the same ligand in the bulk. Many cell surface receptors bind their ligands reversibly, so a surface bound population can be maintained only with a substantial concentration in the bulk. In some other cases, the cell surface receptor binds ligands irreversibly but then the complex is internalized and replaced by a fresh unbound receptor. The newly appearing receptors can be labelled continuously with ligand in the bulk. The bulk fluorescence will be discriminated against by TIRF. Observing rapid dynamical processes at biological membranes. TIRF has been used to quantitatively compare intracellular ionic concentration transients proximal to the membrane versus deeper in the interior (3). It has also been used qualitatively to observe fluorescence loaded secretory vesicle fusion with the membrane (8). Reduction of instrumental luminescence background. In standard epiillumination, the full power of the illumination light excites some fluorescence from the glass and glues in the objective. In TIR, this background is 421

D. Axelrod eliminated, thereby making fluorescence detection of single molecules more feasible (9, 10). Related to this application is the capability of seeing fluorescence fluctuations as fluorescent molecules enter and leave the thin evanescent field region in the bulk. These visually obvious fluctuations can be quantitatively autocorrelated to obtain kinetic information about the molecular motion ('fluorescence correlation spectroscopy', FCS). (j) Fluorescence lifetimes at surfaces. TIR has been combined with fluorescence lifetime imaging of cultured cells labelled with a terbium chelate (11). Such chelates have a very long (millisecond) lifetime compared with organic fluorophores, but as techniques for measuring fast fluorescence lifetimes in a microscopic spatially resolved region are further developed, they will be readily adapted to TIRF.

6. Comparison with other optical sectioning microscopies Confocal microscopy (CM) is another technique for apparent optical sectioning, achieved by exclusion of out-of-focus emitted light with a set of image plane pin-holes. CM has the clear advantage in versatility; its method of optical sectioning works at any plane of the sample, not just at an interface between dissimilar refractive indices. However, other differences exist which, in some special applications, can favour the use of TIRF: (a) The depth of the optical section in TIRF is ~ 0.1 um whereas in CM it is a relatively thick ~ 0.6 um. (b) In some applications (e.g. FRAP, FCS, or on cells whose viability is damaged by light), illumination and not just detected emission is best restricted to a thin section; this is possible only with TIRF. (c) Since TIRF can be adapted to and made interchangeable with existing standard microscope optics, even with 'home-made' components, it is much less expensive than CM. However, at the time of this writing, TIRF microscopy modules or kits are not commercially available; the mirrors and prisms must be purchased separately from optical supply companies and configured by the end user. Cell/substrate contacts can be located by a non-fluorescence technique completely distinct from TIRF, known as 'interference reflection microscopy' or 'reflection contrast microscopy' (RCM). Using conventional illumination sources, RCM visualizes cell/substrate contacts as dark regions. RCM has the advantage that it doesn't require the cells to be labelled, but the disadvantages that it contains no information of biochemical specificities in the contact regions and that it is less sensitive to changes in contact distance (relative to TIRF) within the critical first 100 nm of the surface. 422

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Acknowledgements The author is grateful to Susan Sund and Rhonda Dzakpasu for useful discussions. Figure 1 was provided by Susan Sund. This work was supported by a grant NSF MCB9405928.

References 1. Axelrod, D., Hellen, E.H., and Fulbright, R.M. (1992). In Topics in fluorescence spectroscopy, Vol. 3: biochemical applications (ed. J.R. Lakowicz), p. 289. Plenum Press, New York. 2. Burghardt, T.P. and Thompson, N.L. (1984). Opt. Eng., 23, 62. 3. Omann, G.M. and Axelrod, D. (1996). Biophys. J., 71, 2885. 4. Thompson, N.L., Pearce, K.H., and Hsieh, H.V. (1993). Eur. Biophys. J., 22, 367. 5. Gingell, D., Heavens, O.S., and Mellor, J.S. (1987). J. Cell Sci., 87, 677. 6. Thompson, N.L., Burghardt, T.P., and Axelrod, D. (1981). Biophys. J., 33, 435. 7. Axelrod, D. (1989). In Fluorescence microscopy of living cells in culture. Part B (ed. D.L. Taylor and Y.-L. Wang), Meth. Cell Biol. 30, p. 333. Academic Press, New York. 8. Steyer, J.A., Horstmann, H., and Aimers, W. (1996). Nature, 388, 474. 9. Vale, R.D., Funatsu, T., Pierce, D.W., Romberg, L., Harada, Y., and Yanagida, T. (1996). Nature, 380, 451. 10. Dickson, R.M., Norris, D.J., Tzeng, Y.-L., and Moerner, W.E. (1996). Science, 274, 966. 11. Phimphivong, S., Kolchens, S., Edmiston, P.L., and Saavedra, S.S. (1995). Anal. Chim. Acta, 307, 403.

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12 Nanovid microscopy GRETA M. LEE

1. Introduction Nanovid microscopy, nanometre-sized particles viewed with video-enhanced microscopy (1-3) is used to directly observe the lateral mobility of gold labelled membrane proteins and lipids, to observe the precise location and mobility of gold labelled pericellular and extracellular matrix macromolecules, and to follow endocytosis of gold labelled molecules. The technique involves the use of 30-40 nm colloidal gold particles conjugated to antibodies, streptavidin, lectins, or other ligands. The association of the antibody to the gold is non-covalent but extremely stable. Since a single colloidal gold particle is typically conjugated to many antibody molecules (4), an individual gold particle may bind to a small group of like membrane molecules rather than to a single molecule (5). The gold tag is visualized using video contrast enhancement to detect the light scattering of the gold particle. The gold can be viewed with bright-field, DIC, or epipolarization optics. Examples of bright-field and DIC images are shown in Figure 1. Individual 20-40 nm gold particles are not directly visible by eye through the oculars but can be detected with video contrast enhancement of the Airy disk (see Chapter 3) produced by light scattering of the gold particle. Video contrast enhancement involves both camera (analogue) and image processor (digital) adjustments as described below. The degree of scattering is proportional to the sixth power of the particle radius and to the fourth power of the illuminating wave number (the reciprocal of the wavelength) (2). Because the light scatter is so strong, gold particles appear much larger in the video image than in electron micrographs that show the true size relative to the cell. This is demonstrated in Figure 2 where the same cell is viewed by video light microscopy and electron microscopy (prepared as described in ref. 6). Note how much smaller the gold particles are relative to the cell in the electron micrograph. Also note that similar sized gold particles can have varying levels of contrast in the video image. This can be due to slight variations in focal level. In addition, single gold particles cannot be distinguished from doublets in the video image, but three adjacent gold particles have higher contrast and appear as a larger spot. Whether two or three gold particles are imaged as a single

Greta M. Lee

Figure 1. Examples of colloidal gold labelling of the plasma membrane (A, B) and the pericellular matrix (C, D). (A) and IS) show a lamella of a C3H-10T1/2 fibroblast labelled with 30 nm colloidal gold conjugated to an antibody to Thy-1, a glycophosphatidyl inositol-linked protein found on the cell surface. Note how the gold particles are easily recognized on the cell surface in the bright-field image (A) but are difficult to distinguish from intraceflular structures in the DIC image (B). The gold particles do not align in the two images because they were mobile. (C) and (D) show a bovine chondrocyte labelled with 40 nm colloidal gold conjugated to an antibody to type VI collagen. The label is found in the pericellular matrix which is a three-dimensional structure extending out from this rounded cell. Individual gold particles are readily detectable in the bright-field image (C) but are difficult to to see in the DIC image (B). Arrowheads indicate cell edges. All figures are at the same magnification. Bar = 2 um.

spot or as separate spots is dependent on their spacing and on the resolution limit of the optics. For two adjacent points to be resolved as separate points, the centres of the Airy disks for those two points must be separated by a minimum distance determined by the resolving power of the optics (see Chapter 1), Thus an objective and a condenser with high numerical aperture will be hetler for resolving closely spaced gold particles. The variation in image size for single gold particles and the inability to distinguish between single and two closely spaced gold particles make double labelling with colloidal gold of two different sizes impractical with videoenhanced light microscopy. However, colloidal gold can be used with a fluorescent probe or 1-2 um latex beads. 426

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Figure 2. Bright-field video image of a region of a gold labelled cell before (A) and after image processing for automatic detection of gold particles (B), The electron micrograph (C) of the boxed region in (B) demonstrates that individual 30 nm gold particles can be detected with a video microscope system. The lower case letters indicate corresponding single gold particles (a-d) and aggregates (e and f). This C3H fibroblast was labelled with 30 nm gold anti-pgp-1, which binds to an integral membrane glycoprotein. After fixation and imaging by nanovid microscopy, the cell was processed for whole mount transmission electron microscopy. The distances between the gold particles are shorter in the electron micrograph than in the video images due to cell shrinkage during the dehydration steps. Bars (B) = 2 um; (C) = 0.2 um. Reproduced from ref. 9 with permission.

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1.1 Use of nanovid microscopy to analyse molecular mobility in membranes To analyse the pattern of movement of the labelled molecule, a series of images over time is recorded. The time frame for recording depends on the rate of movement of the labelled molecules and on the question being asked. For fast diffusing molecules, such as lipids in a membrane, images are recorded at video rate. For slower moving molecules, such as capping of membrane proteins, images may be recorded at one frame per second or slower. The sophistication of the analysis can range from tracing the movement on a video monitor with a felt tip marker to analysing the centroids (x, y coordinates of the centre) of each gold particle in each image. For the latter, the analysis can involve: (a) Manually locating the centroid with a pointer. (b) Manually identifying the particles to be tracked and using an image processing program to compute the centroid. (c) Using the image processor to both identify the particle and compute the centroid. The centroids are then used to compute the trajectory of the labelled molecule. These trajectories are used to analyse the pattern of movement and/or to compute the diffusion coefficient. One of the purposes of this approach is to directly observe intramembrane domains or 'corrals' as well as free diffusion and directed transport within the plane of the membrane. Unfortunately, Brownian motion produces trajectories with a variety of random patterns (7, 8) that can complicate the detection of larger domains (9). Thus the interpretation of the data requires careful statistical analysis (5, 10, 11). With more sophisticated analysis, temporary confinement can be separated from pure random motion (12, 13).

1.2 Use of nanovid microscopy to study the extracellular matrix Another, less widely recognized, use of nanovid microscopy is to study location, mobility, and assembly of extracellular matrix molecules on living and fixed cells. This approach is especially useful for studying the pericellular matrix on living cells because this matrix is made up of hydrated hyaluronan and proteoglycans which collapse with fixation and dehydration. Nanovid microscopy was used to analyse the roles of hyaluronan and aggrecan in forming the three-dimensional, dynamic pericellular matrix on chondrocytes (14). Colloidal gold particles can also be used as markers on extracellular matrix fibrils, such as collagen fibrils, to follow their movements on the cell surface as the cell arranges its extracellular matrix (Lee and Loeser, submitted). 428

12: Nanovid microscopy Protocol 1. Probe preparation: method for conjugating purified antibodies to 30 or 40 nm colloidal golda Equipment and reagents 96-multiwell plate pH meter Microcentrifuge Purified monoclonal or polyclonal antibody, dialysis tubing, and clips 30 or 40 nm colloidal gold 20M Carbowax

• 10% bovine serum albumin (BSA) dissolved in dH2O • Glycerol • 2 mM borate buffer pH 8.9 . 0.2 M K2C03, 0.2 M H3PO4 . 20 mM Tris, 150 mM NaCI pH 8.2

Method 1. Dialyse the antibody (roughly 300 ul of 0.1 mg/ml) against 2 mM borate buffer pH 8.9, for 1 h at 4°C. Centrifuge the protein to remove any aggregates. 2. Adjust 10 ml colloidal gold suspension to pH 8.9 with 0.2 M K2CO3 and 0.2 M H3P04. Caution: gold plugs non-gel type pH electrodes. Work quickly and soak the electrode in pH 4 citrate buffer immediately after use. Alternatively, use pH paper. 3. Do a protein concentration curve: put a 96-multiwell plate on a white surface. Using five to ten wells, do a dilution series of the protein adding 10 ul to each well. Add 100 ul colloidal gold to each. Mix well. Wait 10 min. Add 10 ul 10% NaCI, mix, wait 5 min. Look at the colour— pink indicates enough protein to stabilize 100 ul of gold, blue indicates aggregation of the gold due to an inadequate amount of protein. Select the lowest concentration of protein which gives a pink colour as the concentration to use. 4. Calculate the volume of protein necessary to stabilize the remaining gold solution or the volume of gold that can be stabilized with the amount of protein available. Add the dialysed protein to the gold suspension, vortex for 2 min, let sit at room temperature for 8 min. 5. Stabilize the gold by adding 10% BSA to 1%, and 1% 20M Carbowax to 0.05%. Centrifuge at 15000 g for 12 min. Discard the supernatant and resuspend the pellet in 0.05% Carbowax, 1% BSA, 20 mM Tris, 150 mM NaCI pH 8.2. Give two more washes in the same solution to remove any unbound antibody. 6. Resuspend the pellet in a small quantity (100-500 ul) of the above solution (the gold suspension should be the colour of port wine). Add glycerol to 50%. Store at-20°C. • Adapted from Aurobeads colloidal gold for macromolecule labelling, by Janssen.

429

Greta M. Lee General tips: (a) Filter all solutions with a 0.22 um filter. Particulates can cause the gold to aggregate and bacteria will be concentrated with the gold by the wash steps. (b) As an alternative to using incredibly clean glassware, use sterile, disposable plastic test-tubes. (c) To reduce the sticking of conjugated gold to the sides of Eppendorf tubes during centrifugation, add the wash solution first. (d) Osmolarity is important for antibody stability. Before and after conjugation to gold, minimize the time antibodies are left in very dilute salt solutions. An antibody that has lost its proper conformation by prolonged exposure to dilute solution will have little to no antigen affinity after conjugation to the gold. (e) To prepare paucivalent gold probes (5), mix an irrelevant antibody with the desired antibody before adding to the gold. Do a protein concentration curve on each to determine the correct ratio for mixing. Protocol 2. Cell culture and labelling Equipment and reagents • Coverslips and glass scribe to cut spacers • Silicon grease or Parafilm • 30-40 nm colloidal gold conjugated to a primary antibody, streptavidin,a or a lectin

« Tissue culture mediumb • 10% BSA • VALAP: vasoline:lanolin:paraffin (1:1:2, by weight)

Method 1. Grow cells in a monolayer on No. 1 22 mm2 coverslips. (Ideally, a No. 1.5 coverslip should be used, but in this thickness category there are a few coverslips that are too thick for focusing with a high NA oil immersion objective. Use of No. 1 coverslips avoids the frustration.) 2. Remove the glycerol and any unbound antibody by adding 5-20 ul of gold suspension (as prepared in Protocol 7) for each coverslip to 500 ul tissue culture medium containing 1% BSA. Centrifuge as in Protocol 1. Repeat the wash step once or twice. 3. Resuspend the pellet in the desired volume, approx. 100 ul per coverslip. 4. Place two No. 1 or No. 2 coverslip strips as spacers on a clean glass slide; a thin film of silicon vacuum grease can be used to hold them in place. Alternatively, two strips of Parafilm can be used as spacers. 5. Place 100 ul of gold suspension on the slide between the spacers, and invert the coverslip with cells onto the drop. 430

12: Nanovid microscopy 6. Incubate for 20 min or longer in a moist chamber at room temperature or 37°C depending on your experiment. 7. Carefully dry the sides with the spacers then seal with melted VALAP. 8. Slowly flow 200 ul of tissue culture medium with BSA or serum through the chamber to rinse out unbound gold. Flowing medium through too quickly can shear the cells. 9. Dry the remaining edges of the coverslip and seal with VALAP. A vacuum flask attached to the house vacuum line and with a Pasteur pipette inserted in the intake tubing works well for drying. 10. Clean the top of the coverslip with a small drop of dH20. 11. Prepare a control coverslip without colloidal gold for comparison purposes. * When using streptavidin gold, the cells need to first be labelled with a biotinylated probe. b Hepes-buffered Ham's F-12 (Sigma, N8641) is a good medium as the the bicarbonate concentration is low enough that fair pH control can be maintained outside of a CO2 atmosphere.

Protocol 3. Viewing the labelled cells Equipment • Microscope with DIC optics, high NA, oil immersion condenser and objective, a green band pass filter, and IR (e.g. Schott KG5) and UV (e.g. Schott GG475) filters, 2-4 x adapter for the video camera, and stabilized light source • Video camera with manual gain and offset adjustments • Video monitor

Computer with frame grabber Image processor software capable of handling real time background subtraction Image storage device (e.g. VCR, large hard disk, Zip drive, optical disk recorder) Air curtain incubator or heated stage and objective for live mammalian cells

Method 1. Place the specimen on the stage. For an inverted microscope, place a drop of oil on the slide and bring the condenser lens into contact with the oil. For an upright microscope, place a drop of oil on the condenser and bring the oil into contact with the slide. 2. With x 20 objective and phase-contrast or DIC optics, focus on the specimen. (See Chapter 1, Protocol 9 for instructions in using DIC optics.) 3. Adjust the condenser for Kohler illumination (Chapter 1, Protocol 2). 4. Locate a cell of desired morphology. 5. Switch to the x 100 objective using a small drop of oil on the objective and focus on the specimen with DIC optics.

431

Greta M. Lee Protocol 3.

Continued

6. Readjust the condenser for Kohler illumination. 7. Obtain a good image of the desired cell on the video monitor. Save the image to disk for future reference. 8. Reduce the light intensity and then go to bright-field optics by removing the analyser from the light path. 9. If the camera has an indicator for too much light, or if you have an oscilloscope, or your software has a live histogram function, increase the light intensity to the maximum allowable for the camera or as high as possible without saturating the pixels. 10. Use analogue contrast enhancement to view the gold on the video monitor. Adjust the camera by increasing the gain to give a bright image then bringing up the black level until the screen appears a normal grey. (Do not overly increase the gain as the resulting increase in noise will make analysis more difficult.) If the gold particles are not visible, the adjustments are repeated until any dirt specks on the camera face plate are displayed on the monitor. 11. Delicately adjust the fine focus until gold particles are visible as small black spots.a Small organelles within the cell may also show up. With bright-field, the organelles can be distinguished from the gold by slight focal adjustments which make the gold particle disappear but change the organelles from black to white (1). 12. Use digital contrast enhancement to further improve the image. A background image (a blank area only slightly out-of-focus) is subtracted, and then the contrast is enhanced by adjusting the gain and offset of the image with the image processor. • If the selected cell is not labelled, there are usually gold particles that are non-specifically attached to the substratum that can be imaged. These also serve as a reference for distinguishing gold particles from organelles. Examine the control slide without gold labelling to improve confidence in distinguishing gold from organelles.

2. General information on equipment and methods 2.1 Microscope Both bright-field and DIC microscopy can be used. With bright-field, more of the gold particles show up and the gold is easier to distinguish from organelles; however, the optical sectioning is not quite as good as with DIC. When using bright-field to view the gold, it is useful to record corresponding DIC images to provide information regarding location on the cell, cell shape, etc. For bright-field, having the polarizer and Wollaston prisms in the light path does not seem to affect the image quality. 432

12: Nanovid microscopy All lenses and filters in the light path need to be clean because the contrast enhancement necessary to see the gold particles will also show up any dirt. The use of a 2-4 X adapter or projection lens for the camera is dependent on the size of the imaging area of the camera and whether individual gold particles are to be tracked or just visualized. With a 1" newvicon camera, the adapter is needed for tracking but is not necessary for localization. A 1/3" CCD chip gives a higher magnification in the video image and thus the adapter may not be needed. The light source needs to be extremely stable if image analysis for tracking of bright-field images is to be semi-automated. This is because slight variations in grey level due to light fluctuations are amplified by the processing needed for the image processor to detect the gold particles. Thus the light fluctuations result in images that are too dark or light for semi-automated analysis.

2.2 Camera A video camera is used to both visualize the gold and to give analogue contrast enhancement. A newvicon tube camera with manual adjustments of gain and black level works well, but there is variation in how far the gain and black level can be adjusted among different manufacturers. Thus, it is wise to test the camera before purchase. A method to indicate adequate and excess light to the camera is extremely helpful for bright-field adjustments. For image acquisition of rapidly moving particles, we have found the Hamamatsu C2400 performs well. A CCD camera with both gain and offset or black level adjustments can also be used. Some CCD cameras have a mottle on the face plate that is visible at high contrast. Output rate of some of the high resolution digital CCD cameras may be a limiting factor for nanovid microscopy because precise focusing is difficult at less than 15 frames per second. There is adequate light especially with bright-field microscopy, and thus cooled, integrating cameras designed for low light level work are not needed. On the other hand, sensitivity to electronic noise is important in this application because the contrast is pushed so high that all noise is also amplified making subsequent image analysis difficult. The presence of electronic noise also appears to be a function of the electrical grounding within the microscope as we have found that cameras attached to an Olympus 1X70 microscope pick up very little electronic noise.

2.3 Image processor and frame grabber For acquiring images of particles diffusing at 1-10 X 10~9 cm2/sec, real time background subtraction and digital contrast enhancement are needed. Real time image averaging capabilities are helpful for slower moving gold particles. This can be typically performed faster on the frame grabber board than in the computer. We have found that Image-1 from Universal Imaging (West Chester, PA) simultaneously performs real time background subtraction, 433

Greta M. Lee contrast enhancement, and image storage to an optical disk recorder. MetaMorph, also from Universal Imaging, is supplied with different video boards and thus real time background subtraction and enhancement may not be available. NIH-Image, freeware available for both the Macintosh and the PC can also be used for image acquisition and analysis. The Scion AG-5 board (Macintosh) is compatible with NIH-Image and newvicon and video CCD cameras and allows video rate background subtraction and frame averaging, but there is only digital output of the processed image which means the images must be stored on a computer prior to making a video tape.

2.4 Image storage Depending on the recording rate, images can be stored on a hard disk drive, an optical memory disk recorder (Panasonic), or video tape. An optical disk recorder gives video rate storage and retrieval, the resolution is good, and images can be retrieved individually and selected by the image processor; however, the initial cost is high. For movies, when later analysis is not desired, half-inch VHS is acceptable, but S-VHS is better if the recording will be copied later. With NIH-Image and the Scion AG-5 board, cropped images can be stored to the hard disk at or near video rate. Video rate storage is only essential for fast diffusing particles (i.e. gold labelled lipids in a membrane). For slower moving particles, a running average of two frames will give a sharper image, and images can be saved at ten or less frames per second. A minimum of 200 images in a series is recommended for analysing trajectories of individual gold particles. An Iomega Jaz drive with 1 GB disk cartridges is a simple way to store large numbers of digitized images. Alternatively, a writable CD can be used as an economical storage medium.

3. Image analysis to analyse molecular mobility (single particle tracking) To track the movement of individual gold particles, the centroids of each gold particle of interest needs to be located in a series of images. One can manually point to each particle with a cursor to find the centroids, but this is a tedious and difficult process. The alternative is to use the image processor to automatically locate the particles with user-defined selection criteria. For brightfield images, the gold particles need to be separated from the background by thresholding. A histogram function for adjusting grey levels can also be helpful. Smoothing or low pass filters make the gold particles appear more uniform. For the actual determination of the centroids with Image-1's 'object measurement mode' or NIH-Image's 'particle analysis', parameters must first be set to define size and shape limits so that the gold particles can be distinguished from noise and to eliminate aggregates. DIC images do not have enough contrast for use with the object measurement mode or particle analysis 434

12: Nanovid microscopy routines, but MetaMorph will perform semi-automated single particle tracking with subpixel resolution (15) for bright-field, DIC, and fluorescence images. The centroid data can be output to an Excel spread sheet or to customized programs (9, 13) for assembly of trajectories and calculation of diffusion coefficients.

References 1. DeBrabander, M., Nuydens, R., Geuens, G., Moeremans, M., and DeMey, J. (1985). Cytobios, 43, 273. 2. DeBrabander, M., Geerts, H., Nuyens, R., Nuydens, R., and Corneliessen, F. (1993). In Electronic light microscopy techniques in modern biomedical microscopy (ed. D. Shotton), p. 141. Wiley-Liss, Inc., New York. 3. Geerts, H., DeBrabander, M., Nuydens, R., Geuens, S., Moeremans, M., DeMey, J., et al. (1987). Biophys. J., 52, 775. 4. DeRoe, C., Courtoy, P. J., and Baudhuin, P. (1987). J. Histochem. Cytochem., 35, 1191. 5. Lee, G. M., Ishihara, A., and Jacobson, K. A. (1991). Proc. Natl. Acad. Sci. USA, 88, 6274. 6. Van den Pol, A. N., Ellisman, M., and Deerinck, T. (1989). In Colloidal gold: principles, methods and applications, Vol. 1 (ed. M. A. Hayat), p. 452. Academic Press, San Diego. 7. Berg, H. C. (1983). Random walks in biology, p. 5. Princeton University Press, Princeton. 8. Rudnick, J. and Gaspari, G. (1987). Science, 237, 384. 9. Lee, G. M., Zhang, F., Ishihara, A., McNeil, C. L., and Jacobson, K. A. (1993). /. Cell BioL, 120, 25. 10. Qian, H., Sheetz, M. P., and Elson, E. L. (1991). Biophys. J., 60, 910. 11. Sheetz, M. P., Turney, S., Qian, H., and Elson, E. L. (1989). Nature, 340, 284. 12. Sako, Y. and Kusumi, A. (1994). J. Cell Biol, 125, 1251. 13. Simson, R., Sheets, E. D., and Jacobson, K. (1995). Biophys. J., 69, 989. 14. Lee, G. M., Johnstone, B., Jacobson, K., and Caterson, B. (1993). J. Cell Biol, 123, 1899. 15. Gelles, J., Schnapp, B. J., and Sheetz, M. P. (1988). Nature, 331, 450.

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Al List of suppliers Agar Scientific Limited, 66A Cambridge Road, Stansted, Essex CM24 8DA, UK. Aldrich Chemical Co., The Old Brickyard, New Road, Gillingham, Dorset, SP8 4BR, UK. Alfa Aesar Alfa Aesar, Johnson Matthey Catalog Company, 30 Bond Street, Ward Hill, MA 01835-8099, USA. Alfa Aesar, Johnson Matthey Rare Earth Products, Waterloo Road, Widnes, Cheshire WA8 0QH, UK. Amersham Amersham International plc., Lincoln Place, Green End, Aylesbury, Buckinghamshire HP20 2TP, UK. Amersham Corporation, 2636 South Clearbrook Drive, Arlington Heights, IL 60005, USA. (Distributor for Janssen gold products.) Anderman and Co. Ltd., 145 London Road, Kingston-Upon-Thames, Surrey KT17 7NH, UK. Astrocam, Cambridge, UK. Bangs Laboratories Inc., Carmel, IN, USA. BDH (see Merck BDH) Beckman Instruments Beckman Instruments UK Ltd., Progress Road, Sands Industrial Estate, High Wycombe, Buckinghamshire HP12 4JL, UK. Beckman Instruments Inc., PO Box 3100, 2500 Harbor Boulevard, Fullerton, CA 92634, USA. Becton Dickinson Becton Dickinson and Co., 21 Between Towns Road, Cowley, Oxford OX4 3LY, UK. Becton Dickinson and Co., 2 Bridgewater Lane, Lincoln Park, NJ 07035, USA. Bellco Glass Co., 340 Edrudo Road, Vineland, New Jersey 08360-0117, USA. Bio Bio 101 Inc., do Stratech Scientific Ltd., 61-63 Dudley Street, Luton, Bedfordshire LU2 0HP, UK.

List of suppliers Bio 101 Inc., PO Box 2284, La Jolla, CA 92038-2284, USA. Bio-Rad Laboratories Bio-Rad Laboratories Ltd., Bio-Rad House, Maylands Avenue, Hemel Hempstead HP2 7TD, UK. Bio-Rad Laboratories, Division Headquarters, 3300 Regatta Boulevard, Richmond, CA 94804, USA. BioRad Microscience, Hemel Hempstead, Hertfordshire, UK. BioSoft, Cambridge, UK. Biosynth Biosynth Ag, Rietlistr. 4, Unteres Buriet, PO Box 125, Staad, SG 9422, Switzerland. Biosynth International Inc., 1665 West Quincy Avenue, Suite 155, Naperville, IL 60540, USA. Boehringer Mannheim Boehringer Mannheim UK (Diagnostics and Biochemicals) Ltd., Bell Lane, Lewes, East Sussex BN17 1LG, UK. Boehringer Mannheim Corporation, Biochemical Products, 9115 Hague Road, PO Box 504 Indianopolis, IN 46250-0414, USA. Boehringer Mannheim Biochemica, GmbH, Sandhofer Str. 116, Postfach 310120 D-6800 Ma 31, Germany. British Drug Houses (BDH) Ltd., Poole, Dorset, UK. Calbiochem Calbiochem Novabiochem (UK) Ltd., Highfield Science Park, Nottingham NG7, UK. Calbiochem Corporation, PO Box 12087, San Diego, California, USA. Citifluor Products, The Chemical Laboratory, The University of Kent, Canterbury CT2 7NH, UK. Clark Electromedical Instruments, PO Box 8, Pangbourne, Reading RG8 7HU, UK. Collaborative Biomedical Products Collaborative Biomedical Products, Two Oak Park, Bedford, MA 01730, USA. Collaborative Biomedical Products, do Becton Dickinson UK Ltd., Between Towns Road, Cowley, Oxford OX4 3LY, UK. Corel Corporation, Ottawa, Ontario, Canada. Dako Ltd., 16 Manor Courtyard, Hughenden Avenue, High Wycombe HP13 5RE, UK. Data Cell Ltd., Hattori House, Vanwall Business Park, Maidenhead, Berkshire SL6 4UB, UK. Data Translation, 100 Locke Drive, Marlboro, Massacheusetts 01752-1192, USA. Decon Laboratories Ltd., Conway Street, Hove, East Sussex BN3 3LY, UK. Difco Laboratories Difco Laboratories Ltd., PO Box 14B, Central Avenue, West Molesey, Surrey KT8 2SE, UK. 438

List of suppliers Difco Laboratories, PO Box 331058, Detroit, MI 48232-7058, USA. Ditric Optics Inc., Hudson, MA, USA. Du Pont Dupont (UK) Ltd. (Industrial Products Division), Wedgwood Way, Stevenage, Hertfordshire SG1 4Q, UK. Du Pont Co. (Biotechnology Systems Division), PO Box 80024, Wilmington, DE 19880-002, USA. Ealing Electro-Optics plc., Watford, Hertfordshire, UK. Eppendorf Eppendorf-Netheler-Hinz GmbH, 22331 Hamburg, Germany. Eppendorf, c/o Merck Ltd., Merck House, Seldown Road, Poole, Dorset

BH15 1TD, UK. European Collection of Animal Cell Culture, Division of Biologies, PHLS Centre for Applied Microbiology and Research, Porton Down, Salisbury, Wiltshire SP4 0JG, UK. E-Y Laboratories Inc., 107 North Amphlett Blvd., San Mateo, CA 94401, USA. Falcon (Falcon is a registered trademark of Becton Dickinson and Co.) Fisher Scientific Co., 711 Forbest Avenue, Pittsburgh, PA 15219-4785, USA. Flow Laboratories, Woodcock Hill, Harefield Road, Rickmansworth, Hertfordshire WD3 1PQ, UK. Fluka Fluka-ChemieAG, CH-9470, Buchs, Switzerland. Fluka Chemicals Ltd., The Old Brickyard, New Road, Gillingham, Dorset SP8 4JL, UK. Foster Findlay Associates Ltd., Newcastle Technopole, Kings Manor, Newcastle upon Tyne NE1 6PA, UK. Gibco BRL Gibco BRL (Life Technologies Ltd.), Trident House, Renfrew Road, Paisley PA3 4EF, UK. Gibco BRL (Life Technologies Inc.), 3175 Staler Road, Grand Island, NY 14072-0068, USA. Glen Spectra Ltd., Stanmore, Middlesex, England. Graticules Division (see Pyser SGI). Hamamatsu Photonics Hamamatsu Photonic Systems, 360 Foothill Road, Box 6910, Bridgewater, NJ 08807-0910, USA. Hamamatsu Photonics, Lough Point, 2 Gladbeck Way, Windmill Hill, Enfield, Middlesex EN2 7JA, UK. Arnold R. Horwell, 73 Maygrove Road, West Hampstead, London NW6 2BP, UK. Hybaid Hybaid Ltd., 111-113 Waldegrave Road, Teddington, Middlesex TW11 8LL, UK. 439

List of suppliers Hybaid, National Labnet Corporation, PO Box 841, Woodbridge, NJ 07095, USA. HyClone Laboratories, 1725 South HyClone Road, Logan, UT 84321, USA. ICN Biomedicals Inc., 1263 South Chillicothe Road, Aurora, Ohio 44202, USA. Imaging Associates Ltd., 8 Thame Park Business Centre, Wenman Road, Thame, Oxon OX9 3XA, UK. International Biotechnologies Inc., 25 Science Park, New Haven, Connecticut 06535, USA. Invitrogen Corporation Invitrogen Corporation, 1600 Faraday Avenue, Carlsbad, CA 92008, USA. Invitrogen Corporation, Invitrogen B.V., PO Box 2312, 9704 CH, Groninen, The Netherlands. Jackson Immunoresearch Laboratories Inc., 872 West Baltimore Pike, PO Box 9, West Grove, PA 19390, USA. JASC Inc., Eden Prairie, MN, USA. Kodak: Eastman Fine Chemicals, 343 State Street, Rochester, NY, USA. Leica UK Ltd., Davy Avenue, Knowlhill, Milton Keynes MK5 8LB, UK. Life Technologies Inc., 8451 Helgerman Court, Gaithersburg, MN 20877, USA. Media Cybernetics, 8484 Georgia Avenue, Silver Spring, Maryland 20910, USA. Merck Merck Industries Inc., 5 Skyline Drive, Nawthorne, NY 10532, USA. Merck, Frankfurter Strasse, 250, Postfach 4119, D-64293, Germany. Merck BDH, Merck House, Poole, Dorset BH15 1TO, UK. Micro Instruments Ltd., 18, Hanborough Business Park Lodge, Freeland, Oxford, UK. Millipore Millipore (UK) Ltd., The Boulevard, Blackmoor Lane, Watford, Hertfordshire WD1 8YW, UK. Millipore Corp./Biosearch, PO Box 255, 80 Ashby Road, Bedford, MA 01730, USA. Molecular Probes, Eugene, OR, USA. Molecular Probes Europe BV, Leiden, The Netherlands. Narishige Narishige Europe Ltd., Unit 7, Willow Business Park, Willow Way, London SE26 4QP, UK. Narishige USA Inc., 404 Glen Cove Avenue, Sea Cliff, New York 11579, USA. National Diagnostics Ltd., Unit 4, Fleet Business Park, Itlings Lane, Hessle, Hull HU13 9LX, UK. New England Biolabs (NBL) 440

List of suppliers New England Biolabs (NBL), 32 Tozer Road, Beverley, MA 01915-5510, USA. New England Biolabs (NBL), c/o CP Labs Ltd., PO Box 22, Bishops Stortford, Hertfordshire CM23 3DH, UK. Nicholson Precision Scientific, 7851 Beechcraft Avenue, Gaithersburg, MD 29879, USA. Nikon Corporation, Fuji Building, 2-3 Marunouchi 3-chome, Chiyoda-ku, Tokyo, Japan. Nyegaard AS, Postbox 4220, Torshov, Oslo 4, Norway. Omega Optical Inc., Brattleboro, Vermont, USA. Optimas Corporation UK, 40 Churchill Square, Kings Hill, East Mailing, Kent ME19 6DU, UK. Oriel, Stratford, CT, USA. Oxoid Ltd., Wade Road, Basingstoke, Hampshire RG24 0PW, UK. Ted Pella Inc., 4595 Mountain Lakes Blvd. Redding, CA 96003, USA. Perkin-Elmer Perkin-Elmer Ltd., Maxwell Road, Beaconsfield, Buckinghamshire HP9 1QA, UK. Perkin Elmer Ltd., Post Office Lane, Beaconsfield, Buckinghamshire HP9 1QA, UK. Perkin Elmer-Cetus (The Perkin-Elmer Corporation), 761 Main Avenue, Norwalk, CT 0689, USA. Pharmacia Biotech Europe, Procordia EuroCentre, Rue de la Fuse-e 62, B1130 Brussels, Belgium. Pharmacia Biosystems Pharmacia Biosystems Ltd. (Biotechnology Division), Davy Avenue, Knowlhill, Milton Keynes MK5 8PH, UK. Pharmacia LKB Biotechnology AB, Bjorngatan 30, S-75182 Uppsala, Sweden. Photometries Inc., 3440 East Britannia Drive, Tucson, AZ 85706, USA. Polysciences Inc., 400 Valley Road, Warrington, PA 18976-2590, USA. Promega Promega Ltd., Delta House, Enterprise Road, Chilworth Research Centre, Southampton, UK. Promega Corporation, 2800 Woods Hollow Road, Madison, WI 53711-5399, USA. Pyser SGI, Morley Road, Tonbridge, Kent TN1 1RN, UK. Qiagen Qiagen Inc., c/o Hybaid, 111-113 Waldegrave Road, Teddington, Middlesex TW11 8LL, UK. Qiagen Inc., 9259 Eton Avenue, Chatsworth, CA 91311, USA. Rolyn Optics, 706 Arrow Grand Circle, Covina, CA 91722-2199, USA. Schleicher and Schuell Schleicher and Schuell Inc., Keene, NH 03431A, USA. 441

List of suppliers Schleicher and Schuell Inc., D-3354 Dassel, Germany. Schleicher and Schuell Inc., do Andermann and Co. Ltd. Shandon Scientific Ltd., Chadwick Road, Astmoor, Runcorn, Cheshire WA7 1PR, UK. Sigma Chemical Company Sigma Chemical Company (UK), Fancy Road, Poole, Dorset BH17 7NH, UK. Sigma Chemical Company, 3050 Spruce Street, PO Box 14508, St. Louis, MO 63178-9916, USA. Sorvall DuPont Company, Biotechnology Division, PO Box 80022, Wilmington, DE 19880-0022, USA. Spectrum Medical Industries Inc., 1100 Rankin Road, Houston, TX 77073, USA. Stratagene Stratagem Ltd., Unit 140, Cambridge Innovation Centre, Milton Road, Cambridge CB4 4FG, UK. Stratagene Inc., 11011 North Torrey Pines Road, La Jolla, CA 92037, USA. Synoptics Ltd., 271 Cambridge Science Park, Milton Road, Cambridge CB8 4WE, UK. Taab Laboratories Equipment Ltd., 3 Minerva House, Calleva Industrial Park, Aldermaston, Berkshire RG7 4QW, UK. The Binding Site The Binding Site Ltd., PO Box 4073, Birmingham B29 6AT, UK. The Binding Site Inc., 5889 Oberlin Drive, Suite 101, San Diego, CA 92121, USA. United States Biochemical, PO Box 22400, Cleveland, OH 44122, USA. Universal Imaging Corporation, 502 Brandywine Parkway, West Chester, PA 19380, USA. Vector Laboratories Vector Laboratories Ltd., 16 Wulfric Square, Bretton, Peterborough PE3 8RF, UK. Vector Laboratories, Inc., 30 Ingold Road, Burlingame, CA 94010, USA. Wellcome Reagents, Langley Court, Beckenham, Kent BR3 3BS, UK. Wilmad Glass, US Route 40 and Oak Road, Buena, NJ 08310, USA. Yakult Honsha Co. Ltd., 1-1-19 Higashi-Shinbashi, Minato-ku, Tokyo 105, Japan. Yakutt Pharmaceutical Ind. Co. Ltd., 1-19 Higashi Shinbashi, Minato-ku, Tokyo 16, Japan. Carl Zeiss (Oberkochen) Ltd., PO Box 78, Woodfield Road, Welwyn Garden City, Hertfordshire AL7 1LU, UK. Zymed Laboratories Inc., 52 South Linden Avenue, Suite 3, South San Francisco, CA 94080, USA.

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A2 Suppliers for specialist items Adobe Systems, Inc., 345 Park Avenue, San Jose, CA 95110-2704, USA. AVT GmbH, Benzstrasse 2a, D-63741 Aschaffenburg, Germany. Barco International, Noordlaan 5, B-S520 Kuurne, Belgium. Bio-Rad Microscience Ltd., Hemel Hempstead, Hertfordshire HP2 7TD, UK. Bitplane AG, Technoparkstrasse 1, CH-8005 Zurich, Switzerland. Cargille Laboratories, Inc., 55 Commerce Road, Cedar Grove, NJ 070091289, USA. Carl Zeiss Jena GmbH, Produktbereich Mikroskopie, D-07740 Jena, Germany. Carolina Biological Supply Comp., 2700 York Road, Burlington, NC 27215, USA. Clay Adams Co., Parsippany, NJ, USA. COHU, Inc., 5755 Kearny Villa Road, San Diego, CA 92123-5623, USA. Colorado Video, Inc., Boulder, CO 80306, USA. DAGE-MTI, Inc., 701 North Roeske Avenue, Michigan City, IN 46360, USA. Data Translation, Inc., 100 Locke Drive, Marlboro, MA 01752-1192, USA. Datacube, Inc., 300 Rosewood Drive, Danvers, MA 01923, USA. Diagnostic Instruments, Inc., 6540 Burroughs Street, Sterling Height, Michigan 48314-2133, USA. Eastman Kodak Company, 343 State Street, Rochester, NY 14650-2010, USA. EOS Electronics AV Ltd., Barry, South Glamorgan, UK. FAST Electronic GmbH, Landsberger Strasse, 76, D-80007 Munich, Germany. For-A Company Ltd., Tokyo 160, Japan. Hamamatsu Photonics Deutschland GmbH, Arzbergerstrasse 10, D-82211 Hersching, Germany. HaSoTec GmbH, Burgwall 20, D-18055 Rostock, Germany. Imaging Technology, Inc., 55 Middlesex Turnpike, Bedford, MA 01730, USA. Improvision, Inc., Viscount Centre II, University of Warwick Science Park, Coventry CV4 7HS, UK. Inovision Corp., 6321 Angus Drive, Raleigh, NC 27613, USA. Jasc Software, Inc., 11011 Smetana Road, Minnetonka, MN 55343, USA.

Suppliers for specialist items Javelin Systems, 23456 Hawthrone Blvd, Torrance, CA 90505-4716, USA. Kappa Messtechnik GmbH, Kleines Feld 6, D-37130 Gleichen, Germany. Leica Microsystems Holding GmbH, Ernst-Leitz-Strasse, D-35578 Wetzlar, Germany. Matrox Electronic Systems (UK) Ltd., 6 Cherry Orchard West, Kembrey Park, Swindon, Wiltshire SN2 6UP, UK. Media Cybernetics, 8484 Georgia Avenue, Silver Spring, MD 20910, USA. Mitec GmbH, Prof. Messerschmidt Strasse 3, D-85579 Neubiberg, Germany. Mitsubishi Corp., 5665 Plaza Drive, Cypress, CA 90630, USA. Modulation Optics, Inc., Greenville, NY 11548, USA. Molecular Probes, Inc., 4849 Pichforde Avenue, Eugene, OR 94702-0469, USA. Motion Analysis, 3617 Westwind Blvd, Santa Rosa, CA 95403, USA. MS MacroSystem Computer GmbH, Borgacker 2-6, D-58454 Witten, Germany. NIH, Research Services Branch, Bethesda, MD, USA. Nikon Europe BV, Schipholweg 321, 1170 AE Badhoevedorp, The Netherlands. O. Kindler GmbH & Co., Ziegelhofstrasse 214, D-79110 Freiburg, Germany. Olympus, Inc., Two Corporate Center Drive, Melville, NY 11747-3157, USA. Optronics, 175 Taft Court, Suite 111, Goleta, CA 93117, USA. PCO Computer Optics GmbH, Ludwigsplatz 4, D-93309 Kelheim, Germany. Photometries Ltd., 3440 East Britannia Drive, Tucson, AZ 35706, USA. PTI (Photon Technology International), Inc., Deer Park Drive, Suite F, Monmouth Junction, NJ 08852, USA. Photonic Science, Millham, Mountfield, Robertsbridge, East Sussex TN32 5LA, UK. Polaroid Export Europe, Wheathampstead House, Codicote Road, Wheathampstead Hertfordshire AL4 8SF, UK. (CCD cameras, slide makers) Polaroid, 575 Technology Square, 9P, Cambridge, MA 02139, USA. (Polarizers) Princeton Instruments, Inc., 3660 Quakersbridge Road, Trenton, NJ 08619, USA. Proxitronic GmbH, Robert-Bosch-Strasse 34, D-64625 Bensheim, Germany. Quantel Ltd., Turnpike Road, Newbury, Berkshire RG14 2NE, UK. Reichert/Cambridge Instruments, see Leica Scanalytics, Inc., 8550 Lee Highway, Fairfax, VA 22031-1515, USA. Scion Corp., 82 Worman's Mill Court, Suite H, Frederick, MD 21703, USA. Sharp, Inc., 22-22 Nagike-cho Abeno-ku, Osaka 545, Japan. Sigma Chemical Company, 3050 Spruce Street, St Louis Missouri, USA. Sony Corp., 7-35 Kitashinaggawa 6-chome, Shinagawa, Tokyo 141, Japan. Stemmer Imaging GmbH, Gutenbergstrasse 11, D-82178 Puchheim, Germany. Tektronix, Inc., 26600 SW Parkway, Wilsonville, OR 97070, USA. Theta Systeme Dr. Tatarczyk GmbH, John F. Kennedy Strasse 9, D-82194 Grobenzell, Germany. 444

Suppliers for specialist items Uniblitz, SD10 Vincent Assoc., 1255 University Avenue, Rochester, NY 14607, USA. Universal Imaging Corp., 502 Brandywine Parkway, West Chester, PA 19380, USA. VayTek, Inc., 305 E Lowe Avenue, PO Box 732, Fairfield, IA 52556, USA. Vital Images, 505 N Fourth Street, Fairfield, Iowa 52556, USA.

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Index aberrations chromatic 53-4 and point spread function 51-4 and refractive index of specimen 53 spherical 51-3 Airy disc 13, 14, 50-1, 107, 108 Allen video-enhanced contrast (AVEC) 75, 84, 104-5, 107-8 applications 117-23 interpretation 116-17 limitations 117-22 setting up microscope 110-16 anaxial illumination, video microscopy 110, 118-19 antibodies conjugation to colloidal gold 429-30 monoclonal 187-8 polyclonal antisera 187 purification 188 specificity 188-9 storage 189 structure 186-7 atomic force microscopy 342 cellular substructures 362-3 chromosomes 363, 365-6 autofluorescence, induction by glutaraldehyde 204 back focal plane, objective 8, 50 Beer-Lambert law, and absorption measurements 132 B-glucuronidase, histochemical method 381-2 biolistic process, see particle bombardment calcium ions properties of specific dyes 223-7, 230-1 quantification in cells 129, 133, 221-74 ratio imaging 129, 133, 230-1 second messenger in cells 221-2 see also ion imaging cameras CCD 89-91, 93-4, 126-7, 433 film 30-3, 180-1 SIT 126 video 33-8, 89-97 video-intensified microscopy (VIM) 91-5, 123-7 carbohydrates, identification using lectins 185 C-banding of chromosomes, method 162-3 CCD cameras chromosome analysis 182

ion imaging 242 for nanovid microscopy 433 video microscopy 89-91, 93-4, 126-7 cell culture for chromosome studies 152-5, 157 mammalian cells 152-5 for nanovid microscopy 430-1 cell proliferation, and nucleolus organizing regions (NORs) 338 cell-substrate contacts, visualization 420, 422 chromosome banding AgNOR-banding method 164, 166-7 C-banding method 162-3 classification 161-2 definition 161 G-banding method 164 kinetochore staining 166-8 observation 160, 178 chromosome preparation cytocentrifugation 155-6 for immunocytochemistry 155-6 mammalian cells 152-5 plant cells 156-9 quality assessment 159-60 chromosomes atomic force microscopy 363, 365-6 fixation 154-9, 167 fluorescence microscopy 151, 158, 160-1, 178-81 near-field scanning optical microscopy (NSOM) 363-7 observation methods 160, 177-83 photomicrography 180-1 preparation methods 151-60 shear force microscopy 366-8 uniform (solid) staining 160-1 colloidal gold conjugation of antibodies 429-30 label for immunohistochemistry 197, 209-10 nanovid microscopy 425-35 reflection-contrast microscopy 280 silver enhancement 209-10 condensers dark-field illumination 21 light microscope 2-4, 7, 10 confocal microscopy 45-71 choice of objectives 64—5 chromosome analysis 181-2 comparison with other techniques 58-61, 182,422 ion imaging 243 noise 58-60, 61 optical sectioning 62-3

Index confocal microscopy (contd.) out-of-focus light 60-2 photobleaching 242 point scanning 54-5 and point spread function 48-51, 246-8 principle 47-8 setting up 68-70 slit scanning 55-6 specimen preparation 63-8 spinning (Nipkow) disc 56-7 two-photon imaging 57-8 ultraviolet 54 use of 62-70 conjugate planes, light microscope 7-10 contrast methods bright-field 16-17, 40-1 dark-field 17-23, 40-1, 117-18 differential interference contrast (DIC) 25-6, 40, 77 fluorescence microscopy 26, 41, 45 Hoffman modulation contrast 110, 118 nanovid microscopy 425-35 phase-contrast 22-3, 40 polarized light 23-8, 40 Rheinberg illumination 17 test specimen 38-41 video-enhanced contrast (VEC) 73, 75, 77-8, 106-8 video-intensified microscopy 29, 40, 75-8, 123-7 coverslips sealing 109-10 thickness 2, 108-9, 246 culture chambers, for living specimens 248 cytocentrifugation, chromosome preparation 155-6 dark ground contrast, see contrast methods, dark-field diatoms, test specimens for resolution 10-12 differential interference contrast (DIC) 25-6, 40,77 observation of chromosomes 160, 178 and video microscopy 107, 110-11, 113, 115-16, 120-2 dry mass measurement, interference microscopy 39 electron microscopy, and reflection-contrast microscopy 298 electrophysiology, and ion imaging 270 electroporation, introduction of dyes into cells 235-6 embedding paraffin for light microscopy 313-14 resin 301-5

evanescent illumination, see total internal reflection fluorescence (TIRF) extracellular matrix, study by nanovid microscopy 428 eyepieces light microscope 2, 6-9 Ramsden disc 6, 9 fading of fluorochromes, see photobleaching field of view, light microscope 2—3 filters

fluorescence microscopy 179, 239-40 light microscopy 16-18, 32-3, 178, 180 fixation chromosomes 154-9, 167 glutaraldehyde 64, 301-4 for histomorphometry 313-14 for immunohistochemistry 167, 189-91, 300-4 paraformaldehyde 65-6 fixatives, formulae 190-1 fluorescence measurement using video microscopy 132-3 standards 246-8 video microscopy 121-2 fluorescence lifetime imaging 422 fluorescence microscopy 26, 41, 45 antifade mountants 65-6, 175-6, 179-80 applications 45-6 choice of objectives 64-5, 178 chromosomes 151, 158, 160-1, 178-81 comparison with confocal microscopy 58-61 epi-illumination 45,178-9 filters 179, 239-40 illumination 178-9, 239 of ions in cells 237-45 photobleaching 57, 69, 179-80, 223, 226-9, 242 setting up 245-8 signal-to-noise ratio 254-6 fluorescence recovery after photobleaching (FRAP) 129 fluorescence in situ hybridization (FISH), see in situ hybridization fluorescent analogue cytochemistry 127-9 fluorochromes

calcium-sensitive 222-7, 230-1 for chromosome studies 179 excitation and emission wavelengths 179 photobleaching 57, 69, 179-80, 226-9 pH-sensitive 222-5, 228-31 G-banding of chromosomes, method 164 glutaraldehyde fixation 64, 301-4 induction of autofluorescence 204 green fluorescent protein 46, 380, 385-6, 395 448

Index haematoxylin and eosin, staining method 314-15 histomorphometry 131-2, 311-40 anisotropy 326 colour imaging 334 image analysis 327-38 nuclear volume measurement 321-3, 329, 336 shape measurement 337-8 specimen preparation 313-18 statistical analysis 326-7 volume fraction measurement 323-6, 337 Hoffman modulation contrast 110, 118 horseradish peroxidase label for immunohistochemistry 196-7, 204-7, 214, 317-18 silver enhancement 205-6 illumination colour temperature 32 fluorescence microscopy 178-9, 239 ion imaging 239 K6hler 7-10, 31 light microscope 1-4,7-10 reflection-contrast microscopy 282-3 two-photon microscopy 57-8, 244 video microscopy 101-2,112-13 image formation light microscope 3-6 reflection-contrast microscopy 286-90 image processing analogue 75, 95-7 chromosome analysis 183 dangers 183 digital 75, 97-101 histomorphometry 327-38 ion imaging 249, 267 for nanovid microscopy 433-4 video microscopy 78-89, 95-101 image recording drawing 29-30 photomicrography 30-3, 38, 180-1 video cameras 33-7, 38, 73-149 see also photomicrography immersion oil, choice of 109 immunocytochemistry, see immunohistochemistry immunoglobulins, see antibodies immunohistochemistry 185—220 adhesion of sections to slides 210-12 of cell smears 215—18 chromosome preparation 155-6 controls 213 double labelling 214-15 fixation for 167, 189-91 kinetochores 166-8 labelling methods 202-13, 218-19 labels 194-201, 204-10, 214, 317-18, 429-30 mountants 204, 212-13

problem solving 213-14 quantification 218 reflection-contrast microscopy 275-6, 280, 288, 290-1, 295, 305-8 resin sections 305-7 specimen preparation 189-94, 298-301 unmasking hidden antigens 192-4 Inoue' system, video-enhanced contrast (VEC) 107-8 in situ hybridization applications to chromosomes 168-70 labels 171-3 method 173-7 near-field scanning optical microscopy (NSOM) 365-8 probe preparation 170-3 reflection-contrast microscopy 275-6, 295-6 resolution 170 interference microscopy dry mass measurement 39 measurement of optical path differences 29 see also differential interference contrast (DIC) internal reflection microscopy comparison with total internal reflection fluorescence (TIRF) 422 visualization of cell-substrate contacts 422 introduction of materials into living cells, see electroporation; ion-sensitive dyes; microinjection; particle bombardment ion imaging calcium measurement in cells 221-74 calibration 253, 260-4 combination with other techniques 269-70 confocal microscopy 243 controls 270-1 and electrophysiology 270 equipment for 240-5 filters 239-40 fluorochromes for 222-31 image processing 249, 267 light sources 239 multiphoton imaging 244-5 multiple detectors 245 pH measurement in cells 221-74 quantification 252-67 ratio imaging 249-52 ratiometric dyes 230-1 signal-to-noise ratio 254-6 statistical analysis 264-7 ion-sensitive dyes introduction into cells 231-7 photobleaching 223, 226-7, 242, 263 kinetochores, immunocytochemical labelling 166-8 Kohler illumination 7-10, 31

449

Index lectins, identification of carbohydrates 185 light, interactions with matter 5-6 light microscope condensers 2-4, 7, 10 conjugate planes 7-10 eyepieces 2, 6-9 field of view 2-3 illumination 1-4, 7-10, 31 image formation 3-6 linear measurement 319-20 magnification changer 3, 105, 106 objectives 2-6, 9-14, 64-5, 102-4 resolution 6, 10-14, 45 setting up 7-10, 318-19 light sources, see illumination linear measurement, light microscope 319-20 living specimens culture chambers 248 effects of irradiation 248-9 observation 109, 110, 248 reflection-contrast microscopy 275, 289, 308 specimen preparation 108-9 temperature control 249 luminescence 129 magnification, empty 10-11, 14-15 magnification changer, light microscope 3, 105, 106 mammalian cells chromosome preparation 152-5 culture 152-5 membrane mobility, study by nanovid microscopy 428, 434-5 microinjection advantages 386-7 equipment 387-91 identification of cells 391-2, 394-5 introduction of materials into living cells 386-95 technique 392-4 and total internal reflection fluorescence (TIRF)410, 412 microspectrofluorimetry 129 molecular imaging 129 monolayers, near-field scanning optical microscopy (NSOM) 358-62 mountants antifading for fluorescence microscopy 65-6, 175-6, 179-80 for immunohistochemistry 204, 212-13 reflection-contrast microscopy 307 movement, detection by video microscopy 87-8, 101, 122-3, 131-2 nanovid microscopy 425-35 analysis of mobility in membranes 428, 434-5

camera 433 cell culture 430-1 image processing 433-4 image storage 434 microscope 432-3 observation of colloidal gold particles 425-35 principle 425-7 study of extracellular matrix 428 viewing cells 431-2 near-field scanning optical microscopy (NSOM) 341-71 applications 344, 355-67 chromosomes 363-7 distance regulation 349-50 in situ hybridization 365-8 instrumentation 344-9 monolayers 358-62 operation 353-4 principle 342-3 probe design 344-9 probe fabrication 351-3 resolution 346, 355, 359, 362-3, 367, 369 and shear force microscopy 349-50, 353-4 single molecule detection 355-8 subcellular structures 362-3 viruses 362 neurobiology, applications of video microscopy 130 neurons, visualization by video-enhanced contrast 130 Nomarski interference, see differential interference contrast (DIC) nucleolus organizing regions (NORs) measure of cell proliferation 338 silver staining 164, 166, 315-16 numerical aperture and light-gathering power 102-3 objectives 2, 6, 10-14, 50, 102-3 and resolution 6, 10-14, 45

objectives back focal plane 8, 50 cleaning 114, 246 for confocal microscopy 64—5 coverslip thickness 246 for fluorescence 64-5, 178 immersion 2, 53, 64—5 light-gathering power 102-3 light microscope 2, 6, 9-14, 64-5, 102-4 light transmission 103 numerical aperture 2, 6, 10-14, 50, 102-3 reflection-contrast microscopy 277-81, 284 resolution 6, 10-14 water immersion 109, 110 optical microscope, see light microscope

450

Index out-of-focus light removal by confocal microscopy 46, 60-2 removal by deconvolution 46, 61 paraformaldehyde, fixation 65-6 particle bombardment apparatus 374-5 applications 373 identification of transformed cells 380-6 introduction of materials into living cells 373-86 particle preparation 375-7 target tissue 377-80 PH properties of specific dyes 223-5, 228-31 second messenger in cells 221-2 phase-contrast 22-3 observation of chromosomes 160, 178 video microscopy 119 pH measurement in cells 133, 221-74 ratio imaging 133, 230-1 see also ion imaging photobleaching, fluorochromes 57, 69, 179-80, 223, 226-9, 242 photography, video microscopy 142-3 photomicrography 30-3, 38 black and white 32-3, 180 chromosomes 180-1 colour 32-3, 180-1 photon scanning tunnelling microscopy (PSTM) 345, 349 plant cells, chromosome preparation 156-9 point spread function and aberrations 51-4 and confocal imaging 48-51, 246-8 and resolution 246-8 polarized light contrast methods 23-8, 40 video-enhanced contrast (VEC) 113-14, 119-20,122 quantification absorption 132 fluorescence 132-3, 237, 252-67 immunohistochemistry 218 ions in cells 132-3, 230-1, 237, 252-67 Ramsden disc, eyepieces 6, 9 ratio imaging calcium measurement in cells 129-33, 230-1 ions in cells 129, 133, 230-1, 249-52, 257-60 pH measurement in cells 133, 230-1 problems 258-60 quantification 257-67

Rayleigh criterion, resolution 13-14, 82 reflection-contrast microscopy 121, 275-310 applications 275-6, 290-8 colloidal gold 280 detection of rare events 296-7 and electron microscopy 298 focusing 284-6 image formation 286-90 immunohistochemistry 275-6, 280, 288, 290-1, 295, 305-8 in situ hybridization 275-6, 295-6 light sources 282—3 living cells 275, 289, 308 mounting medium 307 objectives 277-81, 284 optical systems 276-86 specimen preparation 290-1, 298-305 unstained material 294 refractive index, measurement 39 resin sections immunohistochemistry 305-7 specimen preparation 301-5 resolution diatoms as test specimens 10-12 in situ hybridization 170 ion imaging 252—3 near-field scanning optical microscopy (NSOM) 346, 355, 359, 362-3, 367, 369 and numerical aperture 6, 10-14 objectives 6, 10-14,45 photographic film 86 photon scanning tunnelling microscopy (PSTM) 349 point spread function 246-8 Rayleigh criterion 13-14, 82 Sparrow criterion 14, 82 star test 13-14 video microscopy 75-7, 82, 86 Rheinberg illumination 17 RNA transcription, in vitro 67-8, 70 scanning probe microscopy 341-2 second messengers 221-2 shear force microscopy cellular substructures 363-4 chromosomes 366-8 and near-field scanning optical microscopy (NSOM) 349-50, 353-4 signal-to-noise ratio, fluorescence microscopy 254-6 silver enhancement colloidal gold 209-10 horseradish peroxidase 205-6 silver staining, nucleolus organizing regions (NORs) 164, 166, 315-16, 338 single molecule detection, near-field scanning optical microscopy (NSOM) 355-8

451

Index slides, adhesion of specimens 66, 210-12, 248, 307-8 specimen preparation chromosomes 151-60 for confocal microscopy 63-8 histomorphometry 313-18 immunohistochemistry 189-94, 298-301 living specimens 108-9 reflection-contrast microscopy 290-1, 298-305 resin sections 301-5 for video microscopy 108-10 specimens, adhesion to slides 66, 210-12, 248, 307-8 staining chromosomes 160-7 haematoxylin and eosin 314-15 star test, resolution 13-14 statistical analysis histomorphometry 326-7 ion imaging 264-7 stereology definition 312 symbols 313 see also histomorphometry stray light 79-80, 84, 107-8, 119, 121 surface fluorescent microscopy, see total internal reflection fluorescence (TIRF) surface phenomena, microscopical detection 399-423

time lapse microscopy 137-8, 420 total internal reflection fluorescence (TIRF) applications 399, 420-2 combination with other microscopical techniques 401, 417-18 comparison with internal reflection microscopy 422 experimental tips 417-20 and microinjection 410, 412 microscopical configurations 407-17 principles 399-401 theory 401-7 transformed cells, identification 380-6 two-photon microscopy 57-8, 242, 244-5 illumination 57-8, 244 ion imaging 244-5 photobleaching 242

ultraviolet microscopy, confocal 54 video cameras image recording 33-8 for nanovid microscopy 433 video-enhanced contrast (VEC) 73, 75-84, 106-23 illumination 101-2 Inoue system 107-8 polarized light 113-14, 119-20, 122 setting up the microscope 110-14 visualization of neurons 130 see also nanovid microscopy video-intensified microscopy (VIM) 29, 40, 73, 75-8, 123-7 applications 127-30 cameras 91-5, 123-7 image acquisition 124-7 microscope requirements 123-4 video microscopy 73-149 anaxial illumination 118-19 applications 117-23 cameras 89-97 chromosome analysis 182 colour images 127 dark-field 117-18 detection of movement 87-8, 122-3, 131-2 and differential interference contrast 107, 110-11, 113, 115-16,120-2 digital image processing 76, 84-9, 97-9 editing video tapes 143-6 fluorescence 121-2 fluorescence measurements 132-3 Hoffman modulation contrast 118 illumination 101-2, 112-13 image intensification 29, 40, 77-9 image recording 133-46 light sources 101-2 microscope requirements 101-6 phase-contrast 119 printouts for presentation 141-2, 143 recording equipment 133-7 reflection-contrast 121, 275-310 resolution 75-7, 82 size measurements 131 storage media 140 time lapse recording 137-8 viruses, near-field scanning optical microscopy (NSOM) 362

452